Comparative Physiology and Evolution of the Autonomic Nervous System [1 ed.] 3718651378, 9783718651375

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Co m p a r a t i v e P h y s i o l o g y a n d E v o l u t i o n of th e A u t o n o m i c Ne r v o u s S y s t e m

The Autonomic Nervous System

A series o f books discussing all aspects o f the autonomic nervous system. Edited by G. Burnstock, Department o f Anatomy and Developmental Biology, University College London, UK.

Volume 1

Autonomic Neuroeffector Mechanisms edited by G. Burnstock and C. H. V. Hoyle Volume 2

Development, Regeneration and Plasticity of the Autonomic Nervous System edited by L A . Hendry and C. E. Hill Volume 3

Nervous Control of the Urogenital System edited by C. A. Maggi

Volume 4

Comparative Physiology and Evolution of the Autonomic Nervous System edited by S. Nilsson and S. Holmgren Volume 5

Disorders of the Autonomic Nervous System edited by D. Robertson and L Biaggioni

Volume 6

Autonomic Ganglia edited by E. McLachlan Other volumes in preparation Autonomic Control of the Respiratory System P. Barnes

Central Nervous Control of Autonomic Function D. Jordan

Nervous Control of Blood Vessels T. Bennett and S. Gardiner

Autonomic-Endocrine Interactions K. Unsicker

Nervous Control of the Heart J. T. Shepherd and S. Vatner

Nervous Control of the Eye A. Sillito and G. Burnstock

Autonomic Innervation of the Skin I. Gibbins and J. Morris

Nervous Control of the Gut and Associated Organs M. Costa

Co m p a r a t i v e P h y s i o l o g y a n d E v o l u t i o n of th e A u t o n o m i c Ne r v o u s S y s t e m

Edited by Stefan Nilsson and Susanne Holmgren Department of Zoophysiology, Zoological Institute, University of Goteborg, Sweden

First published 1994 by Harwood Academic Publishers GmbH. Published 2021 by Routledge 2 Park Square, Milton Park, Abingdon, Oxon OX14 4RN 605 Third Avenue, New York, NY 10017

Routledge is an imprint of the Taylor & Francis Group, an informa business Copyright © 1994 by Taylor & Francis. All rights reserved. No part of this book may be reprinted or reproduced or utilised in any form or by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying and recording, or in any information storage or retrieval system, without permission in writing from the publishers. Notice: Product or corporate names may be trademarks or registered trademarks and are used only for identification and explanation without intent to infringe. Library of Congress Cataloging-in-Publication Data Comparative physiology and evolution of the autonomic nervous system I edited by Stefan Nilsson and Susanne Holmgren. p. cm. -- (The Autonomic nervous system, ISSN 1047-5125 : v. 4) Includes bibliographical references and index. ISBN 3-7186-5137-8 1. Autonomic nervous system--Physiology. 2. Autonomic nervous system--Evolution. 3. Physiology, Comparative. I. Nilsson, Stefan, 1946II. Holmgren, Susanne, 1946111. Series: Autonomic nervous system (Chur, Switzerland): v. 4. [DNLM: 1. Autonomic Nervous System--physiology. 2. Physiology, Comparative. 3. Evolution. WL 600 C737 1993] QP368.C65 1993 612.8'9--dc20 DNLM/DLC for Library of Congress 93-25356 CIP

ISBN 13: 978-3-7186-5137-5 (hbk)

Geoffrey Burnstock — Editor of The Autonomic Nervous System Book Series

This volume is dedicated to Professor John Z. Young and Professor Geoffrey Burnstock, pioneers in the comparative physiology of the autonomic nervous system

Contents Series Preface — Historical and Conceptual Perspective of the Autonomic Nervous System Book Series

ix

Preface

xiii

List of Contributors

xvii

1 Comparative Anatomy and Evolution of the Autonomic Nervous System Ian Gibbins 2

Comparative Aspects on the Biochemical Identity of Neurotransmitters of Autonomic Neurons Susanne Holmgren and Jörgen Jensen

3 Chromaffin Systems Robert M. Sanier

1

69 97

4 The Gastrointestinal Canal Jörgen Jensen and Susanne Holmgren

119

5 Glands Ann-Cathrine Jönsson

169

6 The Circulatory System Judy L. Morris and Stefan Nilsson

193

7 The Spleen Stefan Nilsson

247

8 Lungs and Swimbladders Graeme Campbell and John R. McLean

257

9 Urinogenital Organs Kazumasa Uematsu

311

10 Chromatophor es David J. Grove 11

331

The Iris Stefan Nilsson

353

Index

363

Biosystematic Index

373 VÜ

Preface to the Series - Historical and Conceptual Perspective of the Autonomic Nervous System Book Series The pioneering studies of Gaskell (1886), Bayliss and Starling (1899), and Langley and Anderson (see Langley, 1921) formed the basis of the earlier and, to a large extent, current concepts of the structure and function of the autonomic nervous system; the major division of the autonomic nervous system into sympathetic, parasympathetic and enteric subdivisions still holds. The pharmacology of autonomic neuroeffector transmission was dominated by the brilliant studies of Elliott (1905), Loewi (1921), von Euler and Gaddum (1931) and Dale (1935), and for over 50 years the idea of antagonistic parasympathetic cholinergic and sympathetic adrenergic control of most organs in visceral and cardiovascular systems formed the working basis of all studies. However, major advances have been made since the early 1960s that make it necessary to revise our thinking about the mechanisms of autonomic transmission, and that have significant implications for our understanding of diseases involving the autonomic nervous system and their treatment. These advances include: (1) Recognition that the autonomic neuromuscular junction is not a ‘synapse’ in the usual sense of the term where there is a fixed junction with both pre- and postjunctional specialization, but rather that transmitter is released from mobile varicosities in extensive terminal branching fibres at variable distances from effector cells or bundles of smooth muscle cells which are in electrical contact with each other and which have a diffuse distribution of receptors (Hillarp, 1959; Burnstock, 1986a). (2) The discovery of non-adrenergic, non-cholinergic nerves and the later recognition of a multiplicity of neurotransmitter substances in autonomic nerves, including monoamines, purines, amino acids, a variety of different peptides and nitric oxide (Burnstock et al., 1964; Burnstock, 1986b, 1993b; Burnstock and Milner, 1992; Rand, 1992; Snyder, 1992). (3) The concept of neuromodulation, where locally released agents can alter neurotransmission either by prejunctional modulation of the amount of transmitter ix

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COMPARATIVE PHYSIOLOGY AND EVOLUTION

(4)

(5)

(6)

(7)

(8)

(9)

released or by postjunctional modulation of the time course or intensity of action of the transmitter (Marrazzi, 1939; Brown and Gillespie, 1957; Vizi, 1979; Kaczmarek and Leviton, 1987). The concept of cotransmission that proposes that most, if not all, nerves release more than one transmitter (Burnstock, 1976; Hokfelt, Fuxe and Pernow, 1986; Burnstock, 1990a) and the important follow-up of this concept, termed ‘chemical coding’, in which the combinations of neurotransmitters contained in individual neurones are established, and whose projections and central connections are identified (Furness and Costa, 1987). Recognition of the importance of ‘sensory-motor’ nerve regulation of activity in many organs, including gut, lungs, heart and ganglia, as well as in many blood vessels (Maggi and Meli, 1988; Burnstock, 1990a, 1993a), although the concept of antidromic impulses in sensory nerve collaterals forming part of ‘axon reflex’ vasodilation of skin vessels was described many years ago (Lewis, 1927). Recognition that many intrinsic ganglia (e.g. those in the heart, airways and bladder) contain integrative circuits that are capable of sustaining and modulating sophisticated local activities (Burnstock et al., 1987). Although the ability of the enteric nervous system to sustain local reflex activity independent of the central nervous system has been recognized for many years (Kosterlitz, 1968), it had been generally assumed that the intrinsic ganglia in peripheral organs consisted of parasympathetic neurones that provided simple nicotinic relay stations. The major subclasses of receptors to acetylcholine and noradrenaline have been recognized for many years (Dale, 1914; Ahlquist, 1948), but in recent years it has become evident that there is an astonishing variety of receptor subtypes for autonomic transmitters (see Br. J. Pharmacol., [1991], 102, 560-561). Their molecular properties and transduction mechanisms are being characterized. These advances offer the possibility for more selective drug therapy. Recognition of the plasticity of the autonomic nervous system, not only in the changes that occur during development and aging, but also in the changes in expression of transmitter and receptors that occur in fully mature adults under the influence of hormones and growth factors following trauma and surgery, and in a variety of disease situations (Burnstock, 1990b). Advances in the understanding of ‘vasomotor’ centres in the central nervous system. For example, the traditional concept of control being exerted by discrete centres such as the vasomotor centre (Bayliss, 1923) has been supplanted by the belief that control involves the action of longitudinally arranged parallel pathways involving the forebrain, brain stem and spinal cord (Loewy and Spyer, 1990).

In addition to these major new concepts concerning autonomic function, the discovery by Furchgott that substances released from endothelial cells play an important role, in addition to autonomic nerves, in local control of blood flow, has also made a significant impact on our analysis and understanding of cardiovascular function (Furchgott and Zawadski, 1980; Ralevic and Burnstock, 1993). The later identification of nitric oxide as the major endothelium-derived relaxing factor (Palmer et al., 1988) (confirming the independent suggestions by Ignarro and by Furchgott) and endothelin as an endothelium-derived constricting factor

PREFACE TO THE SERIES xi

(Yanagisawa et al., 1988) have also had a major impact in this area. In broad terms these new concepts shift the earlier emphasis on central control mechanisms towards greater consideration of the sophisticated local peripheral control mechanisms. Although these new concepts should have a profound influence on our considerations of the autonomic control of cardiovascular, urogenital, gastrointestinal and reproductive systems and other organs like the skin and eye in both normal and disease situations, few of the current textbooks take them into account. This is largely because revision of our understanding of all these different specialist areas in one volume by one author is a near impossibility. Thus, this Book Series of 14 volumes is designed to try to overcome this dilemma by dealing in depth with each major area in separate volumes and by calling upon the knowledge and expertise of leading figures in the field. Volume 1, published early 1992, dealt with the basic mechanisms of Autonomic Neuroeffector Mechanisms which set the stage for later volumes devoted to autonomic nervous control of particular organ systems, including Heart, Blood Vessels, Respiratory System, Eye, Skin, Gastrointestinal Tract and Urogenital Organs (volume 3, published in early 1993). Another group of volumes will deal with Central Nervous Control o f Autonomic Function, Autonomic Ganglia, Autonomic-Endocrine Interactions, Development, Regeneration and Plasticity o f the Autonomic Nervous System (volume 2, published in 1992) and this volume, Comparative Physiology and Evolution o f the Autonomic Nervous System. Abnormal as well as normal mechanisms will be covered to a variable extent in all these volumes, depending on the topic and the particular wishes of the volume editor, but one volume will be specifically devoted to Disorders o f the Autonomic Nervous System. A general philosophy followed in the design of this Book Series has been to encourage individual expression by Volume Editors and Chapter Contributors in the presentation of the separate topics within the general framework of the series. This was demanded by the different ways that the various fields have developed historically and the differing styles of the individuals who have made the most impact in each area. Hopefully, this deliberate lack of uniformity will add to, rather than detract from, the appeal of these books. G. Burnstock Series Editor REFERENCES Ahlquist, R.P. (1948). A study of the adrenotropic receptors. Am. J. Physiol., 153, 586-600. Bayhss, W.B. (1923). The Vasomotor System, London: Longman. Bayliss, W.M. and Starling, E.H. (1899). The movements and innervation of the small intestine. J. Physiol. (Lond.), 24, 99-143. Brown, G.L. and Gillespie, J.S. (1957). The output of sympathetic transmitter from the spleen of a cat. J. Physiol. (Lond.), 138, 81-102. Burnstock, G. (1976). Do some nerve cells release more than one transmitter? Neuroscience, 1, 239-248. Burnstock, G. (1986a). Autonomic neuromuscular junctions: current developments and future directions. J. A nat., 146, 1-30. Burnstock, G. (1986b). The non-adrenergic non-cholinergic nervous system. Arch. Int. Pharmacodyn. Ther., 280, Suppl., 1-15. Burnstock, G. (1990a). Co-transmission. The Fifth Heymans Lecture - Ghent, February 17, 1990. Arch. Int. Pharmacodyn. Ther., 304, 7-33. Burnstock, G. (1990b). Changes in expression of autonomic nerves in aging and disease. J. Auton. Nerv. Syst., 30, 525-534.

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Burnstock, G. (1993a). Introduction: Changing face of autonomic and sensory nerves in the circulation. In: Vascular Innervation and Receptor Mechanisms: New Perspectives, edited by L. Edvinsson and R. Uddman. pp. 1-22. Academic Press Inc, San Diego, USA. Burnstock, G. (1993b). Physiological and pathological roles of purines: an update. Drug Dev. Res. 28, 195-206. Burnstock, G., Campbell, G., Bennett, M. and Holman, M.E. (1964). Innervation of the guinea-pig taenia coli: Are there intrinsic inhibitory nerves which are distinct from sympathetic nerves? Int. J. Neuropharmacol., 3, 163-166. Burnstock, G., Allen, T.G.J., Hassall, C.J.S. and Pittam, B.S. (1987). Properties of intramural neurones cultured from the heart and bladder. In Histochemistry and Cell Biology o f Autonomic Neurons and Paraganglia. Exp. Brain Res. Ser. 16, edited by C. Heym, pp. 323-328. Heidelberg: Springer Verlag. Burnstock, G. and Milner, P. (1992). Structural and chemical organisation of the autonomic nervous system with special reference to nonadrenergic, noncholinergic transmission. In Autonomic Failure, 3rd edn, edited by R. Bannister, pp. 107-125. Oxford: Oxford University Press. Dale, H. (1914). The action of certain esters and ethers of choline and their reaction to muscarine. J. Pharmacol. Exp. Ther., 6, 147-190. Dale, H. (1935). Pharmacology and nerve endings. Proc. Roy. Soc. Med., 28, 319-332. Elliott, T.R. (1905). The action of adrenalin. J. Physiol. (Lond.), 32, 401-467. Furehgott, R.F. and Zawadski, J.V. (1980). The obligatory role of endothelial cells in the relaxation of arterial smooth muscle by acetylcholine. Nature, 288, 373-376. Furness, J.B. and Costa, M. (1987). The Enteric Nervous System, Edinburgh: Churchill Livingstone. Gaskeil, W.H. (1886). On the structure, distribution and function of the nerves which innervate the visceral and vascular systems. J. Physiol. (Lond.), 7, 1-80. Hillarp, N .-Ä . (1959). The construction and functional organisation of the autonomic innervation apparatus. Acta Physiol. Scand., (Suppl. 157), 46, 1-38. Hökfelt, T., Fuxe, K. and Pernow, B. (Eds). (1986). Coexistence of neuronal messengers: a new principle in chemical transmission. In Progress in Brain Research, Vol. 68, Amsterdam: Elsevier. Kaczmarek, L.K. and Leviton, I.B. (1987). Neuromodulation. The Biochemical Control of Neuronal Excitability, pp. 1-286. Oxford: Oxford University Press. Kosterlitz, H.W. (1968). The alimentary canal. In Handbook o f Physiology, vol. IV, edited by C.F. Code, pp. 2147-2172. Washington DC: American Physiological Society. Langley, J.N. (1921). The Autonomic Nervous System, Part 1, Cambridge: W. Heffer. Lewis, J. (1927). The Blood Vessels o f the Human Skin and Their Responses, London: Shaw & Sons. Loewi, O. (1921). Über humorale Übertragbarkeit der HerznervenWirkung. XI. Mitteilung. Pflügers Arch. Gesamte Physiöl., 189, 239-242. Loewy, A.D. and Spyer, K.M. (1990). Central Regulations o f Autonomie Functions, New York: Oxford University Press. Maggi, C.A. and Meli, A. (1988). The sensory-efferent function of capsaicin-sensitive sensory nerves. Gen. Pharmacol., 19, 1-43. Marrazzi, A.S. (1939). Electrical studies on the pharmacology of autonomic synapses. II. The action of a sympathomimetic drug (epinephrine) on sympathetic ganglia. J. Pharmacol. Exp. Ther., 65, 395-404. Palmer, R.M.J., Rees, D.D., Ashton, D.S. and Moncada, S. (1988). L-arginine is the physiological precursor for the formation of nitric oxide in endothelium-dependent relaxation. Biochem. Biophys. Res. Commun., 153, 1251-1256. Ralevic, V. and Burnstock, G. (1993). Neural-Endothelial Interactions in the Control o f Local Vascular Tone. pp. 1-117. Medical Intelligence Unit, R.G. Landes Company, Medical Publishers, Austin, Texas. Rand, M. J. (1992). Nitrergic transmission: nitric oxide as a mediator of non-adrenergic, non-cholinergic neuro-effector transmission. Clin. exp. Pharm. & Physiol. 19, 147-169. Snyder, S.H. (1992). Nitric oxide: first in a new class of neurotransmitters? Science 257, 494-496. Vizi, E.S. (1979). Prejunctional modulation of neurochemical transmission. Prog. Neurobiol., 12, 181-290. von Euler, U.S. and Gaddum, J.H. (1931). An unidentified depressor substance in certain tissue extracts. J. Physiol., 72, 74-87. Yanagisawa, M., Kurihara, H., Kimura, S., Tomobe, Y., Kobayashi, M., Mitsui, Y., Yazaki, Y., Goto, K. and Masaki, T. (1988). A novel potent vasoconstrictor peptide produced by vascular endothelial cells. Nature, 332, 411-415.

Preface Langley (1898) proposed the term ‘autonomic nervous system\ and provided a generally acceptable subdivision into three parts: the sympathetic, the parasympathetic and the enteric divisions. The classification is based primarily on the anatomy of mammals and later studies of nerve function helped to firmly corroborate this subdivision. A separation of the sympathetic and the posterior parts of the parasympathetic division may be difficult in some of the non-mammalian vertebrates. In addition, it is often impossible to decide whether neurons of the enteric nervous system are sensory or motor. Despite this, the classification made by Langley has withstood well the challenges of nearly the past 100 years (see Chapter 1, this volume, for a full discussion on autonomic nervous anatomy). The progress of our knowledge about the nature and function of the autonomic nerves appears soundly linked to our ability to literally see the neurotransmitter. Common sense prescribes “/ will believe it when I see i t ” so there is little wonder that at least two of the major leaps forward in our understanding of autonomic nerve function are associated with the visualization of the neurotransmitters themselves. One of these events is the development of the fluorescence histochemical technique for monoamines (‘Falck-Hillarp technique’). The technique was developed by Nils-Ake Hillarp and Bengt Falck in the early 60’s on the basis of the observations by Olavi Eranko that catecholamines produce fluorescent products when exposed to formalin (Eranko, 1952, 1955; Falck et al., 1962). The technique clearly helped the triggering of a formidable explosion of studies into the localization, ultrastructure, physiology, pharmacology and biochemistry of adrenergic and other monoaminergic nerves, both autonomic and central. A second significant breakthrough was the development of immunochemical techniques (e.g., radioimmunoassay and immunohistochemistry) that make it possible to quantify and actually see several cell components. These include peptide neurotransmitters (peptidergic neurons), amines such as 5-hydroxytryptamine (serotonergic neurons) and ‘marker enzymes’ such as tyrosine hydroxylase and dopamine-j3-hydroxylase (adrenergic neurons) or choline acetyltransferase (cholinergic neurons). The neurotransmitter of the purinergic neurons, ATP or a related purine derivative, has been fighting an uphill battle when it comes to visualization: ATP and its metabolites are present in all living cells, and histochemical techniques xiii

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specific for these compounds will thus be inadequate as evidence for transmitter storage in neurons. The term ‘comparative physiology’ has come to mean physiology of nonmammalian animals (although, to some people, it may mean comparing guinea-pigs with rats). Knowledge of the anatomy and organization of the autonomic nervous system in non-mammalian vertebrates is reasonable, relatively speaking. This is to a large extent thanks to several, often painstakingly detailed, late nineteenth and early twentieth century anatomical descriptions covering all the major vertebrate groups. There were also early ground-breaking studies on the general physiology of autonomic nerve functions in which non-mammalian animals, often frogs, were used. Pioneering work on the functional anatomy and physiology of the autonomic nervous system in fish was done by John Z. Young in the early 1930’s. In these studies, Young used ‘pharmacological tools’ to manipulate and characterize the autonomic innervation of several organs in elasmobranch and teleost fish. Later the Melbourne laboratory, then led by Geoffrey Burnstock, conducted systematic research on the autonomic nerve functions in amphibians and reptiles besides their work on mammalian systems. Such comparative physiological studies of the autonomic nervous system provide information about the control and coordination of organs and organ systems in the non-mammalian vertebrates. In addition, the comparative approach opens another perspective on the function of vertebrate control systems by using the diversity of the animal kingdom as a variable. In this way, comparative physiology offers the potential of new general wisdom regarding the evolutions of nerve functions. The philosophy behind this volume is to summarize the current knowledge of the functional anatomy and physiology of the autonomic nervous system in the nonmammalian vertebrates. This aim frequently requires reference to the better-known mammals, and this group is therefore also included albeit less extensively. The ambition is to cover autonomic nerve functions in a selection of organs and organ systems, and also to describe the general anatomy of the autonomic nervous system in the vertebrate classes. Are there any evolutionary or phylogenetic trends and patterns in the design and function of the autonomic nerves? Perhaps this question would be best left to the reader of this volume. Detailed knowledge of autonomic nerve functions is restricted to a handful of mammals (guinea-pigs and rabbits, cats and rats, mice and men) and the number of extant vertebrate species is quite large (there are, for instance, more than 20,000 species of teleost fish). It would therefore seem that any phylogenetic discussions will still be biased more by the unevenly distributed research efforts, limited to a few organs of a few species, than by ‘true’ evolutionary patterns. Göteborg, June 1992, Stefan Nilsson Susanne Holmgren

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REFERENCES Eranko, O. (1952). On the histochemistry of the adrenal medulla of the rat, with special reference to acid phosphatase. Acta Anat., 16 (Suppl. 17), 308. Eranko, O. (1955). Distribution of adrenaline and noradrenaline in the adrenal medulla. Nature, 8, 88-89. Falck, B., Hillarp, N .-A ., Thieme, G. and Torp, A. (1962). Fluorescence of catecholamines and related compounds condensed with formaldehyde. J. Histochem. Cytochem., 10, 348-354. Langley, J.N. (1898). On the union of cranial autonomic (visceral) fibres with the nerve cells of the superior cervical ganglion. J. Physiol. (Lond.), 23, 240-270.

Contributors Campbell, Graeme

McLean, John R.

Gibbins, Ian L.

Morris, Judy L.

Department of Zoology, University of Melbourne, Parkville, Victoria 3052, Australia

Department of Zoology, University of Melbourne, Parkville, Victoria 3052, Australia

Department of Anatomy and Histology, and Centre for Neuroscience, School of Medicine, Flinders University of South Australia, Bedford Park, SA 5042, Adelaide, Australia Grove, David J. Nuffield Fish Laboratory, School of Ocean Sciences, University College of North Wales, Bangor, UK Holmgren, Susanne

Department of Zoophysiology, University of Göteborg, Medicinaregt, 18 S-41390 Göteborg, Sweden Jensen, Jörgen

Department of Zoophysiology, University of Göteborg, Medicinaregt, 18 S-41390 Göteborg, Sweden

Jönsson, Ann-Cathrine

Department of Zoophysiology, University of Göteborg, Medicinaregt, 18 S-41390 Göteborg, Sweden

xvii

Department of Anatomy and Histology, and Centre for Neuroscience, School of Medicine, Flinders University of South Australia, Bedford Park, SA 5042, Adelaide, South Australia Nilsson, Stefan Department of Zoophysiology, University of Göteborg, Medicinaregt, 18 S-41390 Göteborg, Sweden Santer, Robert M.

Department of Anatomy, University of Wales College of Cardiff, PO Box 900, Cathays Park, Cardiff CF1 3YE, UK Uematsu, Kazumasa

Faculty of Applied Biological Science, Hiroshima University, 1-1-89, Higashisenda-machi, Naka-ku, Hiroshima 730, Japan

1 Comparative Anatomy and Evolution of the Autonomic Nervous System Ian Gibbins D epartm ent o f A n a to m y and H istology, and Centre f o r Neuroscience, School o f M edicine, Flinders University o f South Australia, A delaide, Australia. This chapter presents an overview of the functional anatomy of the peripheral autonomic nervous system of each of the vertebrate classes. When combined with developmental studies, there are several clues as to the likely course of evolution of the different divisions of the autonomic nervous system. The enteric nervous system probably is the most primitive division, with apparently homologous neurons occurring in cephalochordates. An oculomotor pathway to ocular tissues is present in all gnathostome classes. Cranial autonomic innervation of the upper jaw via the palatine branch of the facial nerve first appears in amphibians. The cranial autonomic innervation of the lower jaw is primitively derived from the posttrematic branches of the facial and glossopharyngeal nerves, as seen in elasmobranchs. In amphibians, the glossopharyngeal pathway predominates, but in amniotes, the facial pathway is dominant. The vagus provides autonomic pathways to the foregut and its derivatives, and to the heart, via pathways that seem to be highly conserved. The sympathetic division is likely to have evolved from aggregations of chromaffin tissue primarily associated with the origins of the segmental blood vessels and the posterior cardinal veins. The pelvic plexuses are a complex mixture of pathways containing sympathetic and sacral parasympathetic components, mainly involved with the control of the hindgut and its .derivatives. Overall, there is a clear evolutionary trend for an increase in the level of organisation of autonomic pathways. This trend presumably is related to a corresponding increase in the ability of vertebrates to flourish independently of environmental constraints. KEY WORDS sympathetic; parasympathetic; enteric; anatomy; evolution

ANATOM ICAL DIVISIONS OF THE AUTONOM IC NERVOUS SYSTEM Since the pioneering work of John Newport Langley at the turn of the century, the autonomic nervous system (ANS) has been considered to consist of three main divisions (Langley, 1898, 1903, 1921). The divisions were defined by a combination of anatomical and functional criteria, primarily derived from studies of the mammalian ANS. The three primary divisions recognised are: the sympathetic division, l

2 COMPARATIVE PHYSIOLOGY AND EVOLUTION

the parasympathetic division, and the enteric division. A major criterion for distinguishing these divisions was the level of origin of the preganglionic inputs to the peripherally located postganglionic neurons. The distribution of cell bodies of sympathetic preganglionic neurons is restricted to the thoracic and lumbar levels of the spinal cord. Parasympathetic preganglionic neurons are found both in the brain stem and in the sacral spinal cord, whilst the great majority of enteric neurons lack any direct preganglionic input. Langley himself recognised that the parasympathetic division as defined above is a somewhat artificial assemblage. Consequently, he further divided it into three components, again based upon the locations of the preganglionic neurons (Langley, 1903, 1921). First, the cranial and sacral parasympathetic outflows clearly were seen as being separate in both anatomical and functional terms, although they share more characters in common with each other than with the sympathetic division. Then, the cranial component of the parasympathetic division can be considered to comprise two distinct parts. One part is formed by the parasympathetic pathway to the eye. This outflow is associated with the oculomotor nerve (third cranial nerve; CN III), with preganglionic cell bodies in the midbrain, and postganglionlic nerve cell bodies in the ciliary ganglia. The other part of the cranial parasympathetic outflow consists of pathways associated with the facial (seventh cranial nerve; CN VII), glossopharyngeal (ninth cranial nerve; CN IX) and vagus (tenth cranial nerve; CN X) nerves. These nerves are associated with the branchial arches and their derivatives and the corresponding preganglionic neurons are all located in the hindbrain. When a wide range of species is examined, the distinction between the sympathetic outflows and the sacral parasympathetic outflows also may appear to be somewhat artificial (Young, 1933; see Nilsson, 1983). Consequently, the terms “cranial autonomic outflow” and “spinal autonomic outflow” often will be used here rather than “sympathetic” and “parasympathetic” to describe the fundamental divisions of the ANS. Furthermore, in the pelvic region, it sometimes may be impossible to assign any particular neuron unambiguously to a sympathetic or parasympathetic pathway. The evidence for this conclusion will be presented further below, but in the meantime the term “pelvic plexus” will be used for autonomic pathways in the pelvic region, without any implication as to the nature of those pathways (see Gibbins, 1990a). Over the last twenty years, there have been very many studies on the structure and function of the ANS in different vertebrate species. In particular, modern neuro-anatomical studies have used a variety of labelling techniques, including immunohistochemistry and retrograde axonal tracing, to provide a detailed picture of the distribution and function of specific populations of autonomic neurons. These experiments have shown that there is a general organisational principle, termed “chemical coding”, that applies to much of the nervous system including the ANS (Costa, Furness and Gibbins, 1986; Furness et al., 1989). This principle is derived from observations that a single autonomic neuron commonly may contain many different potential neurotransmitters. Most prominent of these transmitters are the neuropeptides. The precise combination of peptide and non-peptide trans-

COMPARATIVE ANATOMY AND EVOLUTION

3

mitters found in any particular neuron generally is related to its terminal projection. In other words, the combinations of transmitters form “chemical codes” that can be used to identify different functional populations of neurons. Moreover, distinctive combinations of neuropeptides may occur in homologous populations of neurons in different species, implying that there are also species-specific factors determining the expression of the chemical codes. However, to date there have been no obvious evolutionary trends detected in the chemical coding of autonomic neurons across the vertebrate classes (see Gibbins, 1989). The functional consequences of the co-existence of many different transmitters in autonomic neurons are complex, and are beyond the scope of this chapter. For recent reviews of this area see Furness et al. (1989) and Morris and Gibbins (1992). In this chapter, I will compare what is known about the functional anatomy of the ANS in each of the vertebrate classes both in the context of Langley’s scheme and of more recent neurochemical studies. Such a comparison should help elucidate some of the general principles of organisation of the ANS as well as their evolution. For more information and access to the literature on the non-mammalian ANS, the reader is referred to the excellent reviews by Hirt (1934), Nicol (1952), Burnstock (1969), Pick (1970) and Nilsson (1983). Somewhat more restricted reviews by Huber (1900) and Kuntz (191 lb) continue to offer valuable insights for the modern reader.

THE STRUCTURE OF AUTONOM IC NEURONS Peripheral autonomic neurons have a wide range of shapes and sizes that vary both within and between species (for detailed descriptions, see Huber, 1900; Gabella,’ 1976). In most cases, the full extent of the dendritic tree of autonomic neurons can only be revealed by certain types of stains, such as some silver impregnation methods or methylene blue under optimal conditions. More recently, much information about the shapes of mammalian postganglionic and enteric neurons has been obtained by injecting them intra-cellularly with markers that completely fill a cell and its processes (e.g. Purves, 1988; Bornstein and Furness, 1988). There have been several schemes proposed for classifying autonomic neurons according to morphological criteria, such as the number and branching patterns of their dendrites, or the overall size of the cell body (e.g. Cajal, 1911; De Castro, 1932; Dogiel, 1896; Gunn, 1951, 1968; Kuntz, 1945). However, none of these schemes has gained universal acceptance. The most simple autonomic neurons are the postganglionic neurons lying in cranial and spinal autonomic pathways of amphibians (Figures l.la,c). These cells are globular or ellipsoid in shape. They are generally considered to be monopolar, bearing a single axon but no dendrites. However, recent experiments in which individual neurons were filled with an intracellular injection of horseradish peroxidase have shown that many of these neurons have prominent axon collaterals that probably end within the ganglia (Forehand and Konopka, 1989). Monopolar neurons also occur in the parasympathetic and pelvic ganglia of small mammals. However, their homologues in larger mammals are multipolar (Figure 1.2a).

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COMPARATIVE PHYSIOLOGY AND EVOLUTION

FIGURE 1.1 Neurons in wholemounts of the vagus nerve and myenteric plexus of the toad, Bufo marinus (see also Figure 1.8). a) Neurons containing immunoreactivity to somatostatin in the vagus. These neurons are monopolar and project to the lung and pulmonary artery. In addition to containing somatostatin, they are also cholinergic, b) Neuron containing somatostatin in the myenteric plexus of the duodenum. Its single axon projects anally (towards the bottom of the plate). Note the prominent lamellar dendrites, c) Monopolar neurons containing immunoreactivity to vasoactive intestinal peptide (VIP) in the distal oesophageal ramus of the vagus, d) VIP-containing neurons in a node of the myenteric plexus of the stomach. Unlike the vagal VIP neurons, the enteric neurons have prominent dendritic processes.

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5

FIGURE 1.2 a) Neurons containing immunoreactivity to VIP in a section of the sphenopalatine ganglion of a guinea-pig, Cavia. The neurons have prominent dendrites, which are intermeshed to form a dendritic glomerulus (star). Similar glomeruli also occur in sympathetic ganglia, b) Binucleate neurons containing VIP in a section of a lumbar sympathetic ganglion of a guinea-pig. c) Chromaffin cells with long processes in a whole-mount of the inter-atrial septum of the urodele amphibian, Necturus maculosus. These cells act functionally as interneurons. Glyoxylic acid induced fluorescence, d) Section of a sympathetic ganglion of a toad, Bufo marinus, labelled for immunoreactivity for neuropeptide Y (NPY). NPY is found only in the small C-cells (arrow heads) projecting to blood vessels, and is absent from the large C-cells (arrows) innervating glands in the skin.

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COMPARATIVE PHYSIOLOGY AND EVOLUTION

The postganglionic neurons in the sympathetic ganglia of most vertebrates generally are multipolar (Huber, 1900; Terni, 1931; Gabella, 1976). They commonly have between six and ten primary dendrites, which themselves may bear many branches. Cajal (1911) has observed that the dendrites on sympathetic neurons of birds are remarkable for their length and complexity. In larger mammals, such as humans, the dendrites may branch profusely and intertwine with those of other neurons to form dendritic glomeruli (Cajal, 1911; De Castro, 1932). Similar glomeruli have been observed in cranial parasympathetic ganglia of mammals (Figure 1.2a). In some cases the dendrites may completely encircle other postganglionic neurons. The function of such close associations between postganglionic neurons is unknown, but they raise the possibility that such neurons may participate in direct interactions with each other. Although most peripheral autonomic neurons have only a single nucleus (which is not surprising!), multinucleate neurons have been reported in some species. Most notable of these are elasmobranchs and guinea-pigs. In elasmobranchs, some sympathetic neurons may contain up to five nuclei (Young, 1933; Pick, 1970), whereas the majority of sympathetic neurons in guinea-pigs are binucleate (Huber, 1900; Figure 1.2b). The functional consequences of these odd arrangements are unknown. In addition to the principal neurons of various shapes and sizes, most autonomic pathways include chromaffin cells. These cells contain high levels of catecholamines and make up the adrenal chromaffin tissue. They also may occur as scattered clumps of cells known as paraganglia, or as small groups of cells within ganglia (see below, and Nilsson, 1983). In general, chromaffin cells are small, ovoid, typically 10-15 ¿on in diameter. When clumped together, the majority of chromaffin cells lack any substantial processes. When present, processes of such cells are relatively short. However, more isolated cells may possess a single process many tens of microns long which is varicose in a manner similar to that of axons of postganglionic neurons. Examples of such cells have been reported in the pelvic plexus of guineapigs (Furness and Costa, 1976) and the systemic arches of the sleepy lizard (Tiliqua rugosa; Berger et al.y 1982). In some locations, chromaffin cells may be bipolar (e.g. toad pulmonary artery; Haller and Rogers, 1978) or bear a number of short processes. The most complex chromaffin cells described to date are those in the cardiac plexus of urodele amphibians (Figure 1.2c). These cells are multipolar and probably function as interneurons (see below). Over the last ten years, it has become clear that the overall shapes and sizes of peripheral autonomic neurons are related to their functions. For example, in sympathetic ganglia of anuran amphibians and mice, vasomotor neurons are significantly smaller than secretomotor neurons (Morris et al., 1986a; Horn, Stofer and Fatherazi, 1987, 1988; Gibbins, 1991; Figure 1.2d). These size differences are correlated with differences in the conduction velocities of action potentials so that the larger secretomotor neurons conduct action potentials faster than the smaller vasoconstrictor neurons. The size of neuronal cell bodies together with the extent and complexity of their dendritic arborizations seem to be determined by the volume of their terminal innervation field. Furthermore, the degree of convergence of preganglionic inputs onto postganglionic cell bodies is related to the size and

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7

complexity of their dendritic trees. Thus, larger neurons with more complex dendrites generally innervate larger volumes of target tissue and receive more synaptic inputs. This trend is observed both within and between individual animals. One consequence of these observations is that the number of postganglionic neurons in a particular pathway increases much less than might be expected as animals increase in size during either their developmental or evolutionary history (see Purves and Lichtman, 1985; Purves, 1988).

THE ENTERIC DIVISION The enteric division of the ANS probably contains more neurons than any of the other autonomic divisions. It comprises a large collection of interconnected neurons located within the wall of the gastro-intestinal tract from the oesophagus to the rectum. The enteric neurons originate from a population of neural crest cells, distinct and separate from the rest of the ANS (see Horstadius, 1950; Le Douarin, 1982). In general, the enteric division has several features that are unique among autonomic pathways. First, most of the enteric neurons do not receive any direct preganglionic input from the central nervous system. Second, enteric neurons include not only various functional classes of motor neurons, but also interneurons (or associative neurons) and sensory neurons. As a consequence of these characteristics, the enteric nervous system can generate and modify reflex activity independently of the rest of the nervous system. Nevertheless, the activity of enteric neurons can be influenced by the central nervous system via direct synaptic inputs from central neurons projecting down along the vagus to the foregut, and along the sacral spinal nerves to the hindgut. Furthermore, enteric nervous activity can be modified by sympathetic neurons projecting to the gastro-intestinal tract (for details, see Furness and Costa, 1987). In mammals, the enteric neurons are collected into two main ganglionated plexuses. The myenteric plexus (Auerbach’s plexus) lies between the outer longitudinal layer and the inner circular layer of smooth muscle of the muscularis externa. The submucous plexus (Meissner’s plexus) lies in the submucosal connective tissue underlying the epithelium of the gut. However, submucosal ganglia are rare or absent in the oesophagus and stomach. There are numerous connections between the two plexuses, and they cooperate in the regulation of smooth muscle motility, secretion and absorption, and local blood flow. The precise arrangement of the myenteric and submucous ganglia varies both along the gut in a single species and in the same region of the gut in different species. A limited immunohistochemical study of enteric neurons in the intestine of a monotreme, the platypus (Ornithorhyncus: Osborne, Campbell and Evans, 1989), suggests that the myenteric plexus in these primitive mammals may be more loosely organised than those in placentals. Recent experimental studies combining microanatomical and physiological approaches have shown that there are very many different functional populations of enteric neurons, and that they are connected in a precise and highly organised

8 COMPARATIVE PHYSIOLOGY AND EVOLUTION

way. For example, in the small intestine of guinea-pigs, Dogiel (1896) originally described two main morphological classes of neurons in the myenteric plexus. Type I cells had smaller cell bodies with short branching dendrites, often with a somewhat lamellar appearance, and with a single axon. Type II cells were larger and bore several long smooth processes with limited branching. None of these processes could be identified unambiguously as an axon. It is now clear that the type I cells are motor neurons and interneurons, and the type II cells are sensory neurons (see Furness and Costa, 1987). From the relatively small amount that is known, the overall arrangement of the enteric plexuses in other amniotes seems to be remarkably similar to that in mammals, despite some differences in the structure of the gastrointestinal tract itself (Nilsson, 1983). For example, in chelonians, there is both a well developed ganglionated myenteric and a submucosal plexus for the whole length of the gut (Geoclemys: Hukuhara et al., 1976; Pseudemys: Timmermans et al., 1991). A similar situation probably applies in birds, although there are few, if any, submucous ganglia in the gizzard (Bennett, 1969, 1974; Bennett and Cobb, 1969; Ali and McLelland, 1978). The enteric plexuses of amphibians are less highly organised than those of amniotes. In the myenteric plexus of the intestines, nerve cell bodies are not aggregated into distinct ganglia. Instead, neurons are scattered along nerve fibre tracts running more or less longitudinally between the longitudinal and circular muscle layers (Bufo: Boyd, Burnstock and Rogers, 1964; Wong et al., 1971; Gibbins, 1981; Rana: Klein, 1873; Gunn, 1951; Torihashi, 1990; Salamandra: Buchan, Polak and Pearse, 1980; Necturus: Holmgren et al., 1985). However, in the stomach, the nerve fibre bundles have a more plexiform arrangement and small groups of neurons occur at the nodes of the plexus (Cole, 1926; Gibbins, 1981; Figure 1. Id). In general, the submucous plexus lacks nerve cell bodies (Müller, 1908; Gunn, 1951; Wong et al., 1971; Gibbins, 1981). The neurons are distinctive in that they commonly are very irregular in shape with broad lamellar processes rather than fine branching dendrites (Rana: Cole, 1925; Gunn, 1951; Torihashi, 1990; Bufo: Gibbins, 1981; Figure 1.2a,d). Overall, the microanatomy of the enteric plexuses in the various groups of fish is similar to that in amphibians (Nilsson, 1983; Bjenning and Holmgren, 1988). The intrinsic neurons generally do not form well organised ganglia, and submucosal neurons are rare or absent (Acipenser, Silurus: Kolossow and Iwanow, 1930; Scylliorhinus, Saccobranchus, Ophiocephalus: Kirtsinghe, 1940; Tinea: Baumgarten, 1965; Parasilurus: Hukuhara and Fujiwara, 1975; Chelmon: Wong and Tan, 1978; Platycephalus: Anderson, 1979; Salmo: Ezeasor, 1979; Holmgren et al., 1982; Myxocephalus, Pleuronectes: Watson, 1981). Although ganglia are usually absent, the neurons tend to be more aggregated at junctions between fibre bundles in the myenteric plexus, especially in the stomach and oesophagus. The neurons themselves usually are irregular in shape, often bearing broad lamellar processes, reminiscent of those on amphibian enteric neurons (Kolossow and Iwanow, 1930; Kirtsinghe, 1940). However, within the intestine of some teleost species, there is a distinctive population of very small spindle-shaped neurons, usually monopolar,

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that contain large amounts of serotonin (Salmo, Carrasius, Platycephalus, Aldrichetta, but not Anguilla, Torquigener, Rhombosolea: Anderson, 1983). Such neurons have not been described in the enteric plexuses of any other vertebrate group. The intestine of lampreys (Lampetra) contains a loose network of nerve fibres and irregularly shaped, multipolar neurons. The neurons are especially prominent near the origin of the liver and in the rectal part of the small intestine (Johnels, 1956). Most of these neurons contain serotonin (Baumgarten et al., 1973; Sakharov and Salimova, 1980). A plexus of neurons with similar morphology to the enteric neurons of lampreys occurs in the intestine of Amphioxus (= Branchiostoma), a cephalochordate. These neurons probably comprise the closest homologue to ah ANS in non-vertebrates (Boeke, 1935; Kirtsinghe, 1940). These observations suggest that the enteric neurons form a primitive component of the ANS. In the absence of any information regarding their functions in amphioxus and lampreys, it is impossible to say if both sensory and motor enteric neurons were present primitively. Nevertheless, it seems clear that there is a significant increase in the level of organisation between the anamniotes and the amniotes. Presumably, this is related to the increased demand for precise control, not only of digestive function itself, but also of water and electrolyte balance, in animals adapted for life on land.

CRANIAL AUTONOM IC OUTFLOWS Following the lead of Langley, it is clear that the cranial autonomic (parasympathetic) outflows fall into two natural groups: the pathway to the ciliary ganglion following the oculomotor nerve (CN III); and the pathways associated with the facial nerve (CN VII), glossopharyngeal nerve (CN IX) and vagus nerve (CN X), all of which are associated with the branchial arches and their derivatives. CILIARY GANGLION AND THE OCULOMOTOR PATHWAY The ciliary ganglion and its relationships to the surrounding neuronal pathways is one of the most highly conserved components of the ANS. A ciliary ganglion is present in most vertebrates (e.g. Figure 1.11). Notable exceptions include cyclostomes (Johnston, 1905; Cords, 1929; Pick, 1970), and groups that are secondarily eyeless, such as caecilian amphibians (Norris and Hughes, 1918), or that have rudimentary eyes, such as the paddlefish, Polyodon (Norris, 1925) or the urodele amphibian, Amphiuma (Norris, 1908). In some species, there is not a discrete ganglion, but rather just a series of loose aggregations of ganglion cells scattered along the ciliary nerves, forming a “ciliary plexus” (Norris and Hughes, 1920; Figure 1.3). This arrangement has been described in such diverse groups as selachians [Squalus, Mustelus: Norris and Hughes, 1920; Scyllium (= Scylliorhinus): Young, 1933], urodele amphibians (Desmognathus: Strong, 1890; Ambystoma: Herrick, 1894; Coghill, 1902), anuran amphibians (Rana, Xenopus: Taxi, 1976); snakes

FIGURE 1.3 Simplified diagram of the cranial nerves of a shark, Squalus. Sites where autonomic ganglion cells occur are indicated by filled dots. There are two main locations: the ciliary plexus, and the post-trematic rami of the facial, glossopharyngeal and vagus nerves. Sensory ganglia are indicated with dense stipple. The position of the orbit is shown by a light dotted line. Adapted from Norris and Hughes (1920). Ill, oculomotor nerve; Vj sup, superficial ramus of the ophthalmic division of the trigeminal nerve; prof, ramus of the ophthalmic division of the trigeminal nerve (the “profundus”); V-VII, common roots of the trigeminal (V) and facial (VII) nerves; V2-VII, maxillary division of the trigeminal nerve and accompanying facial nerve fibres; V3-VII, mandibular division of the trigeminal nerve and accompanying facial nerve fibres; VIIhyo, hyomandibular ramus of the facial nerve; VIImd, internal and external mandibular rami of the facial nerve; VIIpal, palatine ramus of the facial nerve; IX, glossopharyngeal nerve; X, X j, X2, X3, X4, vagus nerve and its main branchial rami; Xvisc, visceral ramus of the vagus.

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(Elaphe: Auen and Langebartel, 1977) and guinea-pigs (Watanabe, 1972; Gibbins and Morris, 1987). In some cases, the “ciliary ganglia” are so small and diffuse that they may not be visible macroscopically (Coghill, 1902; Norris and Hughes, 1920; Young, 1933; Gibbins and Morris, 1987). In all likelihood, these small ganglia occur at constant locations within the ciliary plexus (Norris and Hughes, 1920; Young, 1933). Even in species that have a discrete ciliary ganglion, an “accessory ciliary ganglion” or other groups of scattered ganglion cells may be present along the ciliary nerves, right up until their entry into the sclera. Accessory ciliary ganglia have been described in holocephalan elasmobranchs (Chimaera: Cole, 1896; Hydrolagus: Nicol, 1950), teleosts (Lampanyctus: Ray, 1950), amphibians (Rana: Strong, 1890) and a variety of mammals including rats (Stone et al., 1988), cats, macaque monkeys, humans (Grimes and von Sallman, 1960; Bryson et al., 1966). Although preganglionic fibres reach ciliary ganglion cells via the oculomotor nerve, the ciliary ganglion is closely associated with the first (ophthalmic) division of the trigeminal nerve. In elasmobranchs, teleosts and reptiles, this branch of the trigeminal nerve originates from a separate division of the trigeminal sensory ganglion and is referred to as the ramus ophthalmicus profundus or, simply, “the profundus” (e.g. Herrick, 1899; Willard, 1915; Norris and Hughes, 1920; Young, 1931; Ray, 1950; see Nilsson, 1983). Situated on the profundus nerve is a separate sensory ganglion, which in many early papers was confused with the ciliary ganglion (see Beard, 1887, and Norris and Hughes, 1920, for references and discussions of this point). The ciliary ganglion contains several different functional populations of postganglionic neurons, all of which usually are assumed to be cholinergic. It innervates the iris and surrounding structures, such as the ciliary muscles and, in at least some cases, the choroidal blood vessels. In some species, the different functional classes of neurons may be distinguished morphologically. This may be seen most dramatically in the ciliary ganglion of birds. The ganglion contains about 6000 neurons in pigeons and turkeys, and about 3000 neurons in chickens (see Gabella, 1976). At least half of the neurons are large and have myelinated axons; they innervate the striated muscle of the iris and ciliary bodies. The smaller cells innervate the smooth muscle and vasculature of the choroid. Not only can these different classes of neurons be distinguished by their size, but they can also be distinguished by the distinctive nature of their respective preganglionic inputs. In young birds, the large neurons are surrounded by large calyciform (“cup-shaped”) endings which develop into tight clusters of terminals surrounding the cell bodies. Each postganglionic neuron receives an input from only one preganglionic neuron. Ganglionic transmission occurs via a mixture of chemical (cholinergic) and electrical transmission (see Bennett, 1974; Gabella, 1976). The smaller neurons have more conventional preganglionic chemical synapses. Recent studies in pigeons have demonstrated that the two preganglionic pathways arise from different subnuclei in the brain, clearly indicating that pupillary diameter and accommodation can be controlled independently of choroidal blood flow (Reiner et al., 1983; Gamlin and Reiner, 1991). The iris of reptiles also contains striated muscle (see Nilsson, 1983), but observations of the ciliary ganglion in chelonians suggest that morphologically

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different populations of neurons are not present, at least in this group of reptiles (Soliman, 1964). On the other hand, the ciliary ganglion of Menidia, a teleost fish, contains a distinctive collection of larger ganglion cells at its caudal end (Herrick, 1899). However, their target is unknown. The ciliary ganglion cells of mammals generally are considered to be morphologically homogeneous. In smaller species, such as rodents, they are mostly monopolar. Even in larger species, such as cats or humans, they have only relatively limited dendritic processes (Slavich, 1932; Warwick, 1954; Purves, 1988). In laboratory mammals, each postganglionic ciliary neuron has only a few (up to 6) preganglionic inputs, with a larger number of inputs being found on cells with more complex dendritic geometry (Hume and Purves, 1983). Functional evidence indicates that there are at least three different populations of postganglionic ciliary neurons: neurons innervating the constrictor and dilator muscles of the iris; neurons innervating the ciliary muscles; and neurons providing vasodilator innervation to the ciliary bodies, the choroid and surrounding blood vessels. Experiments in which single ciliary ganglion cells have been injected with horseradish peroxidase have shown that a single neuron can innervate both the constrictor and dilator muscles of the iris (Jackson, 1986). However, neither these experiments nor any others have shown any correlations between the morphology of mammalian ciliary ganglion cells and their specific targets. Recently, immunohistochemical studies have provided evidence that different populations of ciliary ganglion cells in some mammalian species can be distinguished by their content of neuropeptides (rats: Stone et al., 1988; but probably not guineapigs: Gibbins and Morris, 1987). In addition, some ciliary ganglion cells contain enzymes associated with the synthesis of catecholamines (tyrosine hydroxylase and dopamine-/3-hydroxylasfe), although few if any ciliary ganglion cells normally synthesise detectable levels of catecholamines (rats, cats, dogs, monkeys: Landis et al., 1987; Uemura et al., 1987). Nevertheless, it is still not clear how these immunohistochemically identified classes correlate with the functionally characterised populations of ciliary ganglion cells. Neurons containing vasoactive intestinal peptide (VIP) are likely to be the vasodilator neurons. However, they may receive their preganglionic input primarily via the facial nerve rather than the oculomotor nerve (Stjernschantz and Bill, 1980; Ruskell, 1985). Indeed, there is good neurochemical evidence that VIP-containing neurons in the sphenopalatine ganglion innervate the iris, ciliary bodies and uveal blood vessels of guinea-pigs, rats and cats. (Uddman et al., 1980; Bjorklund et al., 1985; Gibbins and Morris, 1987). These neurons clearly lie on a facial nerve pathway (Ruskell, 1985; see below). There is some circumstantial evidence that a close association between oculomotor and facial nerve pathways is a long-standing one in the evolution of the vertebrates. Thus, a branch of the palatine ramus of the facial nerve enters the ciliary plexus of selachians. At its junction with the ciliary nerves, there is a small ganglion that apparently projects to nearby blood vessels, including the efferent branchial artery of the pseudobranch (Norris and Hughes, 1920; Young, 1933). The details of the connections and pathways of the autonomic components of the facial nerve will be discussed in the following section.

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THE BRANCHIOMERIC CRANIAL AUTONOMIC PATHWAYS: THE FACIAL (CN VII), GLOSSOPHARYNGEAL (CN IX) and VAGUS (CN X) NERVES The facial, glossopharyngeal and vagus nerves are complex nerves providing motor, sensory and autonomic innervation to the branchial arches and their derivatives. Around the turn of the century, there was considerable interest in the homologies and development of these cranial nerves in an attempt to understand their evolutionary history. The studies provided a large body of information about the neuroanatomy of these pathways in a wide variety of vertebrates, especially in the anamniotes. Some of the most useful early studies of the different groups are: cyclostomes: Johnston, (1905); elasmobranchs: Cole (1896); Norris and Hughes (1920); Young (1933); teleosts: Herrick, (1899, 1900); Ray (1950); ganoid fish: Norris (1925); anurans: Strong (1895); caecilians: Norris and Hughes (1918); urodeles: Herrick (1894); Coghill (1902); lizards: Willard (1915); chelonians: Ogushi (1913); Soliman (1964). Tissues in the upper jaw are generally innervated by autonomic pathways running with the facial nerve. In all vertebrates, the facial nerve divides into an anterior, or dorsal, component and a posterior, or ventral, component, known as the hyomandibular ramus. In non-mammalian classes, the anterior component possesses a major branch known as the palatine ramus (or palatine nerve; Figures 1.3, 1.6, 1.7, 1.11), which runs rostrad from the roots of the facial nerve to the floor of the orbit. Here, it ramifies to varying degrees and generally sends anastomosing rami to the first, or ophthalmic, division of the trigeminal nerve (the ramus ophthalmicus profundus) and to the second, or maxillary, division of the trigeminal nerve (the infra-orbital nerve). Except in elasmobranchs and caecilian amphibians, there generally are also communications between the glossopharyngeal nerve and the facial nerve (e.g. “Jacobson’s anastomosis”; Figure 1.7). Thus, an infra-orbital plexus is formed, which, in principle, could provide innervation to the palate and any related structures in the upper jaw. Preganglionic input to this plexus would arise primarily from the facial nerve, with perhaps some contribution from the glossopharyngeal nerve. Whereas secretomotor and vasodilator innervation of targets in the upper jaw arises from the palatine nerve pathway, corresponding targets in the lower jaw seem to be innervated via the chorda tympani and glossopharyngeal nerves. The status of the chorda tympani in different vertebrate groups is controversial (Cole, 1896; Herrick, 1899; Norris and Hughes, 1920) and still has not been completely resolved (Lombard and Bolt, 1979). It is most likely to be a constant ramus of the hyomandibular branch of the facial nerve, and probably corresponds to the internal mandibular branch of the facial nerve of fish (Lombard and Bolt, 1979; Figures 1.3, 1.4, 1.7, 1.11). The vagus nerve does not usually provide an autonomic innervation to cranial tissues themselves in extant vertebrates: it carries extensive autonomic pathways mainly to the foregut and its derivatives. Unfortunately, the outstanding early work on cranial pathways generally has not

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FIGURE 1.4 Cranial autonomic ganglia and microganglia in a cat, Felis. Nearly all of the neurons in these ganglia contain immunoreactivity to VIP. The ciliary ganglion and the ganglia of the retro-orbital plexus are not shown. Adapted from Gibbins, Brayden and Bevan, (1984). V3, mandibular division of the trigeminal nerve; VII, facial nerve; VIIct, chorda tympani; VIIgspn, greater superficial petrosal nerve; VIIvid, Vidian nerve (nerve of the pterygoid canal); IX, glossopharyngeal nerve; IXipn, lesser petrosal nerve; IXtr , tympanic ramus of the glossopharyngeal nerve; cn, carotid nerve (sympathetic); In, lingual nerve; OG, otic ganglion; SCG, superior cervical ganglion (sympathetic); SLG, sublingual gland ganglion; SMG, submandibular ganglion; SPG, sphenopaiatine ganglion.

been followed up with modern multi-disciplinary techniques, except in the case of common laboratory mammals. Only in these species have detailed correlations been made between the neurochemistry, neuroanatomy and functions of cranial autonomic neurons. Consequently, I will begin this account with an overview of the autonomic pathways in the facial, glossopharyngeal, and vagal nerves of mammals, before turning to non-mammalian species. Mammals For many years, conventional wisdom has stated that there are two main ganglia associated with the autonomic outflows of the facial nerve and one main ganglion

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FIGURE 1.5 Cranial autonomic ganglia in cats and lizards, a) Wholemount of the chorda tympani, a branch of the facial nerve, showing a microganglion containing VIP-immunoreactive neurons. Cat. b) Wholemount of the lesser petrosal nerve, a branch of the glossopharyngeal nerve within the tympanic plexus, with a small microganglion containing VIP-immunoreactive neurons. Cat. c) Wholemount of the pre-orbital ganglion of a lizard, Tiliqua rugosa. The neurons contain immunoreactivity for galanin. d) Section of a ganglion associated with the submandibular salivary gland of a lizard, Pogona vitticeps. The neurons contain immunoreactivity for VIP.

16 COMPARATIVE PHYSIOLOGY AND EVOLUTION

in the glossopharyngeal nerve autonomic pathway. Thus, the sphenopalatine ganglion (also known as the pterygopalatine ganglion) and the submandibular ganglion lie on branches of the facial nerve, and the otic ganglion lies on a branch of the glossopharyngeal nerve. However, scattered all along the various ramifications of both the facial and glossopharyngeal nerves are many small aggregations of autonomic nerve cell bodies forming microganglia (Figure 1.4, 1.5). Furthermore, there are many interconnections between the facial and glossopharyngeal nerves, especially in the region of the tympanic and cavernous plexuses (Rosen, 1950; Suzuki and Hardebo, 1991; see below), and it is not always obvious from anatomical considerations alone within which pathway a particular ganglion or microganglion lies. All the neurons in the ganglia lying on facial and glossopharyngeal nerve pathways are thought to be cholinergic, although this has only been demonstrated directly for some of the ganglia in rats (Leblanc, Trimmer and Landis, 1987; Suzuki, Hardebo and Owman, 1990). However, as in the case of the ciliary ganglion, many of these neurons contain immunoreactivity for enzymes normally involved in the synthesis of catecholamines, such as dopamine-jS-hydroxylase. Nevertheless, the neurons themselves do not synthesise catecholamines (rats: Grzanna and Coyle, 1978; guinea-pigs: Gibbins, 1990b). In the species studied to date (cats, rats, guineapigs, pigs, humans), the great majority of postganglionic neurons in the facial and glossopharyngeal pathways contain neuropeptides, the most widespread of which is VIP (Figure 1.5a,b). Moreover, many of the neurons contain several other peptides, including substance P, neuropeptide Y (NPY) and enkephalin (Lundberg et al.%1988; Gibbins, 1990b; Leblanc et al.9 1987; Leblanc and Landis, 1988). Sphenopalatine ganglion. The sphenopalatine ganglion lies on the medial side of the orbit and receives preganglionic fibres from the facial nerve via the nerve of the pterygoid canal (the Vidian nerve) and the greater superficial petrosal nerve (Figure 1.4). Although the ganglion has close anatomical associations with the maxillary (second) division of the trigeminal nerve, no preganglionic nerves take that route. In many species (e.g. cats, guinea-pigs, humans: Gibbins et al., 1984; Gibbins, 1990a; Nomura and Matsuura, 1972), there is at least one small ganglion on the Vidian nerve, proximal to the main ganglion. Other small ganglia occur in fine nerves running from the sphenopalatine ganglion towards the eye as part of a retro-orbital plexus. The sphenopalatine ganglion contains predominantly secretomotor and vasodilator neurons supplying the nasal mucosa, the lacrimal and Harderian glands, and the uvea. It is also likely to provide vasodilator innervation to the muscles of the jaws and face. Recently, the sphenopalatine ganglion of guinea-pigs has been shown to provide an abundant innervation of hair-follicles and sebaceous glands in the facial skin, lips, and eyelids (Gibbins, 1990b). These neurons contain up to four co-existing neuropeptides, but their functions are unknown. Submandibular ganglion. The submandibular ganglion lies in close association with the lingual nerve, a major branch of the mandibular (third) division of the trigeminal nerve (Figure 1.4). Although it receives most of its preganglionic input from

COMPARATIVE ANATOMY AND EVOLUTION

17

the facial nerve via the chorda tympani, there is also likely to be some input from the glossopharyngeal nerve via connections with the chorda tympani in the tympanic plexus (Laage-Hellman and Stromblad, 1960; Reichert and Poth, 1933; Figure 1.4). In addition to supplying secretomotor and vasodilator innervation to the submandibular salivary gland, the ganglion is supposed also to innervate the sublingual gland (e.g. Kuntz, 1945). However, in at least some species, there are small groups of microganglia located near the hilum of the sublingual gland (e.g. humans: Chorobski and Penfield, 1932; cats: Gibbins et al., 1984; mice: Purves and Lichtman, 1987; guinea-pigs: Gibbins, 1990b; Figure 1.4. Microganglia also occur along the chorda tympani itself (Figure 1.5a), and may provide postganglionic fibres to either of these salivary glands (cats: Gibbins et al., 1984; rats: Ekstrom et al., 1984). In guinea-pigs, the neurons innervating the submandibular gland are neurochemically distinct from those innervating the sublingual gland, clearly indicating that each of these glands is regulated by a distinct population of neurons (Gibbins, 1990b). Otic ganglion. The otic ganglion lies just outside the base of the skull close to the mandibular division of the trigeminal nerve (Figure 1.4). Some species, such as goats and horses, actually possess a paired otic ganglion, one either side of the mandibular nerve, and interlinked by commissural fibres. In cats, there seems to be a complex of three or more microganglia rather than a single otic ganglion (Ohkubo, 1979). Guinea-pigs possess an intracranial accessory otic ganglion lying on the lesser petrosal nerve before it exits from the skull to join the main otic ganglion (Gibbins, 1990b). The otic ganglion receives most of its preganglionic input from the glossopharyngeal nerve via the lesser petrosal nerve. However, there is evidence for an additional preganglionic input from the facial nerve via the chorda tympani (Diamant and Wiberg, 1965; Reichert and Poth, 1933). The main target of the otic ganglion is the parotid salivary gland and probably small glands at the back of the tongue and pharynx, to which it supplies secretomotor and vasodilator innervation. The ganglion also provides vasodilator innervation to the blood vessels of the lower lip and jaw (Segade, Quintanilla and Nunez, 1987; Kaji et al., 1988), and the cerebral circulation (Suzuki, Hardebo and Owman, 1990; Suzuki and Hardebo, 1991). Recent retrograde tracing experiments have demonstrated that the secretomotor neurons are significantly larger than the vasomotor neurons in the otic ganglia of rats (Kaji, Maeda and Watanabe, 1991). A similar relationship has been found for sympathetic neurons in the superior cervical ganglia of mice (Gibbins, 1991; see below). Cranial microganglia. In addition to the main autonomic ganglia of the facial and glossopharyngeal nerve pathways, other aggregations of ganglion cells are found along various ramifications of these nerves. Although there seem to be some species differences, relatively constant collections of microganglia occur in the tympanic plexus (within the middle ear cavity; Figure 1.5b), the cavernous plexus (on the floor of the middle cranial fossa) and along the lingual nerve within the tongue (Gibbins et al., 1984; Ichikawa et al., 1988; Figure 1.4). These ganglion cells are surprisingly

18 COMPARATIVE PHYSIOLOGY AND EVOLUTION

abundant: in humans, there may be several thousand of them (Gibbins, 1990a). Little is known about the function of most of these ganglia, but they probably cause vasodilation of local blood vessels: they are non-noradrenergic and commonly contain VIP (Gibbins et al., 1984; Suzuki, Hardebo and Owman; 1990; Figures 1.5a, b). Furthermore, retrograde tracing studies have shown that a prominent microganglion within the cavernous plexus of rats innervates the cerebral circulation (Hara, Hamill and Jacobowitz, 1985; Hara and Weir, 1988; Suzuki, Hardebo and Owman, 1990). The intralingual neurons receive preganglionic inputs both from the facial nerve (via the chorda tympani; Hellekant, 1972) and the glossopharyngeal nerve (Fitzgerald and Alexander, 1969; Gomez, 1961). At least some of the neurons in the cavernous plexus are likely to lie on facial nerve pathways, but an additional preganglionic contribution from the glossopharyngeal nerve cannot be ruled out (see Gibbins, 1990a). Vagus. The autonomic pathways in the vagus nerve of mammals are complex, and will be discussed only briefly here (see Gibbins, 1990a, for details and references). The vagus provides autonomic innervation to the foregüt and its derivatives, including the oesophagus, stomach, duodenum, airways, thyroid gland and pancreas, in addition to the heart. There is also some anatomical evidence for a vagal innervation of more distal structures such as the proximal colon and kidney, although these observations require functional confirmation. Microganglia are scattered along the course of the vagus nerve (Dolgo-Saburoff, 1936; Botar et al., 1950) and also occur in plexuses associated with the target tissues, such as the heart, airways, thyroid gland and pancreas (e.g Dogiel, 1877; Ploschko, 1897; Perman, 1924; see Gibbins, 1990a). In the gastro-intestinal tract, the vagus provides preganglionic input to some enteric neurons within the myenteric plexus (see Furness and Costa, 1987). Any particular target tissue may be innervated by more than one vagal autonomic pathway, each of which may have opposing effects. For example, there are both excitatory and inhibitory vagal pathways to the smooth muscle of the airways and stomach of guinea-pigs and cats. In some cases, there may be multiple vagal pathways controlling a single effector response, such as acid secretion from the stomach of rats, dogs, or humans (for reviews, see Walsh, 1987, 1988; Furness and Costa, 1987; Gibbins, 1990a). A particularly subtle example of the differentiation of vagal pathways controlling the activity of a single effector can be seen in the innervation of the heart of dogs. A long series of elegant physiological and morphological experiments has demonstrated that each of the major microganglia located in or near the heart has a separate role in regulating the rate and force of heart beat (Randall, 1984). Indeed, the left vagus has functions different from those of the right vagus (Randall, 1984). Recent immunohistochemical observations in guinea-pigs suggest that some of these cardiac ganglia may be distinguished by their neuropeptide content (see Gibbins and Morris, 1991). In addition to the principal ganglion cells, the cardiac ganglia of most mammalian species also contain small collections of chromaffin cells (Jacobowitz, 1967).

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Fish One might expect that the basic arrangement of the cranial autonomic pathways of the facial, glossopharyngeal and vagus nerves would be more obvious in the various classes of extant fishes, in which there has been comparatively little gross morphological modification of the branchial apparatus from its ancestral vertebrate state. The status of the branchiomeric autonomic pathways in cyclostomes is difficult to establish from the literature. Almost certainly, there are no well defined autonomic ganglia along the facial and glossopharyngeal nerves in the head. In Lampetra and Petromyzon, collections of bipolar or multipolar cells have been described in branches of the facial nerve running beneath the orbit to supply the mucosa and blood vessels of the roof of the pharynx. Similar cells are also associated with the cerebral blood vessels (Iijima and Wasano, 1980), the linings of the anterior three gill sacs (Johnston, 1905, 1908) and the branchial rami of the vagus nerve (Cords, 1929; Nakao, 1981). These cells were interpreted as autonomic ganglion cells by Johnston (1908) and most later authors (e.g. Kuntz, 1911b; Hirt, 1934; Johnels, 1956; Pick, 1970). Recent ultrastructural work has confirmed this conclusion, and has provided good circumstantial evidence that these neurons innervate the vasculature of the gills and brain {Lampetra: Iijima and Wasano, 1980; Nakao, 1981). There is general agreement that the vagus nerve provides autonomic innervation to the heart (Nakao et al., 1981), and probably the gastrointestinal tract, of lampreys (see Nicol, 1952; Pick, 1970; Nilsson, 1983). There are clearly identified ganglion cells lying within branches of the vagus associated with the heart and the digestive tract (Hirt, 1934; Johnels, 1956). The cardiac neurons probably are responsible for a cholinergic excitation of the heart (see Chapter 6), but the actions of the gastrointestinal neurons are unknown. Ganglion cells are also associated with the heart of myxines, but their functional connections have not been identified (see Nilsson, 1983). In addition to ganglion cells, the heart of cyclostomes contains masses of small cells containing catecholamines. These cells are morphologically similar to chromaffin cells: they do not seem to be under vagal control (see Chapter 6 and Beringer and Hadek, 1973). As in cyclostomes, there are no well defined cranial autonomic ganglia associated with the facial and glossopharyngeal nerves of elasmobranchs (Chimaera: Cole, 1896; Squalus, Mustelus: Norris and Hughes, 1920; Young, 1933; Hydrolagus: Nicol, 1950), holosteans (Amia: Brookover, 1910; Norris, 1925), chondrosteans (Polyodon: Allis, 1920-21; Norris, 1925; Acipenser, Scaphyrhynchus: Norris, 1925), teleosts {Menidia: Herrick, 1899; Gadus: Herrick, 1900; Uranoscopus: Young, 1931; Lampanyctus: Ray, 1950; Trichiurus: Harrison, 1981) or dipnoans (Lepidosiren: Jenkin, 1928). Although a palatine ramus of the facial nerve supplies the palatine mucosa and glands in all these groups, there probably are no autonomic ganglion cells within it (see Cole, 1896). One possible exception is a small ganglion at the junction of an anastomosing ramus of the palatine nerve with the ciliary plexus of sharks (Norris and Hughes, 1920; Young, 1933). This conclusion needs

20 COMPARATIVE PHYSIOLOGY AND EVOLUTION

to be verified using modern microanatomical methods. Nevertheless, there is unlikely to be a significant cranial parasympathetic innervation of the palatine region in any group of fish. Whilst there is apparently no cranial parasympathetic innervation of structures in the upper jaw of fish, there is a cranial autonomic outflow to the branchial region. In selachian elasmobranchs, small groups of ganglion cells are associated with the hyomandibular ramus of the facial nerve and in the post-trematic ramus of the glossopharyngeal nerve (Norris and Hughes, 1920; Young, 1933; Figure 1.3). However, in chimaeroid (holocephalan) elasmobranchs, ganglion cells are absent from the facial nerve although they are present in the branchial rami of the glossopharyngeal nerve (Nicol, 1950). Corresponding groups of ganglion cells occur on the post-trematic branches of the vagus nerve of sharks (Norris and Hughes, 1920; Young, 1933; Figure 1.3) and chimaeroids (Nicol, 1950), but they have been claimed to be found in the pre-trematic rami in skates {Raja: Shore, 1889). Numerous postganglionic neurons occur along the post-trematic ramus of the glosso-pharyngeal nerve and in the pre- and post-trematic branchial rami of the vagus in teleosts {Perea, Salmo, Ictalurus, Sander, Micropterus: Bailly and DunelErb, 1986; Dunel-Erb et al., 1989; Gadus: I. Gibbins, unpub. obs. Figure 1.11). The function of these neurons is not known, but they may include the source of cholinergic vasoconstrictor innervation of the branchial vasculature in teleosts (Nilsson, 1984a; Bailly and Dunel-Erb, 1986; Dunel-Erb, Bailly and Lawent, 1989). The functions of the branchial ganglia in elasmobranchs are obscure (Nilsson, 1984a; Nilsson and Holmgren, 1988), but, in principle, they could innervate the smooth muscle, blood vessels and glands of the branchial arches and pharyngeal region (Nicol, 1950). In addition to providing autonomic innervation of the branchial region, the vagus of all groups of fish has cardiac and visceral rami innervating the heart and gastrointestinal tract, respectively. The paired cardiac rami of the vagus arise from ventral visceral rami, and, in elasmobranchs and at least some teleost species (e.g. Trichiurus), they also receive contributions from the most posterior branchial rami (Norris and Hughes, 1920; Young, 1933; Harrison, 1981). In both teleosts and elasmobranchs, the cardiac nerves run in the dorsal wall of the ducts of Cuvier to the sinus venosus, where they break up into a ganglionated plexus, extending only as far as the proximal regions of the atrium (Menidia: Herrick, 1899; Uranoscopus: Young, 1931; elasmobranchs: Young, 1933; Lampanyctus: Ray, 1950; Pleuronectes: Santer, 1972; Hydrolagus: Nicol, 1950; see also Nicol, 1952). These neurons provide a dense innervation of the sino-atrial region (Saetersdal, Justesen and Krohnstad, 1974; Laurent, Holmgren and Nilsson, 1983; Leknes and Saetersdal, 1980) and are likely to be responsible for cholinergic inhibition of cardiac activity (Axelsson, Ehrenstrom and Nilsson, 1987; see Morris and Nilsson, this volume, Chapter 6). Cardiac ganglion cells are reported to be absent from the South African lungfish, Lepidosiren (Jenkin, 1928), despite physiological evidence for a vagal inhibitory cholinergic innervation of the heart in this species (Morris and Nilsson, this volume Chapter 6). Small ganglia and individual ganglion cells are scattered along the dorsal visceral

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21

rami of the vagus running to the oesophagus and stomach of teleosts (Huber, 1900; Ray, 1950; P. Karila and S. Holmgren, personal communication), but probably not elasmobranchs (Norris and Hughes, 1920; Young, 1933; Nicol, 1950). In particular, a small “intestinal” or “swim-bladder” ganglion occurs around the junction of the vagal rami with the splanchnic nerves (Ray, 1950; Lundin and Holmgren, 1989; see also Pick, 1970). In lungfish, ganglion cells seem to be restricted to the pulmonary branches of the vagus (Jenkin, 1928; Nicol, 1952). As in mammals, it is likely that the vagus provides preganglionic inputs to neurons within the myenteric plexus of at least the foregut of all groups of fish (see Nilsson, 1983). Nothing is known of the vagal innervation of the pancreas of fish other than a tentative observation that some fine rami may run from the intestinal ramus of the vagus towards the pancreatic tissue (Ray, 1950). Amphibians, reptiles and birds From the information given in the preceding sections, it seems clear that there is a large jump in the level of organisation of the branchiomeric autonomic pathways between fish and mammals. In particular, mammals possess a set of well-defined ganglia that are absent in fish. This, in part, may be related to the presence of welldefined salivary glands in mammals, which in turn may be a consequence of major changes in the skeletal structure of the palate and jaws during the evolution of mammals. An examination of the arrangement of the branchiomeric autonomic ganglia in non-mammalian tetrapods may be expected to shed some light on this problem. Unfortunately, discussion is limited by the unavailability of much of the critical data, especially with respect to the distribution, connections and chemistry of ganglion cells within the cranial nerves of non-mammalian species. The palatine ramus o f the facial nerve. As mentioned above, the palatine nerve does not seem to contain any ganglion cells in any group of fish, and, therefore, may not carry autonomic pathways in these species. However, scattered ganglion cells and ganglia are associated with this nerve in all species of tetrapods examined to date. The lowest level of organisation seems to occur in anuran amphibians. In anurans, the palatine nerve runs beneath the floor of the orbit deep to the palatine membrane. It sends branches to the Harderian gland and to the intermaxillary gland in addition to the palatine mucosa itself (Stirling and Macdonald, 1883). There are numerous anastomoses with the ophthalmic and maxillary branches of the trigeminal nerve (Strong, 1895). In Rana and Bufo, small groups of ganglion cells are scattered throughout the palatine plexus, where they are usually found at junctions or branch points (Stirling and Macdonald, 1883; Smirnow, 1890; Strong, 1890, 1895; Huber, 1900). However, there are no constant, large, well-defined ganglia anywhere along the course of the palatine nerve or its ramifications (Stirling and Macdonald, 1883; Strong, 1895). These neurons are believed to innervate the glandular epithelium, small exocrine glands, and blood vessels of the palate and buccal cavity (Stirling and Macdonald, 1883; Strong, 1895), although there has been no modern confirmation of this conclusion. A similar distribution of ganglion cells probably occurs in the palatine plexus of urodele amphibians (Ambystoma:

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COMPARATIVE PHYSIOLOGY AND EVOLUTION

FIGURE 1.6 Diagram of the cranial autonomic ganglia of a lizard, Varanus. This is a dorsal view of the left side, with the position of the orbit shown by the light dotted line. Rostral is towards the top of the figure. Adapted from Willard (1915). Ill, oculomotor nerve; V, trigeminal nerve; V !, ophthalmic division of the trigeminal nerve; V2, maxillary division of the trigeminal nerve; VII, facial nerve; Vll?al, palatine ramus of the facial nerve; CiG, ciliary ganglion; EG, ethmoid ganglion; IOG, infraorbital ganglion; PG, palatine ganglion; POG, preorbital ganglion.

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Strong, 1895; Amphiuma: Norris, 1908). A small collection of ganglion cells occurs regularly at an anastomosis between the palatine nerve and the deep ophthalmic nerve, and may represent a rudimentary palatine ganglion (Coghill, 1902; Norris, 1908). In at least some species of caecilian amphibians, a somewhat greater degree of organisation of the palatine ganglion cells is observed. In Ichthyophis, Dermophis, and Herpele, but probably not in Geotrypes, a small ganglion, the “palatine ganglion” (see below), is found on a lateral branch of the palatine nerve communicating with a medial branch of the infra-orbital nerve (Norris and Hughes, 1918). Neurons in the palatine ganglion are most likely to innervate glands and blood vessels of the palate. There is no communicating ramus between the glossopharyngeal and facial nerves in caecilians (Norris and Hughes, 1918), so presumably all the preganglionic inputs to the palatine plexus in the species must arise from the facial nerve. It is not known if there are further isolated ganglion cells scattered along the more distal ramifications of the palatine nerve in caecilians. In reptiles, the palatine nerve enters the floor of the orbit between the ophthalmic and maxillary (infraorbital) branches of the trigeminal nerve and forms an extensive anastomosing plexus joining all three nerves (Figures 1.6, 1.7). A distinct “palatine ganglion” occurs in a position corresponding to that in caecilian amphibians, i.e. on or near a proximal connection between the palatine and infraorbital nerve, in lizards (Varanus: Watkinson, 1906; Bellairs, 1949; Anolis: Willard, 1915; Lacerta: Adams, 1942; Pogona, Ctenophorus, Tiligua, Phyllodactylus: I. Gibbins and M. Jurjevic, unpublished observations; Figure 1.6), snakes (Python: Gaupp, 1888; Elaphe, Thamnopis: Auen and Langebartel, 1977), chelonians (Trionyx: Ogushi, 1913; Chelydra, Eretmochelys: Soliman, 1964; Figure 1.7), and crocodiles (Crocodilus, Caiman, Alligator: Bellairs and Shute, 1953). A second prominent ganglion also occurs within the palatine plexus of most reptiles. The “ethmoid ganglion” occurs at an anastomosis between a branch of the palatine nerve and the ophthalmic nerve distally within the orbit of lizards (Watkinson, 1906; Bellairs, 1949; Gibbins and Jurjevic, unpublished observations; Figure 1.6) and snakes (Tropidonotus: Gaupp, 1888; Elaphe: Auen and Langebartel, 1977). In some cases, this ganglion appears actually to lie on the distal portion of the intra-orbital course of the ophthalmic nerve. The ethmoid ganglion is absent from crocodiles (Bellairs and Shute, 1953) and chelonians (Ogushi, 1913; Soliman, 1964; Figure 1.7). Lizards, snakes and turtles possess yet another ganglion within the palatine plexus: the “preorbital ganglion” is found in a distal anastomosis between the palatine nerve and the infraorbital nerve, just as the latter nerve leaves the orbit (Figures 1.5c, 1.6, 1.7). In lizards and snakes, it also has connections with the ethmoid ganglion (Willard, 1915; Bellairs, 1949; Soliman, 1964; Auen and Langebartel, 1977; I.L. Gibbins and M. Jurjevic, unpublished observations). In addition to these well formed, but generally small, ganglia of the palatine plexus, there are numerous microganglia and scattered nerve cell bodies along most of its ramifications (Willard, 1915; Auen and Langebartel, 1977; Figure 1.6). Moreover, there are numerous isolated ganglion cells in the terminal branches of the palatine nerve supplying the buccal mucosa and labial salivary glands of lizards. Most of the ganglion cells associated with the palatine nerve and its ramifications contain neuropeptides, such as VIP, NPY and galanin in various species-specific combinations (I.L. Gibbins

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COMPARATIVE PHYSIOLOGY AND EVOLUTION

Chelydra FIGURE 1.7 Diagram of the cranial autonomic ganglia of a turtle, Chelydra. Lateral view of the left side, with the orbit indicated by a light dotted line. Rostral is towards the left. Adapted from Soliman (1964). Ill, oculomotor nerve; V, trigeminal nerve; V !, ophthalmic division of the trigeminal nerve; V2, maxillary division of the trigeminal nerve; V3, mandibular division of the trigeminal nerve; VII, facial nerve; VII^, chorda tympani; VIIpal, palatine ramus of the facial nerve; IX, glossopharyngeal nerve; X, vagus nerve; CiG, ciliary ganglion; PG, palatine ganglion; POG, pre-orbital ganglion; SMG, submandibular ganglion.

and M. Jurjevic, unpublished observations). It is likely that they provide secretomotor and vasodilator innervation to the glands and blood vessels of the palate and upper jaw as well as to the Harderian gland of the orbit. Little is known of the corresponding ganglia in birds. They possess an ethmoid ganglion, and a palatine ganglion (also referred to as the sphenopalatine or pterygopalatine ganglion: see Bolton, 1971, and Bennett, 1974), similar to those in lizards and snakes (Gaupp, 1888; Bellairs and Shute, 1953). The ethmoid ganglion lies more superiorly within the orbit compared with the palatine ganglion. Both ganglia have connections with the Vidian nerve, which presumably transmits preganglionic fibres from the facial nerve. There are also interconnections between the two ganglia, primarily via the medial naso-palatine nerve (Bolton, 1971). As in lizards, there are scattered ganglion cells along the branches of the palatine nerve, as well as in the fine nerves supplying the Harderian gland itself (Gienc and Kuder, 1985). Neurons in both main ganglia provide secretomotor innervation to the Harderian gland and the supra-orbital salt gland (Ash, Pearce and Silver, 1969; Bolton, 1971; Gienc and Kuder, 1985). Whilst the nasal mucosa and glands are

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innervated mainly from the ethmoid ganglion, the palatine mucosa and glands probably are innervated mainly from the sphenopalatine ganglion. The hyomandibular branch o f the facial nerve, the chorda tympani, and the glossopharyngeal nerve. In anuran amphibians, there are numerous intralingual ganglion cells. These neurons contain VIP and galanin, and presumably provide vasodilator and secretomotor innervation to local blood vessels and glands (Baecker, Yanaihara and Forssmann, 1983; Morris et al., 1986a, 1989). The preganglionic pathways to these intralingual neurons are not known definitively. However, ganglion cells occur along the lingual branch of the glossopharyngeal nerve of Rana (Strong, 1895) and Bufo (Morris et al., 1986a). Examination of the innervation of the branchial and pharyngeal region of larval anurans and caecilians indicates that the lingual branch of the glossopharyngeal nerve develops from a post-trematic branchial ramus (Strong, 1895; Norris and Hughes, 1918). Furthermore, the branchial rami of the glossopharyngeal nerve contain small groups of ganglion cells, located near their entry to the floor of the pharynx of tadpoles (Strong, 1895). On the other hand, there does not seem to be a connection between the facial nerve and the lingual nerve in amphibians (Gaupp, 1888; Strong, 1895; Norris and Hughes, 1918). Taken together, these observations suggest that the intralingual neurons of amphibians lie on a glossopharyngeal pathway and that they are homologous with the pathways running in the branchial rami of the glossopharyngeal nerve in fish (see above). Intralingual neurons also occur in lizards (Baecker, Yanaihara and Forssmann, 1983). Moreover, small aggregations of ganglion cells are found near the salivary glands of the lower jaw (Figure 1.5d). Except in skinks (Tiliqua), most of these neurons contain VIP and galanin (Figure 1.5), as they do in anurans, and they probably provide vasodilator and secretomotor innervation to blood vessels and glands in the lower jaw (I.L. Gibbins and M. Jurjevic, unpublished observations). A chorda tympani, clearly homologous with that of mammals, exists in lizards, snakes and turtles (Auen and Langebartel, 1977; Lombard and Bolt, 1979; Figure 1.7). Within the lower jaw it connects with the main branch of the mandibular division of the trigeminal nerve, as it does in mammals, to form a “lingual” nerve (Gaupp, 1888; Ogushi, 1913; Willard, 1915; Soliman, 1964, Auen and Langebartel, 1977). At this point, a small ganglion, the “submandibular ganglion” has been described in crocodiles (Bellairs and Shute, 1953), turtles (Soliman, 1964; Figure 1.7) and snakes (Auen and Langebartel, 1977). In contrast, the glossopharyngeal nerve enters the tongue mainly via its connections with the hypoglossal nerve (cranial nerve XII) and has few if any anastomoses with the lingual nerve itself (Watkinson, 1906; Willard, 1915; Auen and Langebartel, 1977; Figure 1.7). These observations suggest that the facial nerve rather than the glossopharyngeal nerve may be the primary route of preganglionic fibres to the ganglion cells within the tongue of reptiles (see also Gaupp, 1888). Intralingual neurons have been reported in birds (Baecker, Yanaihara and Forssmann, 1983) and some are collected into a series of small submandibular ganglia (Bolton, 1971). These ganglia have primary connections with the facial nerve via the chorda tympani rather than with the glossopharyngeal nerve (Bolton, 1971;

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COMPARATIVE PHYSIOLOGY AND EVOLUTION

Bufo FIGURE 1.8 Distribution of cranial autonomic neurons along the various branches of the vagus nerve of toads, Bufo marinus. Different groups of neurons are distinguished by their neuropeptide content: SOM, somatostatin; VIP, vasoactive intestinal peptide. The somatostatin-containing neurons at the main bifurcation of the vagus project to the lung, whilst the VIP-containing cells project to the oesophagus and stomach. The somatostatin-containing neurons are cholinergic. See also Fig. 1.1. IX, glossopharyngeal nerve and roots; X, vagal roots. The combined sensory ganglion is finely stippled. Adapted from Gibbins et al.t (1987).

Bennett, 1974). As in reptiles, the available observations are consistent with the facial nerve rather than the glossopharyngeal nerve providing autonomic pathways to the lower jaw of birds. It will be important to test these conclusions with functional experiments.

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The vagus nerve. The distribution of ganglion cells along the various ramifications of the vagus nerve of anuran amphibians is well known (Rana: Strong, 1895; Bufo: Gibbins et al., 1987; Figures l.la,c, 1.8). About two-thirds along its course, the vagus divides into two main rami. One ramus branches further to give rise to a pulmonary ramus supplying the lung and pulmonary artery, a cardiac ramus supplying the heart and two rami supplying the proximal oesophagus. The other main branch of the vagus is known as the oesophago-gastric ramus and gives rise to two main rami which innervate the distal oesophagus, stomach, and probably the proximal portion of the duodenum. At the site of origin of the oesophago-gastric ramus, there is a large collection of ganglion cells (Figure 1.8). Immunohistochemical and pharmacological studies have shown that there are two functional populations of neurons in this location. The neurons in one population are cholinergic and contain both somatostatin and galanin: they project down the pulmonary ramus to provide excitatory innervation to the smooth muscle of the lung and the pulmonary artery (Gibbins et al., 1987; Morris et al., 1986a, 1989; Figure 1.1a). The neurons in the other population contain VIP and are probably non-cholinergic: they project down the oesophago-gastric ramus to provide inhibitory innervation to the foregut. More distally in both the pulmonary and oesophageal rami are large collections of ganglion cells, most of which contain VIP (Figure 1.1c), and which provide inhibitory innervation to the lung and gut, respectively. Within the lungs of anurans, the ganglion cells are found mainly in the proximal ramifications of the vagus (Smirnow, 1890; Gibbins et al., 1987). However, in urodeles, ganglion cells can be found in small branches of the vagus nerve for almost the full length of the lung (Stirling, 1882). Once it enters the heart, the cardiac ramus of the vagus runs over the sinus venosus and down the interatrial septum. Along the intracardiac course of the vagus are numerous cholinergic ganglion cells, which, in Bufo, contain both somatostatin and galanin, and inhibit cardiac activity (Campbell et al., 1982; Morris, Gibbins and Osborne, 1989; see Morris and Nilsson, chapter 6). These cells form loose aggregations (“Bidder’s ganglion”, “Remak’s ganglion”; Dogiel, 1877); however, in general, there is no regular arrangement of ganglion cells within the heart (Strong, 1895; Gibbins et al., 1987). In urodeles, the intracardiac ganglia contain prominent chromaffin cells, in addition to the cholinergic ganglion cells. Both classes of cells also contain galanin. The chromaffin cells are unusual in that they have long branching processes: they probably act as interneurons between the principal vagal postganglionic neurons (McMahan and Purves, 1976; Parsons et al., 1989; Figure 1.2c). Interneurons of this sort do not seem to be present in the cardiac ganglia of anuran amphibians. Comparatively little is known of the distribution of ganglia and the different functional autonomic pathways in the vagus nerves of reptiles and birds. In both groups, there is a vagal cholinergic inhibition of the heart, presumably mediated by intrinsic ganglion cells (see Morris and Nilsson, chapter 6) Such ganglion cells have been described along the branches of the vagus as they enter the heart as well as along their intra-cardiac ramifications (Caiman, turtles: Dogiel, 1877). In the rat snake, Elaphe, the intracardiac neurons also contain somatostatin (Donald, O’Shea and Lillywhite, 1990a). Ganglion cells occur in the intrapulmonary vagus of lizards and snakes, where at least some of them contain VIP (Acrochordus: Donald and

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Lillywhite, 1989; Elaphe: Donald, O’Shea and Lillywhite, 1990b; S. Holmgren and I.L. Gibbins, unpublished observations) and substance P (Davies and Donald, 1992). This distribution of vagal autonomic ganglion cells is similar to that seen in amphibians. Small ganglia occur in the pancreas of turkeys (Vaillant, Dimaline and Dockray, 1980): these neurons presumably lie in vagal pathways. EVOLUTION OF THE CRANIAL AUTONOMIC PATHWAYS Pathways to the upper jaw The palatine nerve carrying preganglionic fibres from the facial nerve to the ganglia of the orbit and palate in non-mammalian vertebrates clearly is homologous with the Vidian nerve (nerve of the pterygoid canal) and greater superficial petrosal nerve of mammals. Given the close anatomical relationship between the mammalian sphenopalatine ganglion and the maxillary division of the trigeminal nerve (i.e. the infraorbital nerve), it is most likely that the homologous ganglion in reptiles is the palatine ganglion, which has a similar close relationship with the infraorbital nerve. Other ganglia of the reptilian palatine plexus may persist in mammals as the various microganglia found within the retro-orbital plexus; along the Vidian nerve proximal to the sphenopalatine ganglion; and along the intracranial rami of the facial nerve (such as the greater superficial petrosal nerve). The neurons lying in the facial nerve contributions to the mammalian retro-orbital plexus, especially those with connections to the ciliary nerves, may correspond to the ethmoid ganglion cells of reptiles, since both these populations of neurons are more closely associated with the ophthalmic division of the trigeminal nerve rather than its maxillary division. With the evolution of a complete hard palate in mammals, the floor of the orbit, through which the original palatine nerves run, has become more clearly separated from the roof of the mouth (see Carroll, 1988). Consequently, the sphenopalatine ganglion may be considered as a consolidated relay point for the cranial autonomic regulation of secretory tissues and blood vessels of the upper jaw, taking over the functions of all the various ganglia of a primitive reptilian palatine plexus. A single palatine ganglion also occurs in crocodiles, generally considered to be the most advanced reptiles in terms of their physiology and behaviour (e.g. Meyer, 1984). Unlike other reptiles and birds, crocodiles possess a secondary palate, which has evolved independently from that in mammals. Perhaps, the evolution of a complete secondary palate in both crocodiles and mammals has generated the same adaptive pressures to concentrate their autonomic pathways via a single autonomic ganglion innervating the upper jaw. Alternatively, the presence of a single palatine ganglion in chelonians, crocodiles and mammals might imply that this is the primitive condition in stem reptiles, and that the ethmoid ganglion has developed independently in modern lepidosaurs and birds. Pathways to the lower jaw The cranial autonomic outflow to structures in the lower jaw seems to be derived primitively from the branchial rami of the facial and glossopharyngeal nerves, as is seen in elasmobranchs. In amphibians, it seems most of this function has been

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taken over by pathways following the glossopharyngeal nerve. In reptiles and birds, however, the main pathway apparently follows the facial nerve. The presumed reptilian condition is most similar to that seen in mammals, where the preganglionic cranial autonomic outflow to ganglia and microganglia innervating most targets in the lower jaw runs primarily with the facial nerve via the chorda tympani, with only a minor contribution from the glossopharyngeal nerve. In further support of this idea, the sublingual glands of reptiles are considered to be homologous with the sublingual and submandibular glands of mammals (Gaupp, 1888). The difference in autonomic pathways to the lower jaw of amphibians compared with amniotes may be related to the distinctive structure of the middle ear (Carroll, 1970) and the course of the chorda tympani in amphibians. Indeed, Lombard and Bolt (1979) have argued that the middle ear may have evolved independently in amphibians and amniotes. A long-standing problem still remains with respect to the evolutionary history of the otic ganglion. This ganglion, which lies on a glossopharyngeal pathway, apparently occurs only in mammals, and there is no obvious homologue in nonmammalian vertebrates (Gaupp, 1888). As described above, its main target is the parotid salivary gland, in addition to some blood vessels in the lower jaw and the cerebral circulation. The otic ganglion usually has close anatomical relations with the origin of the maxillary (third) division of the trigeminal nerve, which itself mostly innervates targets in the lower jaw. Therefore, it seems reasonable to assume that the autonomic pathway now including the otic ganglion also originally innervated primarily lower jaw structures. With the rostral shift of the jaw articulation that has occurred uniquely in the evolution of mammals from reptiles (see Kermack and Kermack, 1984; Carroll, 1988), it is possible that the more proximal components of a glossopharyngeal autonomic pathway to the lower jaw became associated with the area of the new jaw articulation. Presumably, this change occurred in parallel with the development of a new salivary gland, the parotid gland, perhaps derived from glandular tissue primitively located more proximally within the lower jaw. Remnants of this primitive pathway may persist in the relatively minor glossopharyngeal input to the submandibular gland ganglia, for example. These conclusions suggest that the lower jaw of mammal-like reptiles, in contrast with modern reptiles, possessed autonomic pathways running in both the facial and glossopharyngeal nerves. Vagal pathways A comparison of the distributions of the various rami of the vagus nerve and the ganglion cells within them suggests that there are some constant features across the vertebrate classes. Most notable is the presence of a cardiac ramus with intracardiac ganglion cells, and a number of rami running to the foregut (oesophagus, stomach, and, in some cases, the proximal small intestine), which carry preganglionic fibres to intrinsic enteric neurons. Of considerable interest is the homology of the vagal innervation of the lung of tetrapods with the various branches of the vagus in fish. The lung and its associated structures develop as a ventral evagination of the foregut at the levels of the fourth to sixth branchial arches (Balinsky, 1970; Hamilton, Boyd and Mossman, 1976; Moore, 1982). Because of this embryological origin from the

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gut, the intrinsic ganglionated plexuses of the lung are sometimes considered to be homologous with the enteric plexuses. However, a careful comparison of the innervation of the heart, lungs and foregut in anuran amphibians with the vagal innervation of the branchial arches and viscera of fish suggests that this is almost certainly not the case. Rather, the ventral origin of the lungs from the branchial arch region of the pharynx suggests that the ganglia in the lung are most likely to be homologous with the small ganglia in the branchial and pharyngeal rami of the vagus in fish. Two further observations support this conclusion. First, in teleosts, the cardiac ramus of the vagus originates from the main ventral visceral ramus of the vagus, which also gives rise to an inferior pharyngeal ramus running to the floor of the pharynx and the oesophagus (Herrick, 1899; Ray, 1950; Harrison, 1981; Figure 1.11). A similar branching pattern is seen in the cardiac, pulmonary, cranial oesophageal, and middle oesophageal rami of the anuran vagus (Figure 1.8), and is consistent with the pulmonary ramus being an anuran homologue of the teleost inferior pharyngeal ramus. Second, the ganglion cells themselves in the pulmonary rami of the vagus in anurans are monopolar and are morphologically identical to other cranial postganglionic neurons, including the vagal postganglionic neurons within the heart; they are quite unlike the multipolar enteric neurons intrinsic to the gut (see Figure 1.1). The ganglia in the cranial and middle oesophageal rami of the anuran vagus as well as those at the origin of the oesophago-gastric ramus which project to the gut, are also likely to be homologous with the branchial neurons of fish. Like the pulmonary rami, these rami to the proximal oesophagus arise from the ventral visceral branch of the vagus and contain monopolar ganglion cells (Figures 1.1c, 1.8). On the other hand, the oesophago-gastric ramus in anurans probably is equivalent to the dorsal visceral ramus of the vagus in fish (Herrick, 1899; Ray, 1950; cf. Figures 1.8 and 1.11), and may represent the primitive route by which vagal preganglionic fibres reach the stomach. Nevertheless, the ganglion cells situated distally along the oesophago-gastric ramus are monopolar, like those in the other vagal rami, and bear no morphological resemblance to the multipolar intrinsic enteric neurons (Figure 1.1). Consequently, all the ganglion cells along the oesophageal and oesophago-gastric rami may be considered better as genuine postganglionic vagal neurons, rather than as displaced enteric neurons. The distribution of ganglion cells in the vagal pathways to the foregut in mammals is somewhat different from that in anurans. Whilst scattered ganglion cells are relatively common in the pulmonary rami of the vagus in mammals, they are rare along the rami supplying the digestive tract (Botar et al., 1950). Since the vagus is well known to provide preganglionic inputs to intrinsic neurons of the mammalian foregut, at least some of which contain and release VIP (Fahrenkrug et al., 1978; Furness and Costa, 1987), it seems that the vagal postganglionic neurons of mammals have become fully incorporated into the enteric plexuses of the foregut. Nevertheless, there is still the possibility that vagal postganglionic neurons innervating the oesophagus of mammals are more akin to the primitive branchial neurons than the enteric neurons.

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SPINAL AUTONOM IC PATHW AYS: SYMPATHETIC A N D PELVIC GANGLIA The functional neuroanatomy of the spinal autonomic pathways is best known in mammals, especially guinea-pigs, cats and humans. Details can be found in Hirt (1934), Kuntz (1945), Mitchell (1953), Pick (1970), Gabella (1976) and Gibbins (1990a), and this information will be summarised briefly here, before more closely examining the corresponding pathways in non-mammalian vertebrates. MAMMALS Although sympathetic preganglionic outflows are restricted to thoraco-lumbar levels of the spinal cord, sympathetic ganglia themselves extend from upper cervical to sacral levels. The sympathetic ganglia can be separated into two main groups on the basis of their location, and, at least to some degree, their peripheral projections and functions. They are usually called the “paravertebral” and the “prevertebral” ganglia. In addition to the principal ganglion cells, most sympathetic and pelvic ganglia contain small aggregations of chromaffin cells. However, the largest collection of chromaffin tissue in adult mammals occurs in the adrenal medulla. These catecholamine secreting cells are innervated by lumbar preganglionic neurons, and consequently are considered to form part of the spinal sympathetic outflow. Paravertebral ganglia The paravertebral sympathetic ganglia lie beside the vertebral column, forming two largely independent “sympathetic chains”. From thoracic to sacral levels, the paravertebral ganglia are more or less segmentally arranged. Adjacent paravertebral ganglia are connected with each other by interganglionic connectives, thereby forming the characteristic chain of ganglia. The largest ganglion is the stellate ganglion. It is located at the lower cervical-upper thoracic level of the chain, and connects with the brachial plexus and cervical sympathetic trunk, as well as giving rise to the cardiac sympathetic nerves. Cervical ganglia are also present. In marsupials and eutherians, the most prominent is the superior cervical ganglion. It has close anatomical relations with the bifurcation of the common carotid artery and with the vagus nerve, and provides sympathetic innervation to cranial tissues. However, in monotremes, the main cervical ganglion is found more caudally, around the level of the sixth cervical spinal nerve (van den Broek, 1908). Preganglionic fibres reach the postganglionic neurons within the ganglia via the ventral spinal roots and communicating rami. In many cases, the preganglionic fibres may travel up or down the sympathetic chain before making synapses with ganglion cells. This clearly must happen for the ganglia at cervical and sacral levels for which there are no direct spinal outflows. The axons of postganglionic neurons may leave the ganglia directly or they also may project along the chain for a number of segments before they leave. In either case, postganglionic fibres may then enter the spinal nerves via communicating rami to be distributed to the rest of the body,

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or they may project to the viscera by well defined nerve trunks, such as the carotid nerves in the head, the cardiac nerves in the thorax, or the splanchnic nerves in the abdomen. Small collections of ganglion cells may lie in the rami communicating with the spinal nerves and along the carotid, cardiac and splanchnic nerves (see Pick, 1970; Gabella, 1976; Gibbins, 1990a, for references and further discussion). Most of the postganglionic neurons in the paravertebral ganglia of laboratory mammals (e.g. guinea-pigs, rats, mice, cats, dogs, pigs) are noradrenergic. They innervate a variety of peripheral targets including blood vessels, pilo-erector muscles, adipose tissue, and exocrine glands, such as the salivary glands. They also innervate airway smooth muscle, the heart and, to some degree, the abdominal and pelvic viscera. Non-noradrenergic sympathetic neurons, at least some of which are cholinergic, are found in many species, although they seem to be absent from marsupials (Morris, Gibbins and Murphy, 1986b). They provide vasodilator innervation to blood vessels supplying skeletal muscle and perhaps bones, and secretomotor innervation to cutaneous sweat glands. In species such as humans, where sweat glands are widely and densely distributed over the body, cholinergic sudomotor neurons may comprise up to 30% of postganglionic neurons in the paravertebral ganglia (Gibbins, 1990a). Recent immunohistochemical studies have demonstrated not only that each of the main populations of postganglionic sympathetic neurons is neurochemically distinct, but also that they are innervated by unique populations of preganglionic neurons (Gibbins, 1992). Prevertebral ganglia The prevertebral sympathetic ganglia are associated with the main ventral branches of the aorta. Thus, there is a coeliac ganglion at the origin of the coeliac artery, a superior mesenteric ganglion at the origin of the superior mesenteric artery, and an inferior mesenteric ganglion at the origin of the inferior mesenteric artery and a renal (or aortico-renal) ganglion at the origin of each renal artery. In some species, such as guinea-pigs, the coeliac and superior mesenteric ganglia are virtually fused into a single complex lying around the origin of the common coeliac-superior mesenteric trunk (Macrae, Furness and Costa, 1986; Lindh, Hokfelt and Elfvin, 1988). On the other hand, the inferior mesenteric ganglion in humans is very small, in keeping with the relatively small size and peripheral distribution of the inferior mesenteric artery (Gibbins, 1990a). Although the prevertebral ganglia are often regarded as being single midline ganglia, they often possess two lobes, one lying either side of their related artery, suggesting that they are primarily paired structures. This can be seen clearly in the bilateral renal ganglia, for example. The main ganglia are connected with each other via a loosely organised plexus known as the pre-aortic or intermesenteric plexus. It continues caudad from the inferior mesenteric ganglion towards the pelvic plexus as the hypogastric or presacral plexus. Within the pre-aortic plexus are irregular microganglia and small collections of chromaffin cells. There are usually thought to be no prevertebral ganglia in the thorax. However, in most mammalian species examined, there is a prominent mediastinal ganglion (the “ganglion of Wrisberg”), which lies near the arch of the aorta (Teitelbaum and

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Uhlenhuth, 1932), and which may be regarded as a thoracic prevertebral ganglion. It is closely associated with the cardiac nerves running from the stellate ganglion and it also may have connections with the vagus nerve. Most, but not all, of the postganglionic neurons in the prevertebral ganglia are noradrenergic and they mainly innervate the abdominal and pelvic viscera. They receive multiple preganglionic inputs from thoraco-lumbar preganglionic neurons via the splanchnic nerves that run through the mesentery from the sympathetic chain. However, some cells in the inferior mesenteric ganglion have preganglionic input from the sacral spinal levels via the pelvic nerves. Such neurons would be functionally “parasympathetic”, according to conventional criteria based on Langley’s classification. Furthermore, many of the neurons also receive peripheral synaptic inputs from enteric neurons with cell bodies in the myenteric plexus of the intestine (Kuntz, 1940; Crowcroft and Szurszewski, 1971; Dalsgaard and Elfvin, 1982; Dalsgaard et al., 1983; Macrae, Furness and Costa, 1986; Lindh, Hokfelt and Elfvin, 1988; Masuko and Chiba, 1988; Webber and Heym, 1988). Thus, some of the prevertebral neurons, such as those controlling secretion from the small intestine, are integral components of peripheral reflex arcs (see Furness and Costa, 1987). Once again, many of the discrete populations of neurons in the prevertebral ganglia can be identified by their distinctive content of neuropeptides (Costa and Furness, 1984). Pelvic ganglia As the pre-aortic plexus extends caudad from the inferior mesenteric ganglion, it is often known as the hypogastric plexus, which in turn runs into the pelvic plexus. The major ganglion in this region is known as the hypogastric or anterior pelvic ganglion in males and the paracervical ganglion in females. In both sexes, these ganglia are associated with a series of smaller ganglia, and contain noradrenergic and non-noradrenergic neurons. Functional studies have shown that some neurons in these ganglia receive preganglionic inputs from lumbar levels of the spinal cord, and, therefore would be considered to lie within “sympathetic” pathways. However, other neurons in the same ganglia have preganglionic inputs from sacral levels of the spinal cord and would therefore be regarded as lying in “parasympathetic” pathways. Furthermore, some individual neurons probably have inputs from both levels of the spinal cord (Szurszewski, 1981), so that they cannot be classified unambiguously as lying in either a sympathetic or a parasympathetic pathway. Presumably, these neurons could be activated by either pathway, depending on the physiological circumstances. Within the more caudal parts of the pelvic plexus, most of the neurons are nonnoradrenergic and most of their preganglionic inputs arise from sacral levels via the pelvic nerves. In most species, the caudal pelvic neurons are aggregated into small ganglia near and within the walls of the distal colon and rectum, the urinary bladder and ureters, and, in males, at the base of the penis. However, in rats, they are aggregated into a single “major pelvic ganglion”. The neurons in the caudal pelvic ganglia provide innervation to the smooth muscle, glands and blood vessels of the pelvic organs. Throughout the pelvic plexus, often closely associated with the ganglia, are prominent clusters of chromaffin cells.

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Components o f spinal autonomic pathways: a continuum? When the various neuronal populations within the different ganglia on the spinal autonomic pathways of mammals are compared using functional, anatomical, and neurochemical criteria, it is evident that the prevertebral and pelvic ganglia form a graded continuum. Thus, using Langley’s criteria, the coeliac-superior mesenteric ganglion complex can be seen as being mainly “sympathetic”, in that most of the postganglionic neurons are noradrenergic and the preganglionic inputs arise almost entirely from thoraco-lumbar levels. Conversely, the caudal pelvic ganglia can be considered mainly “parasympathetic”, in that most of the postganglionic neurons are non-nor adrenergic, and preganglionic inputs arise predominantly from the sacral spinal cord. However, the inferior mesenteric ganglia and the anterior pelvic ganglia contain mixtures of neurons lying in both the sympathetic and parasympathetic pathways, with the inferior mesenteric ganglion containing relatively more “sympathetic” neurons and the anterior pelvic ganglion containing relatively more “parasympathetic” neurons. Consequently, the ganglia on the spinal autonomic pathways should be designated simply as “paravertebral”, “prevertebral”, or “pelvic” ganglia (as proposed by Kuntz, 1911b, for example) according to their locations, without any implications as to the origin of preganglionic inputs, neurochemistry, or functions of their constituent neurons. This problem will be addressed further below, after a consideration of the arrangements in non-mammalian vertebrates. FISH There are clear differences in the level of organisation of the ganglia associated with the spinal autonomic outflow in the different classes of fish, ranging from loosely organised clusters of chromaffin tissue in cyclostomes to the well organised sympathetic chains in teleosts. Cyclostomes The literature on the spinal autonomic pathways in cyclostomes is sparse and confusing. This must be due, at least in part, to the ill-defined nature of tissues that could be considered to be components of these pathways (see Nicol, 1952, Campbell, 1970, Pick, 1970, and Nilsson, 1983, for further discussion). It is clear that cyclostomes do not possess sympathetic ganglia like those seen in mammals, and it is most probable that they do not have autonomic ganglion cells closely associated with the roots of their spinal nerves (Nicol, 1952; Pick, 1970). In lampreys, small collections of cells, presumed to be postganglionic neurons, are clustered around the cloaca, and distal portions of the hindgut and ureters. Occasional cells, which may be autonomic neurons, are also found near the kidneys (Lampetra: Johnels, 1956; Johnels and Ôstlund, 1958; Honma, 1970; Nakao and Ishizawa, 1982). Some of these neurons contain serotonin (5-hydroxytryptamine; Honma, 1970; Baumgarten et al., 1973). Ultrastructural studies have demonstrated the neurons do receive synaptic inputs (Nakao and Ishizawa, 1982), that presumably originate from the more caudal spinal nerves (Johnels and Ôstlund, 1958).

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Although there is no obvious widespread system of spinal autonomic neurons, there are, nevertheless, well defined aggregations of chromaffin cells throughout the body of lampreys (Petromyzon: Gaskell, 1912; Lampetra: Johnels, 1956). The largest collection of chromaffin cells is located in the walls of the anterior part of the posterior cardinal veins, and is particularly marked in the vicinity of the Cuverian duct and the origin of the coeliac artery. Smaller irregular collections of chromaffin cells occur in the walls of the segmental veins, especially near the dorsal root ganglia. In the more caudal regions of the body, chromaffin tissue lies on the dorsal surface of the single cardinal vein, between it and the caudal artery. The chromaffin cells in all these locations closely resemble those lining the walls of the heart (see under “Cranial Autonomic Outflows”, earlier). Some of the chromaffin cells are multipolar, and probably have been described as peripheral neurons by early authors (e.g. Kuntz, 1911b; see Nicol, 1952). It is by no means clear how, or indeed if, these chromaffin cells are connected to the central nervous system. According to Johnels (1956), they are innervated by fibres running from the dorsal rather than ventral roots, which would be consistent with GaskelFs observations on the close association between the segmentally arranged chromaffin tissue and the dorsal root ganglia. Although these fibres have been considered to be autonomic, there is no functional evidence that this is really the case. The status of a spinal autonomic outflow in hagfish (Myxine) is even more difficult to interpret than it is in lampreys. Cells considered to be autonomic neurons have been described along the dorsal aorta (see Nicol, 1952, and Nilsson, 1983). In addition, there is a subcutaneous nervous plexus, formed by terminations of the spinal nerves, which is particularly dense around the slime glands. Within the subcutaneous plexus, there are numerous bipolar or unipolar cells, which can be stained with either silver or methylene blue methods. Consequently, these cells have been interpreted as being peripheral autonomic neurons lying in spinal pathways, that presumably provide motor innervation to smooth muscle surrounding the glands and the subcutaneous venous sinuses (Bone, 1963). It is not clear if similar cells are present in lampreys (Campbell, 1970) Elasmobranchs Sympathetic ganglia. Although cyclostomes lack sympathetic ganglia and their spinal autonomic outflow is poorly defined, sympathetic ganglia and their corresponding spinal pathways clearly are present in elasmobranchs. Despite this, however, they still possess some of the characteristics of the cyclostome spinal autonomic system, the most notable being more or less segmentally arranged aggregations of chromaffin tissue (Figure 1.9). The following account is based upon the remarkably concordant descriptions of Chevrel (1889: Scyllium, Mustelus, Galeus, Acanthias, Centrina, Squatina, Raja, Myliobates, Trygon, Torpedo, Chimaera), Muller and Liljestrand (1918: Squalus, Raja), Young (1933: Scyllium, Mustelus, Torpedo), Nicol (1950: Hydrolagus) and Gannon et al., (1972: Heterodontus). Sympathetic ganglia extend caudad from the region of the subclavian artery and brachial plexus supplying the pectoral fin, to about the level of the anus; they are

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FIGURE 1.9 Diagram of the spinal autonomic ganglia in the trunk region of a shark, Scylliorhinus. Chromaffin bodies are shown in coarse stipple, whilst sympathetic ganglia are solid black. The largest chromaffin bodies are known as the “axillary bodies”, and their accompanying sympathetic ganglia are the “gastric ganglia”. Note the irregular nature of the connectives between the ganglia, and the communicating rami. Adapted from Young (1933). ca, coeliac artery, accompanied by splanchnic nerves arising from the gastric ganglia; sa, segmental artery; sn, spinal nerve.

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FIGURE 1.10 Transverse section through the trunk of a shark (Scylliorhinus) at the level of the kidneys. Note the close association of the sympathetic ganglia (solid black) with the suprarenal chromaffin bodies (sup), the segmental arteries (sa), and the segmental veins (sv). cv, posterior cardinal vein; da, dorsal aorta; ir, interrenal tissue. Adapted from Young (1933).

absent from cranial and caudal regions (see also Shore, 1889). In general, the ganglia are segmentally arranged, although in some segments, there may be more than one ganglion (Figure 1.9). Adjacent ganglia usually are interconnected by a loose network of fine nerve trunks, but there is not a well demarcated sympathetic chain as seen in mammals, and some ganglia may lack direct connections with their neighbours. On the other hand, there may be connections (transverse commissures) between ganglia on each side of the animal. The more cranial ganglia lie on the dorsal walls of the posterior cardinal sinuses at levels corresponding to the nearby segmental arteries. The more caudal ganglia are located more ventrally and are sandwiched between the ventral walls of the posterior cardinal veins and the dorsal surface of the mesonephric kidneys (Figure 1.10). Except in Heterodontus (Gannon, Campbell and Satchell, 1972), the most cranial sympathetic ganglion is significantly larger than the others, and has been named the “gastric ganglion”, since it is the origin of the largest splanchnic nerve, following the coeliac artery to the stomach (Figure 1.9). In the region of the brachial plexus of sharks (but not rays: Chevrel, 1889; Young, 1933), there is a “post-branchial plexus” (Chevrel, 1889), containing smaller “pregastric ganglia” (Nicol, 1950), especially at regions where the plexus is joined by visceral rami of the vagus nerve (Chevrel, 1889). Nevertheless, there are no prevertebral ganglia associated specifically with the coeliac and mesenteric arteries of elasmobranchs. Although there are some ganglion cells scattered around the urino-genital tract, there are no well-defined pelvic ganglia. According to Young (1933) and Nicol (1950), the sympathetic ganglia probably receive preganglionic inputs from the spinal cord via communicating rami with the corresponding segmental spinal nerve. However, postganglionic fibres are supposed not to travel with the spinal nerves. Rather, they follow segmental blood vessels

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directly or run out in the splanchnic nerves to the gastro-intestinal tract and perhaps other viscera (for details, see Nilsson and Holmgren, 1988). In the absence of any prevertebral ganglia, the splanchnic nerves of elasmobranchs contain only postganglionic fibres. Chromaffin tissue. The most characteristic feature of the elasmobranch sympathetic ganglia is their close association with prominent collections of chromaffin tissue, sometimes known as supra-renal bodies (Figure 1.9). The largest collections of chromaffin tissue are aggregated around the gastric ganglia, and the combined gastric ganglion-chromaffin cell mass is known as the “axillary body”. Most ganglia have large numbers of chromaffin cells associated with them, and, indeed, may be completely surrounded by chromaffin tissue, especially in the region of the kidney (Figure 1.10). In Heterodontus, one of the more primitive living sharks (Carroll, 1988), the chromaffin tissue extends along the interganglionic connectives, thereby forming a continuous cord on the dorsal surface of the posterior cardinal vein from the axillary body to the cranial end of the kidney (Gannon, Campbell and Satchell, 1972). Histological and physiological studies together indicate that the chromaffin tissue of elasmobranchs is innervated by preganglionic spinal neurons (Nilsson and Holmgren, 1988). This chromaffin tissue releases catecholamines, predominantly adrenaline, into the circulation in a way analogous with the adrenal medulla of mammals. The most likely targets of these circulating catecholamines are the heart, which also contains chromaffin tissue but which generally lacks a direct sympathetic innervation, and perhaps the branchial vasculature (Nilsson and Holmgren, 1988). Teleosts The sympathetic ganglia of teleost fish are generally arranged into well organised paravertebral sympathetic chains, with adjacent ganglia being connected by a well defined sympathetic trunk. Ganglia are ordered segmentally and extend from cranial to caudal levels. The following descriptions are taken mainly from the following sources: Chevrel (1889: Labrax = Anarhichas, and nearly 70 other species from more than 30 families); Young (1931: Uranoscopus)\ Ray (1950; Lampanyctus); Soust (1981: Platycephalus)\ Nilsson (1983: Gadus). Cranial sympathetic ganglia. Well developed cranial sympathetic ganglia are unique to teleosts and holosteans. In chondrosteans, they are rudimentary or absent (Chevrel, 1894; Allis, 1920-21; Norris, 1925). In most teleost species, there are usually five pairs of cranial sympathetic ganglia associated with the cranial nerves (Chevrel, 1889; Figure 1.11). The first two ganglia are associated with the trigeminal-facial sensory ganglion complex. One ganglion is primarily associated with the trigeminal component, whilst the other is associated with the facial nerve component. The third ganglion lies ventral to the sensory ganglion of the glossopharyngeal nerve, whilst the fourth and fifth ganglia lie either end of the vagal sensory ganglion complex. In Menidia, a relatively primitive teleost, there is a more rostral small group of ganglion cells lying on the cranial margin of the trigeminal ganglion near the roots of the ciliary nerves and probably connecting with the

FIGURE 1.11 Simplified diagram of the cranial nerves (stippled) and cranial sympathetic ganglia (solid black) of a teleost fish, Menidia. A sympathetic ganglion is associated with each of the major cranial sensory ganglia (dense stipple). The ciliary ganglion (CiG) is not sympathetic, but lies in the oculomotor cranial autonomic outflow (III). Nevertheless, it has connections with the cranial sympathetic trunk. The position of the orbit is shown by a light dotted line. Adapted from Herrick (1899). Ill, oculomotor nerve; V, trigeminal roots and ganglia; V^VII, ophthalmic division of the trigeminal nerve and accompanying facial nerve fibres; V2-VII, maxillary division of the trigeminal nerve and accompanying facial nerve fibres; V3-VII, mandibular division of the trigeminal nerve and accompanying facial nerve fibres; VII, facial nerve roots and ganglia; VIIhyo, hyomandibular ramus of the facial nerve; VIIpal, palatine ramus of the facial nerve; IX, glossopharyngeal nerve; X t , X2, X3, main branchial rami of the vagus; X4oes, oesophageal ramus of the fourth branchial ramus of the vagus; Xcar(i, cardiac ramus of the vagus; Xint, intestinal ramus of the vagus; CiG, ciliary ganglion; CG, coeliac ganglion.

COMPARATIVE ANATOMY AND EVOLUTION 39

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palatine plexus (Herrick 1899; Figure 1.11). In some teleost species, the cranial sympathetic ganglia fuse to varying degrees. For example, in Gadus (Herrick, 1900; Nilsson, 1983) and Platycephalus (Soust, 1981), there are only three pairs of relatively large ganglia associated respectively with the ganglia of the facial-trigeminal complex, the glossopharyngeal nerve, and vagal complex. However, in Trichuris (Harrison, 1981) and Lampanyctus (Ray, 1950), the sympathetic ganglia associated with the glossopharyngeal nerve, and possibly also the vagus nerve, are absent, so that there is only a large sympathetic ganglion complex associated with the trigeminal ganglion. Conversely, in eels, a single pair of cranial sympathetic ganglia is associated with the vagal ganglion, whilst the more rostral ganglia are absent {Conger: Chevrel, 1889; Figure 1.12). The great majority of the neurons in the cranial sympathetic ganglia synthesise catecholamines (Soust, 1981; Nilsson, 1983). Postganglionic sympathetic fibres run out with their corresponding cranial nerves to supply blood vessels, pigment cells, and perhaps glands in the cranial and branchial region (Chevrel, 1889). Neurons in the vagal sympathetic ganglia project only down the post-trematic and cardiac rami, with a small contribution to the gastric rami (Grove and Campbell, 1979; Soust, 1981; Nilsson, 1983). Preganglionic neurons projecting to the cranial sympathetic ganglia leave the spinal cord mostly from the third and fourth spinal nerves (Uranoscopus: Young, 1931; Platycephalus: Soust, 1981; Gadus: Nilsson, 1983); there is no preganglionic input from the cranial nerves themselves, and they cannot be considered as homologues of the cranial parasympathetic ganglia of amniotes. Spinal sympathetic ganglia. In the trunk, the more cranial part of the spinal sympathetic chains lie either side of the vertebral column. However, in Uranoscopus, Platycephalus and Serranus, they fuse in the region of the kidneys where the posterior cardinal veins also fuse, and run as a single sympathetic trunk, ventral to the vertebral column, towards the end of the abdominal cavity (Chevrel, 1889; Young, 1931; Soust, 1981). This fusion does not occur in Anarhichas (Chevrel, 1889) or Lampanyctus (Ray, 1950), but in eels, there is only a single sympathetic trunk for most of the length of the abdominal cavity {Conger: Chevrel, 1889; Figure 1.12). In addition to this fusion between the sympathetic chains, there are prominent commissures from one side to the other (Young, 1931; Ray, 1950; Soust, 1981). In many species, there is an especially large “transverse commissure” at about the level of the second or third spinal sympathetic ganglion (Young, 1931; Soust, 1981; Nilsson, 1983). As in the cranial sympathetic ganglia, the postganglionic neurons in the spinal sympathetic ganglia do not project up or down the sympathetic trunk, or, indeed, along the commissures. Rather, they directly enter their corresponding spinal nerve via communicating rami to innervate blood vessels, pigment cells and perhaps cutaneous glands. Nevertheless, preganglionic fibres project for considerable distances in both directions along the chain, as well as from one side to the other via the commissures (Young, 1931; Soust, 1981). Although there are ganglia associated with the first two spinal segments and the brachial plexus, their preganglionic inputs probably leave the spinal cord at the third and fourth spinal segments (Young, 1931;

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Conger FIGURE 1.12 Ventral view of the unusual arrangement of the sympathetic ganglia in the eel, Conger. Caudal to the brachial sympathetic plexus, there is only a single sympathetic chain, lying on the left side of the vertebral column, whilst the aorta lies on the right side. The cranial sympathetic ganglia are compressed into one large ganglion (CrSGi) at the level of the glossopharyngeal and vagus nerves, and a smaller one at the level of the axillary artery (CrSG2). The coeliac ganglion (CG) lies at the same level as the sympathetic chain. Adapted from Chevrel (1889).

Soust, 1981). In Platycephalus, preganglionic outflow from the third and fourth spinal segments run craniad, whilst the outflow from the fifth and sixth segments runs caudad along the sympathetic chain (Soust, 1981). Prevertebral ganglia. Many teleost species possess some form of a coeliac ganglion (see Chevrel, 1889; Figure 1.11). The coeliac ganglion is absent from the primitive

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chondrostean fish, as is the case in elasmobranchs (Acipenser: Chevrel, 1894). Nevertheless, scattered ganglion cells occur along their splanchnic nerves {Huso: Balashov et al,, 1981). A range of intermediate states apparently occurs in teleosts. For example, a primitive arrangement, similar to that in chondrosteans, also exists in Salmo, which generally lacks a well formed ganglion (Campbell and Gannon, 1976). In Gadus, the coeliac ganglion seems to be comprised of the fused second and third spinal sympathetic ganglia on the right side (Nilsson, 1983), whilst in Uranoscopus and the stomachless flatfish, Rhombosolea and Ammotretis, the coeliac ganglion is connected by short splanchnic nerves to the first and second spinal sympathetic ganglia of the right side only (Young, 1931; Grove and Campbell, 1979). In Gadus and Uranoscopus, the transverse commissure also transmits preganglionic fibres to the coeliac ganglion (Young, 1931; Nilsson, 1983), but in Platycephalus and Anarhichasy there is a separate splanchnic nerve supplying preganglionic fibres to this ganglion (Chevrel, 1889; Soust, 1981). Prevertebral ganglion cells, regardless of their precise location, innervate the gastrointestinal tract via the mesenteric nerves. In most species, there are connections between visceral branches of the vagus and the splanchnic nerves. Small ganglia may appear at these junctions. Cholinergic sympathetic neurons. Most of the postganglionic neurons in the sympathetic chains, coeliac ganglion, or scattered along the splanchnic nerves contain catecholamines {Tinea: Baumgarten, 1967; Salmo: Campbell and Gannon, 1976; Gadus: Nilsson, 1983, 1984b; Channa: Ishimatsu, Johansen and Nilsson, 1986). Their main targets are the vasculature, including the renal circulation, visceral smooth muscle (Nilsson, 1984a) and cutaneous chromatophores (Sand, 1935). However, there is pharmacological evidence that the pigment cells in the skin of catfish (Parasilus) are innervated by cholinergic sympathetic neurons, presumably arising from the chain ganglia. Such a prominent cholinergic sympathetic pathway seems to be unique amongst those teleosts studied to date (Fujii and Miyashita, 1976). Small numbers of functionally cholinergic sympathetic neurons occur in the coeliac ganglion of Gadus (Winberg, Holmgren and Nilsson, 1981), Rhombosolea, and Ammotretis (Grove and Campbell, 1979). Pelvic ganglia. In Anarhichas (Chevrel, 1889), Uranoscopus (Young, 1931), Lampanyctus (Ray, 1950), and Gadus (Nilsson, 1983), collections of ganglion cells associated with the ureters and urinary bladder have been described. It is not clear whether these neurons are related to the sympathetic chain (like the coeliac ganglion), or whether they mark the rudiments of a pelvic plexus. The presence of some non-adrenergic neurons in this ganglion, at least some of which innervate the smooth muscle of the urinary bladder (Nilsson, 1983), makes the latter possibility more attractive. Chromaffin tissue. Whilst there are some chromaffin cells in the sympathetic ganglia, the main collection of chromaffin tissue surrounds the posterior cardinal veins at the cranial ends of the kidneys, forming the so-called “head kidney”. In GaduSy there is a small sympathetic ganglion (“satellite ganglion”) associated with

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this tissue. Despite this, the chromaffin tissue is innervated by spinal preganglionic neurons (see Nilsson, 1983, 1984b). There is a similar distribution of chromaffin tissue in ganoid fish, with the major aggregations being around the posterior cardinal veins (Holosteans: Lepisosteus, Nilsson, 1981; Chondrosteans: Acipenser, Chevrel, 1894; Huso, Balashov et al., 1981). Dipnoans As in teleosts, the sympathetic ganglia of lungfish are arranged into well defined paravertebral sympathetic chains. However, unlike teleosts, but like tetrapods, there are no cranial sympathetic ganglia. Furthermore, there may not be any connections between the sympathetic trunk and the vagus, or any of the other cranial nerves. An unusual feature of the dipnoan sympathetic chain is that it tends to form loops around the segmental arteries, with a small ganglion lying both on the cranial and caudal ends of the loop. The ganglia are so small that they may not be visible macroscopically (Lepidosiren: Jenkin, 1928; Protopterus: Holmes, 1950). Despite having well organised, but small, sympathetic ganglia, lungfish possess segmentally arranged aggregations of chromaffin tissue lying around the origins of the segmental arteries, in addition to accumulations in the walls of the posterior cardinal veins and atrium (Protopterus: Holmes, 1950; Abrahamsson et al., 1979). This is reminiscent of the distribution of chromaffin tissue in elasmobranchs and cyclostomes (Nilsson, 1983). AMPHIBIANS There are marked differences in the organisation of the spinal autonomic pathways between the three major groups of extant amphibians, ranging from the primitive urodeles through the familiar anurans to the highly specialised legless amphibians, the caecilians. Urodeles Of the amphibian groups, the most primitive arrangement of the sympathetic ganglia is found in the urodeles. The following description is taken from the work of Andersson (1892: Siredon, Necturus, Ambystoma, and Salamandra) and Singer (1942: Triturus). Ganglionated sympathetic chains extend from the level of the first spinal nerve down into the tail. Except in Necturus, the sympathetic trunk does not project into the cranial region, although there are connections with the vagus nerve in all the species examined. The ganglia are usually very small, and are somewhat irregularly arranged. Nevertheless, there is an underlying segmental distribution. At more cranial levels, the sympathetic chains lie between, and are closely applied to, the posterior cardinal veins and the dorsal aorta. In some species (Triturus, Necturus, Siredon), the more cranial sympathetic trunk on each side is doubled, with a component lying either side of each lateral aorta. The largest sympathetic ganglia occur in the region of the brachial plexus, with a relatively large ganglion lying cranial and caudal to the origin of the subclavian artery. These ganglia are interconnected with each other and other smaller ganglia to form a subclavian plexus. At

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least in Triturus, the subclavian plexus gives rise to cardiac sympathetic nerves running directly to the heart, and to subclavian nerves which follow the subclavian artery into the upper limb. Splanchnic nerves supplying the stomach and proximal small intestine also originate from the subclavian plexus. There does not seem to be a coeliac ganglion situated distally along the splanchnic nerves, and they may be presumed to carry postganglionic fibres. As in fish, chromaffin tissue occurs in the walls of the posterior cardinal veins. Chromaffin cells are also prominent in the ganglia themselves (Andersson, 1892; Vogel and Model, 1977), especially in the more caudal regions of the abdominal sympathetic chains. There is no information regarding the possibility of a sacral spinal autonomic outflow in urodeles. However, Andersson describes a ganglionated “iliac plexus” with communications with the sacral nerves, which may represent the pelvic plexus in these species. Such a possibility needs to be examined with modern neuroanatomical and functional experiments. Anurans In contrast with the urodeles, the sympathetic chains of anuran amphibians are well organised with prominent segmentally arranged ganglia (Figure 1.13). As befits the greatly reduced number of vertebrae and spinal segments in anurans, there are usually only ten pairs of sympathetic ganglia, corresponding to the number of spinal nerves (Pick, 1970). There are no cranial sympathetic ganglia, although the sympathetic trunk extends into the head as far as the ciliary nerves (Bufo: Morris, 1975). The most cranial two pairs of ganglia tend to be larger than the rest (Huber, 1900). Spinal sympathetic ganglia. The relationship between the sympathetic chains and the aorta or spinal nerves varies between different species (Figure 1.13). In some species, such as Rana esculenta (Langley and Orbeli, 1910) and Limnodynastes spp. (Hill, Watanabe and Burnstock, 1975), the ganglia closely follow the course of the lateral and dorsal aortae. These ganglia have long communicating rami with the spinal nerves, and can be considered to lie in a more prevertebral position (Figure 1.13a). At the other extreme is Bufo, in which the ganglia are closely applied to the spinal nerves either side of the vertebral column. Thus, the sympathetic chains in Bufo have a genuine paravertebral location. Consequently, the ganglia have relatively short communicating rami with the spinal nerves (Langley and Orbeli, 1910; Figure 1.13b). Species such as R. temporaria (Gaskell and Gadow, 1884, Langley and Orbeli, 1910), R . catesbeiana (Bishop and O’Leary, 1938), and R. pipiens (Pick, 1970) have an intermediate arrangement of their ganglia. Remarkably, the sympathetic chains in Bufo initially develop closely applied to the dorsal aorta, in locations corresponding to those seen in adult R . esculenta. Only later do they move dorsally to become more closely associated with the spinal nerves (Jones, 1905). All of the postganglionic neurons in the chain ganglia synthesise adrenaline. They fall into two main functional classes that can be distinguished by morphological, electrophysiological and neurochemical criteria (Figure 1.2d). Small neurons (C cells) with slow conduction velocities contain the neuropeptides, NPY and galanin,

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b

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Bufo marinus

FIGURE 1.13 Different locations of the sympathetic chains in two species of anuran amphibians, a) Rana esculenta. The sympathetic ganglia (solid black) closely follow the course of the lateral and dorsal aortae. The coeliac ganglia (CG) are connected to the sympathetic chain via short splanchnic nerves. However, there are long communicating rami between the sympathetic ganglia and the spinal nerves (indicated by their segmental number). Adapted from Taxi (1976). b) Bufo marinus. Here, the sympathetic ganglia are closely applied to the origins of the spinal nerves, with which they are connected via short communicating rami. The coeliac ganglia (CG) and renal ganglia (RG) are connected to the sympathetic chain via long splanchnic nerves. Adapted from Boyd, Burnstock and Rogers, (1964) and Morris and Gibbins (1983).

and innervate blood vessels (Morris et al., 1986a, 1989; Horn, Stofer and Fatherazi, 1987, 1988). Large neurons with fast conduction velocities lack neuropeptides and innervate mucous and poison glands in the skin (Nishi, Soeda and Kukedu, 1965; Dodd and Horn, 1983; Horn, Stofer and Fatherazi, 1987, 1988; Morris et al., 1986a, 1989). These correlations between peptide content, size and function are remarkably similar to those seen in the superior cervical ganglia of mice (see Gibbins, 1991). The levels of the preganglionic outflow to the spinal sympathetic ganglia have been a source of much debate for many years. When taken together, functional studies both in Rana spp. and Bufo spp. have established that sympathetic

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preganglionic neurons leave the spinal cord via the ventral roots of segments two to eight, inclusive (Langley and Orbeli, 1910; Uyeno, 1922; Bishop and O’Leary, 1938; Dodd and Horn, 1983; Horn and Stofer, 1988). Whilst segments two to seven carry preganglionic neurons in visceral, vascular and cutaneous pathways, the outflow from the eighth spinal segment lacks the visceral component (Uyeno, 1922, cf. Langley and Orbeli, 1910, 1911). In R. catesbeiana, there is little cutaneous secretomotor outflow caudal to the sixth segment and the outflow from the eighth ventral root is predominantly vasomotor (Dodd and Horn, 1983; Horn and Stofer, 1988; Horn, Fatherazi and Stofer, 1988, cf. Langley, 1911). However, in Bufo, preganglionic neurons in cutaneous secretomotor pathways extend caudally as far as the eighth spinal segment (Uyeno, 1922: Brücke, 1922; cf. Langley and Orbeli, 1911). As in mammals, both preganglionic and postganglionic fibres travel along the sympathetic chains of anurans (Langley and Orbeli, 1910, 1911; Bishop and O’Leary, 1938; Jan and Jan, 1982; Horn and Stofer, 1988; Horn, Fatherazi and Stofer, 1988). Postganglionic fibres from the first three or four ganglia travel craniad to join the cranial nerves (Gaskell and Gadow, 1884; Figure 1.8). In particular, the vagus nerve is a major pathway for postganglionic sympathetic fibres reaching the stomach, lungs and heart, and thus is a genuine vago-sympathetic trunk (Gaskell and Gadow, 1884; Langley and Orbeli, 1910; Campbell, 1969, 1971; Morris, Gibbins and Clevers, 1981; Nilsson, 1983). Prevertebral ganglia. Anurans have a well-defined coeliac ganglion situated at the origin of the coeliac artery. In species such as R. esculenta and R . pipiens, where the chain ganglia are relatively closely applied to the lateral aortae, the splanchnic nerves carrying preganglionic fibres to the ganglion are very short (Langley and Orbeli, 1910; Figure 1.13a). In some cases, the coeliac ganglion may be at least partially fused with the fifth pair of chain ganglia (Pick, 1970; Taxi, 1976). Conversely, in species like Bufo with true paravertebral ganglia, the splanchnic nerves are long (Morris and Gibbins, 1983; Figure 1.13b). In both genera, the splanchnic nerves carry preganglionic fibres originating from spinal segments two to seven (Bishop and O’Leary, 1938; Pick, 1970; Nilsson, 1983). The coeliac ganglion provides adrenergic innervation to the gastro-intestinal tract and spleen (Gibbins, 1981; Nilsson, 1983). Small collections of ganglion cells occur along the renal nerves (R. temporaria: Langley and Orbeli, 1910; Bufo: Morris and Gibbins, 1983; Figure 1.13b) and around the origin of the inferior mesenteric artery (Langley and Orbeli, 1910). They presumably innervate the vasculature, especially the renal circulation (Morris, 1983). Pelvic ganglia. There are easily identified aggregations of non-adrenergic ganglion cells associated with the urinary bladder, hindgut and the corresponding mesentries of anurans (Huber, 1900). They provide excitatory innervation to the smooth muscle of these organs. In Bufo, they contain somatostatin and galanin (Gibbins, 1983; Morris et al., 1989). The preganglionic inputs to these ganglia arise predominantly from the ninth and tenth ventral roots and reach them via the pelvic nerves (Gaskell,

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1886; Müller, 1908; Langley and Orbeli, 1910, 1911; Boyd, Burnstock and Rogers, 1964; Burnstock, O’Shea and Wood, 1963). Thus, there is good evidence that anuran amphibians possess a more caudal spinal autonomic outflow that is anatomically and neurochemically distinct from the spinal sympathetic outflow, and is comparable in many respects to the sacral parasympathetic outflow to the pelvic plexus of mammals (Gaskell, 1886). Chromaffin tissue. Small groups of chromaffin cells are found in the sympathetic ganglia and along the various sympathetic nerves. They are also prominent around the dorsal aorta, particularly at the levels of the renal arteries (Hill, Watanabe and Burnstock, 1975; Taxi, 1976; Weight and Weitsen, 1977; Morris and Gibbins, 1983; Nilsson, 1983). However, the largest aggregations of chromaffin cells occur in the adrenal tissue on the ventral surface of the kidneys. Caecilians The sympathetic chain in caecilians is much reduced compared with other amphibians, and, indeed, with any other tetrapods (Norris and Hughes, 1918). In Herpele, Geotryptes, Hypogeophis, and probably Dermophis, only two pairs of ganglia are present: one usually is associated with the vagal ganglion and the other, larger, one lies between the levels of the second and third spinal nerves. There may be three or four pairs of ganglia in Caecilia (see Norris and Hughes, 1918, for references and discussion on these different species). The sympathetic trunk runs a short distance caudal from the last ganglion before apparently ending in the fourth spinal nerve. Cranially, the sympathetic trunk runs as far as the trigeminal ganglion, and, presumably, postganglionic sympathetic fibres project out along the trigeminal, facial, glossopharyngeal and vagal nerves. This dramatic reduction of the sympathetic system may be a consequence of their limbless condition with a concomitant reduction in the necessity for specific regional control of the circulation, for example. Furthermore, their specialised burrowing and nocturnal lifestyle may serve to protect these strange creatures from both environmental and physiological extremes. REPTILES AND BIRDS In general, the spinal sympathetic pathways of reptiles and birds are well organised, with clearly defined paravertebral chains. Although there are clear differences in organisation of the sympathetic and pelvic ganglia between the two groups, some reptiles and birds share features not seen in other vertebrates. Lizards The paravertebral sympathetic chains of lizards may contain from 15 to 25 pairs of ganglia, each one corresponding to a spinal nerve. The most cranial ganglion is at the level of the brachial plexus and is considerably larger than the rest (Lacerta, Chamaleo, Hatteria, Varanus, Tejus: Hirt, 1921; Lacerta: Adams, 1942; Tiliqua, Ctenophorus, Tympanocryptus, Pogona, Gemmatophora, Phyllodactylus: I.L.

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Gibbins, unpublished observations). From this ganglion arise nerves running to the heart and upper limb. The sympathetic trunk extends craniad and probably makes extensive communications with the cranial nerves (Hirt, 1921; Adams, 1942). Preganglionic fibres leave the spinal cord at all levels from the brachial to lumbar plexuses: there are no cervical or pelvic preganglionic outflows to the sympathetic chain (Lacerta: von Reibnitz, 1923; Terni, 1931; Gongylus: Terni, 1931). There do not seem to be any prevertebral ganglia associated with the arteries supplying the gastro-intestinal tract (von Reibnitz, 1923). Nevertheless, paravertebral ganglia at the level of the pylorus and proximal small intestine are somewhat enlarged and tend to have longer communicating rami, thereby forming something of a coeliac plexus (Hirt, 1921; Nilsson, 1983). The ganglia in this region send several fine nerves towards the gut. Virtually all of the postganglionic neurons in the sympathetic chain of lizards synthesise catecholamines. The great majority of them are likely to innervate the cardiovascular system, although most of the viscera do have a sympathetic innervation (McLean and Burnstock, 1967a,b; Furness and Moore, 1970; I.L. Gibbins, J.L. Morris and S. Holmgren, unpublished observations). Furthermore, in chameleons (but probably no other major group of lizards), there is a well documented sympathetic innervation of the cutaneous chromatophores (Sand, 1935). Whilst there are no obvious prevertebral ganglia in lizards, there is a well defined pelvic ganglion (Hirt, 1921; I.L. Gibbins, unpublished observations). Little is known of the connections of this ganglion, but it probably receives input from sacral spinal nerves (Hirt, 1921), and may contribute to the non-noradrenergic innervation of the pelvic viscera, such as the urinary bladder (Burnstock and Wood, 1967; Berger and Burnstock, 1979). Small numbers of chromaffin cells occur within sympathetic ganglia, and they have a wide, but irregular, distribution throughout the cardiovascular system. However, the main aggregation of chromaffin tissue is in the adrenal glands situated at the cranial pole of the kidneys. Chelonians The arrangement of the sympathetic ganglia in chelonians seems to be similar to that in lizards (Hirt, 1934). There is a particularly large ganglion at the level of the brachial plexus, which gives rise to cardiac and pulmonary nerves. The cervical sympathetic trunk may bear two to three small ganglia; these ganglia also may contribute to the cardiac nerves, as well as to the innervation of cranial structures {Testudo, Chelone, Emys: Gaskell and Gadow, 1884; Terni, 1931; Chelydra: Huber, 1900). In the head, the cranial sympathetic trunk communicates with at least the vagal, glossopharyngeal and facial nerves (Testudo: Terni, 1931; Chelydra, Eretmochelys: Soliman, 1964) and probably the trigeminal nerve (Trionyx: Ogushi, 1913). Caudally, the trunk communicates with the sacral nerves and continues into the tail (Huber, 1900). Developmental studies suggest that chelonians possess a ganglionated prevertebral sympathetic plexus and pelvic ganglia that have some similarities with that of birds (Thalassochelys, Chelydra: Kuntz, 191 Id; see below). According to Huber (1900), there are two types of neurons in the sympathetic ganglia of tortoises: large monopolar cells, similar to amphibian neurons, and small

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multipolar cells, more like those seen in mammals. Nothing is known of the peripheral projections and functions of these different classes of neurons. Crocodilians The spinal sympathetic ganglia of crocodilians have a number of peculiarities in their arrangement (Alligator: Gaskell and Gadow, 1884; Hirt, 1934; Alligator, Caiman, Crocodylus: Bellairs and Shute, 1953). In the thoraco-lumbar region, the ganglia are segmentally arranged, lying very close to the corresponding spinal nerves to which they are connected by short communicating rami. The most cranial of the thoracic ganglia lies at the level of the brachial plexus (eleventh body segment), and, as in lizards, it is much larger than the other paravertebral ganglia. In addition to feeding into the brachial plexus, the ganglion gives rise to a prominent cardiac nerve. Consequently, it has been named the “cardiac ganglion” (Gaskell and Gadow, 1884), although it clearly corresponds to the stellate ganglion of mammals. Cranial to this ganglion the main sympathetic chains enter the transverse foramen of the cervical vertebrae and bear a small ganglion at each cervical segment up until the third spinal nerve. The cervical sympathetic ganglia at spinal levels five to eight send small branches ventrally which combine to form a prominent, single ganglionated sympathetic trunk running along the midline, ventral to the cervical vertebrae. The median trunk closely follows the subvertebral artery, to which it supplies many fine branches, and then divides at the level of the upper cervical vertebrae to rejoin the main cervical sympathetic trunks as they enter the head. In the cranial region, the sympathetic trunks communicate with all of the major cranial nerves and probably reach the ciliary nerves. A series of prevertebral ganglia follows the aorta from the origin of the coeliac artery caudad to the renal arteries (Hirt, 1934). Recent immunohistochemical studies (S. Holmgren and I.L. Gibbins, unpublished observations) suggest that most, if not all, of the postganglionic sympathetic neurons synthesise catecholamines, and contain both NPY and somatostatin. Snakes The anatomical arrangement of the spinal autonomic system in snakes is a great mystery. In Elaphe and Thamnophis, there is supposed to be a “craniocervical sympathetic trunk” communicating between the sympathetic trunk itself, the first spinal nerve and the cranial nerves (Auen and Langebartel, 1977). Furthermore, there clearly is a noradrenergic innervation of the cardiovascular system in Elaphe (Donald and Lillywhite, 1988). Nevertheless, there has never been a convincing description of a sympathetic chain in any snake species (Hirt, 1934). Limited dissections in our laboratory have failed to find any trace of a “conventional” sympathetic chain in snakes. Possibly, the sympathetic ganglia are somewhat cryptic: perhaps they lie within the vertebral column, or they may be primarily “prevertebral”. Alternatively, snakes may have converged with the legless caecilian amphibians and similarly have, at best, a vestigial spinal sympathetic outflow. Obviously, more work is required in this area!

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Birds Spinal sympathetic ganglia. The sympathetic chains of birds extend from upper cervical to sacral levels (Figure 1.14). The ganglia are well formed and are segmentally arranged with short communicating rami with their corresponding spinal nerves. As in crocodiles, ganglia are present in the cervical sympathetic trunk which runs through the transverse foramina of the cervical vertebrae. The most cranial of the sympathetic ganglia, the “supreme cervical ganglion”, lies between the origins of the glossopharyngeal and vagus nerves as they leave the skull (Gallus, Columba, Anas, Anser: Terni, 1931; Hirt, 1934; Pick, 1970; Figure 1.14). In fowls and pigeons, there is strong functional and histological evidence that preganglionic outflow to the sympathetic chain has a much more restricted distribution, extending only from the last (16th) cervical segment to the fourth lumbar segment (Langley, 1904; Terni, 1931). Thus, there is no cervical preganglionic input to the cervical sympathetic ganglia. Postganglionic neurons in the cervical ganglia project into the cervical spinal nerves and to cranial structures, such as blood vessels, penna-motor muscles, and, presumably, glands (Langley, 1904; Bennett, 1974). Most of the postganglionic neurons in the sympathetic chains contain catecholamines (Bennett, 1974). The major targets of these neurons are blood vessels and the penna-motor muscles, which may be innervated by two functionally distinct classes of neurons: one raising the feathers and one lowering them (Langley, 1904). Prevertebral ganglia. In contrast with reptiles, there are well developed prevertebral ganglia in birds (Nolf, 1934a, b; Pick, 1970; Bennett, 1974). A ganglionated coeliaco-mesenteric plexus surrounds the aorta and the coeliac and mesenteric arteries at their origin, and is connected with the paravertebral chain by a series of splanchnic nerves (Figure 1.14). Small ganglia also are associated with the adrenals and kidneys. Extending from the caudal pole of the kidneys towards the cloaca, there is a group of ganglia forming a pelvic plexus (Hirt, 1934; Nolf, 1934c; Pick, 1970). Many of these neurons do not contain catecholamines, and have been presumed to be cholinergic (Bennett, 1974). Pelvic ganglia - Remakes nerve. The most remarkable feature of the spinal autonomic pathways of birds is the ganglionated nerve of Remak. It extends from the caudal end of the pelvic plexus through the mesentery of the intestine until it merges with the coeliac plexus cranially. It receives inputs from the sacral spinal nerves, but also communicates with the sympathetic chain via the pelvic plexus caudally and the coeliaco-mesenteric plexus cranially (Nolf, 1934c). The ganglia contain neurons with and without catecholamines, but there is a higher relative proportion of catecholamine-containing neurons at the cranial end of Remak’s nerve (Costa, 1966; Bennett, 1974; Young, 1990). The ganglia mostly supply the intestine (Nolf, 1934c), with the non-catecholamine neurons causing a non-cholinergic contraction of the hindgut (Bartlet and Hassan, 1971; Takewaki, Ohashi and Okada, 1977; Komori and Ohashi, 1982).

FIGURE 1.14 Sympathetic ganglia of a bird, Gallus. Note the numerous cervical sympathetic ganglia extending from the supreme cervical ganglion to the level of the brachial plexus. The coeliac plexus surrounds the origin of the coeliac artery from the aorta. Spinal nerves are indicated by dashed lines. Adapted from Pick (1970).

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GENERAL FEATURES AND EVOLUTION OF THE SPINAL AUTONOMIC OUTFLOWS Chromaffin cells and sympathetic neurons Whereas chromaffin tissue seems to be the main source of regulation of the cardiovascular system in cyclostomes, it is clear that this function has been taken over to a very large degree by spinal sympathetic neurons in amniotes. Within the various groups of fish, there are two well-defined intermediate stages: elasmobranchs possess segmentally arranged ganglia, each associated with large amounts of chromaffin tissue, whilst teleosts, which appeared in the fossil record considerably later than the elasmobranchs, have a well organised sympathetic chain, but still retain a large mass of chromaffin tissue on the posterior cardinal veins. In most tetrapods (one exception being urodeles), the separation of chromaffin tissue and sympathetic ganglia has been more complete, with the development of a concentrated mass of adrenal medullary tissue associated with the ventral surface (anuran amphibians) or cranial pole (amniotes) of the kidneys. Developmental studies provide good evidence that a phylogenetic shift from chromaffin cells to neurons is a realistic scenario. Developing sympathetic neurons have many morphological and biochemical features in common with chromaffin cells, both classes of cell being derived embryologically from the neural crest. Furthermore, under appropriate experimental manipulation, sympathetic progenitor cells normally destined to become chromaffin cells can be induced to differentiate into cells with many of the features of sympathetic neurons (for references, see Yntema and Hammond, 1947; Horstadius, 1950; Purves and Lichtman, 1985; Jacobson, 1991). Indeed, recent experiments have shown that the common precursor of chromaffin cells and sympathetic neurons share more biochemical markers with chromaffin cells than with neurons (Anderson et al.9 1991). There seems to be little doubt that sympathetic ganglion cells primitively synthesise and release catecholamines, as would befit a chromaffin cell ancestry. Nevertheless, sympathetic neurons without catecholamines appear to varying degrees in different species, ranging from catfish to humans. In developing rats, cholinergic sympathetic neurons innervating the sweat glands gradually acquire their cholinergic characteristics after initially possessing many features of catecholamine-synthesising neurons (Yodlowski, Fredieu and Landis, 1984; Leblanc and Landis, 1986; Stevens and Landis, 1987; Landis, Siegel and Schwab, 1988). Furthermore, at least some non-noradrenergic neurons in the sympathetic chain ganglia of adult guinea-pigs contain enzymes in the catecholamine synthesis pathway, even though they do not normally contain catecholamines (Gibbins, 1992). Finally, some cholinergic sympathetic neurons in adult fish retain certain properties of catecholamine-synthesising neurons, most notably the ability to take up exogenous catecholamines and their analogues from the external environment (Winberg, Holmgren and Nilsson, 1981). All these observations are consistent with cholinergic neurons having been a secondary development in the evolution of sympathetic pathways.

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Prevertebral and paravertebral ganglia Concomitant with the shift in emphasis from chromaffin tissue to ganglion cells has been a gradual change in the location of the segmental collections of chromaffin and ganglion cells from the segmental arteries to the segmental nerves. In cyclostomes, elasmobranchs and dipnoans, the segmental aggregation of chromaffin cells surround branch points of the dorsal aorta, especially the origins of the segmental arteries, where the cells are presumably well placed to secrete catecholamines into the various levels of the systemic circulation. In amniotes, however, the sympathetic ganglia have a classical paravertebral location. Generally, they are intimately associated with the spinal nerves which provide paths for their preganglionic inputs and their postganglionic outflows. Once again, an intermediate arrangement is found in teleosts, in which the ganglia are segmentally arranged, but closely follow the course of the dorsal aorta. In this context, the situation in anuran amphibia is interesting. Some species, such as Rana esculenta, have sympathetic chains closely following the lateral and dorsal aortae, very much as in teleosts, whilst others, such as Bufo, have paravertebral chains, much more like those in amniotes. The presence or otherwise of prevertebral sympathetic ganglia seems to be a consequence of the position of the sympathetic chains relative to the aorta and vertebral column. In species with paravertebral chains, the prevertebral ganglia tend to be more prominent than in species with “para-aortic” sympathetic chains. When present, the prevertebral ganglia are always closely associated with the origins of the major ventral branches of the aorta supplying the gastro-intestinal tract and its

a Gallus embryos FIGURE 1.15 Transverse sections of chick embryos (Gallus) showing the development of the primary and secondary sympathetic chains. Compare with Fig. 1.10. Adpated from Terni (1931). a) 100 hours post-fertilisation, second thoracic segment. The primary site of neural crest cell aggregation (1°) has been around the origin of the segmental arteries (sa) from the dorsal aorta (da). The secondary migration to more dorsal sites close to the origin of the spinal nerves (2°) has begun. DRG, dorsal root ganglia; nch, notochord, b) 7 days post-fertilisation, fourth thoracic segment. The definitive sympathetic chain is now in position (2°). Neurons remaining in the original position of the primary chain around the dorsal aorta (da) form the prevertebral ganglia and plexuses. DRG, dorsal root ganglia; nch, notochord.

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derivatives, suggesting that they have retained their primitive close relationship with branch points of the aorta. Developmental studies again provide strong circumstantial evidence for this phylogenetic sequence in the position of the sympathetic chains (Figure 1.15). During the development of the main sympathetic chains in toads, reptiles and birds, the sympathetic progenitor cells initially aggregate around the aorta and its branch points, including the segmental arteries (Figure 1.15a; cf. Figure 1.10). This initial migration pathway closely resembles that seen in selachians (Acanthias), holosteans {Amid), teleosts (Opsanus: Kuntz, 1911a) and many amphibians {Ambystoma, Necturns, Rana: Kuntz, 1911c; Vogel and Model, 1977). However, at later stages in the development of toads and amniotes, the primordia of the sympathetic chain ganglia migrate dorsad to lie in their definitive paravertebral position, ventral to the spinal nerves (Bufo: Jones, 1905; Thalassochelys, Chelydra: Kuntz, 191 Id; Lacerta: von Reibnitz, 1923; Gallus: Kuntz, 1922; van Campenhout, 1930; Terni, 1931; Yntema and Hammond, 1947). The prevertebral ganglia remain aggregated around the ventral branch points of the aorta (Figure 1.15b). The ganglionated sympathetic trunk lying in the ventral midline at the cervical levels of crocodiles seems to be a persistent remnant of the initial ventral aggregation of sympathoblasts in this region (Bellairs and Shute, 1953). Why a secondary migration of most of the sympathetic ganglia should occur is a mystery, but it may be related to the necessity for more precise connections with the spinal nerves as part of a general trend towards an increased complexity of connectivity in the spinal autonomic pathways in amniotes. In all vertebrates, there is a major spinal sympathetic outflow corresponding to the level of the brachial plexus. At this level in reptiles and mammals is the stellate ganglion, invariably the largest of the paravertebral sympathetic ganglia. Indeed, in caecilian amphibians, one of the few sympathetic ganglia present at all occurs at this level. The stellate ganglion provides sympathetic innervation of several major structures, including the heart, lungs and forelimb. In fish, this is the level where the largest mass of chromaffin tissue occurs, well sited to regulate the activity of the heart. Thus, it seems likely that the presence of a large collection of sympathetic tissue at this level has had a very long evolutionary history, and that a large stellate ganglion is a primitive condition for tetrapods in general, rather than being an aggregation of a series of smaller segmental ganglia, like the cervical ganglia of birds (see Terni, 1931). The brachial spinal levels also provide preganglionic outflows to sympathetic ganglia innervating cranial effectors. Each major vertebrate group seems to have evolved independently a unique anatomical arrangement of cranial or cervical sympathetic ganglia (van den Broek, 1908; Terni, 1931; Singer, 1942). But, regardless of their anatomical locations, there is never a cervical preganglionic outflow. Pelvic pathways One of the initial propositions to be examined here was that the division between the lumbar and sacral spinal autonomic outflows was an artificial one that could not be maintained across a wide selection of vertebrate groups (see Nilsson, 1983).

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However, it appears that probably all vertebrates, including cyclostomes, do possess a distinct spinal preganglionic outflow to ganglion cells innervating the hindgut and its derivatives in the pelvic viscera. Furthermore, in the few cases where this problem has been specifically examined, there is no overlap between the segmental origin of the preganglionic outflow to the sympathetic chains and the preganglionic outflow to the pelvic ganglia. Nevertheless, the homology of the spinal outflow to the pelvic organs of teleosts and elasmobranchs with the sacral spinal outflow of tetrapods has not been established unambiguously (Young, 1933). Clearly, this question needs to be examined more closely in a wider variety of species. Despite the presence of a well demarcated spinal outflow to the pelvic ganglia, it is none the less true that the pelvic plexus is closely associated with both sympathetic and sacral pathways, especially in amniotes. Furthermore, the cranial to caudal gradient in neurochemical properties of Remak’s nerve in birds closely follows that of the prevertebral and pelvic ganglia in mammals. On the basis of developmental studies, Kuntz (191 Id) concluded that the arrangement of the prevertebral and pelvic plexuses in reptiles is intermediate between that in mammals and the prevertebral plexus-Remak’s nerve-pelvic plexus complex in birds. Embryological experiments suggest that the neurons of the pelvic plexus, including much of Remak’s nerve, arise from a more caudal population of neural crest cells that is anatomically separated from those developing into sympathetic neurons (Jones, 1937; Yntema and Hammond, 1947; Le Douarin, 1982). Thus, the apparent cranial to caudal gradients in both neurochemistry and source of preganglionic inputs to the prevertebral and pelvic ganglia may well be a result of a gradual intermixing between neurons with embryological origins from either thoraco-lumbar or sacral levels of the neural crest.

CONCLUSIONS I began this review by asking whether or not Langley’s scheme for classifying the various components of the ANS could be applied across the vertebrate classes. The evidence, such as it is, seems to say that indeed it can. The enteric neurons comprise a separate autonomic division in all classes of vertebrates. It is almost certainly the most primitive component of the autonomic system, apparently being present in non-vertebrate chordates, such as amphioxus, that lack any other obvious components of an “autonomic” nervous system. The cranial autonomic outflow clearly can be divided into two main divisions: one controlling the eye via the oculomotor nerve and the ciliary ganglion; the other related to the primitive branchiomeric structure of the jaws and pharyngeal region, with preganglionic outflow running via the facial, glossopharyngeal and vagus nerves. The sympathetic chains are supplied by a well defined spinal outflow from the level of the brachial plexus to the lower abdominal region. Similarly, in most species, there is a well defined spinal outflow, caudal to that innervating the sympathetic chains, that provides preganglionic input to neurons in the pelvic plexuses. The true nature of the prevertebral ganglia is still somewhat enigmatic, as is their relationship with the pelvic plexuses. The more

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cranial prevertebral ganglia clearly lie on spinal sympathetic pathways, but the situation is not so obvious for the more caudal ganglia. Further studies are required to define the relationships between the sacral and lumbosacral spinal outflows in a wide range of vertebrate species. Although there are still great gaps in our knowledge of the structure and function of the ANS of the main vertebrate groups, one trend is remarkably clear: there has been a marked increase in the level and precision of organisation of the peripheral autonomic pathways during the evolution of more advanced forms. Overall, there is little difference between the different amniote classes: reptiles, birds and mammals. Significant jumps in organisation occur between cyclostomes and elasmobranchs; elasmobranchs and teleosts; teleosts and anuran amphibians; and anurans and amniotes. Nevertheless, intermediate grades of organisation exist, especially in terms of the spinal sympathetic pathways. Thus, in many respects, dipnoans seem to lie between elasmobranchs and teleosts (Hirt, 1934), and urodeles seem to fall between dipnoans and anurans (Singer, 1942). This ordering does not seem to be at odds with most currently accepted schemes of vertebrate phylogeny, which postulate polyphyletic origins for all the major groups of modern fish, and perhaps also for the main subclasses of extant amphibians (see Carroll, 1988). The increase in organisation and complexity of the peripheral ANS across vertebrate phylogeny is parallel to that seen in the central nervous system. Presumably, it is related most closely to the requirements for increasingly efficient control of the cardiovascular system, gut and other internal organs in response to the high energy demands imposed by lifestyles progressively more independent of environmental constraints (see Hochachka and Somero, 1984; Bray, 1985; Sibly and Calow, 1986; Carroll, 1988; Little 1990).

ACKNOWLEDGEMENTS The author’s own work has been supported by grants from the National Health and Medical Research Council of Australia, the National Heart Foundation of Australia, the Australian Research Council, and the Flinders Medical Centre Research Foundation. I would like to thank Sue Matthew and the staff of the Flinders University Medical Library for valuable assistance in obtaining reference material, and Dr Judy Morris for critical advice on the manuscript and assistance in preparing the plates.

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Morris, J.L. (1983). Effects of nerve stimulation on vascular resistance in the toad kidney. NaunynSchmiedeberg's Arch. Pharmacol., 323, 335-340. Morris, J.L. and Gibbins, I.L. (1983). Innervation of the renal vasculature of the toad {Bufo marinus). Cell Tissue Res., 231, 357-376. Morris, J.L. and Gibbins, I.L. (1992). Co-transmission and neuromodulation. In: The Autonomic Nervous System. Vol. I. Autonomic Neuroeffector Mechanisms., edited by G. Burnstock and C.H.V., Hoyle, pp . 33-119. Chur: Harwood. Morris, J.L., Gibbins, I.L. and Clevers, J. (1981). Resistance of adrenergic neurotransmission in the toad heart to adrenoceptor blockade. Naunyn-Schmiedebergs Arch. Pharmacol., 317, 331-338. Morris, J.L., Gibbins, I.L., Campbell, G., Murphy, R., Furness, J.B. and Costa, M. (1986a). Innervation of the large arteries and heart of the toad (Bufo marinus) by adrenergic and peptide-containing nerves. Cell Tissue Res., 243, 171-184. Morris, J.L., Gibbins, I.L. and Murphy, R. (1986b). Neuropeptide Y-like immunoreactivity is absent from most perivascular noradrenergic axons in a marsupial, the brush tailed possum. Neurosci. Lett., 71, 264-270. Morris, J.L., Gibbins, I.L. and Osborne, P.B. (1989). Galanin-like immunoreactivity in sympathetic and parasympathetic neurons of the toad Bufo marinus. Neurosci. Lett., 102, 142-148. Müller, R. (1908). Uber die Nervenversorgung des Magendarmkanales beim Frosch durch Nervennetze. Pflüger’s Archiv für die gesamte Physiologie, 123,387-405 Müller, E. and Liljestrand, F. (1918). Anatomische und experimentalle Untersuchungen über das autonome Nervensystem der Elasmobranchier nebst Bemerkungen über die Darmnerven bei den Amphibien und Saugetieren. Arch. A nat., 1918, 137-172. Nakao, T. (1981). An electron microscopic study on the innervation of the gill filaments of a lamprey, Lampetra japonica. J. Morphol., 169, 325-336. Nakao, T. and Ishizawa, A. (1982). An electron microscopic study of autonomic nerve cells in the cloacal region of the lamprey, Lampetra japonica. J. Neurocytol., 11, 517-532. Nakao, T., Suzuki, S. and Saito, M. (1981). An electron microscopic study of the cardiac innervation in larval lampreys. Anat. Rec., 199, 555-563. Nicol, J. (1950). The autonomic nervous system of the chimaeroid fish, Hydrolagus colliei. Q. J. Microscop Sei., 91, 379-399. Nicol, J. (1952). Autonomic nervous systems in lower chordates. Biol. Rev., 27, 1-49. Nilsson, S. (1981). On the adrenergic system of ganoid fish: the Florida spotted gar, Lepisosteus platyrhincus (Holostei). Acta Physiol. Scand., I l l , 447-454. Nilsson, S. (1983). Autonomic Nerve Function in Vertebrates. Berlin: Springer-Verlag. Nilsson, S. (1984a). Innervation and pharmacology of the gills. In: Fish Physiology, Vol. 10A, edited by W.S. Hoar and D.J. Randall, pp . 185-227. New York: Academic Press. Nilsson, S. (1984b). Adrenergic control systems in fish. Marine Biol. Lett., 5, 127-146. Nilsson, S. and Holmgren, S. (1988). The Autonomic Nervous System. In: Physiology o f Elasmobranch Fishes, edited by T.J. Shuttleworth, pp . 143-169. Berlin: Springer-Verlag. Nishi, S., Soeda, H. and Kukedu, K. (1965). Studies on sympathetic B and C neurons and patterns of preganglionic innervation. J. Cell. Comp. Physiol., 66, 19-32. Nolf, P. (1934a). Les nerfs extrinsèques de l’intestin chez l’oiseau. I. Les nerfs vagues. Arch. Int. Physiol. Biochim., 39, 113-164. Nolf, P. (1934b). Les nerfs extrinsèques de l’intestin chez l’oiseau. III. Le nerf de Remak. Arch. Int. Physiol. Biochim., 39, 227-256. Nolf, P. (1934c). Les nerfs extrinsèques de l’intestin chez l’oiseau. II. Les nerfs coeliaques et mesenteriques. Arch. Int. Physiol. Biochim, 39, 165-226. Nomura, Y. and Matsuura, T. (1972). Distribution and clinical significance of the autonomic nervous system in the human nasal mucosa. Acta Oto-Laryngol., 73, 493-501. Norris, H.W. (1908). The cranial nerves of Amphiuma means. J. Comp. Neurol., 18, 527-568 Norris, H.W. (1925). Obervations upon the peripheral distribution of the cranial nerves of certain ganoid fishes (Amia, Lepidosteus, Polyodon, Scaphirhyncus, and Acipenser). J. Comp. Neurol. 39, 345-432. Norris, H.W. and Hughes, S.P. (1918). The cranial and anterior spinal nerves of caecilian amphibians. J. Morphol., 31, 489-560. Norris, H.W. and Hughes, S.P. (1920). The cranial nerves of Squalus acanthias. J. Comp. Neurol., 31, 293-402. Ogushi, K. (1913). Zur Anatomie der Hirnnerven und des Kopfsympathicus von Trionya japonicus nebst einigen kritischen Bemerkungen. Morphol. Jhrb., 45, 441-480.

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2 Comparative Aspects on the Biochemical Identity of Neurotransmitters of Autonomie Neurons Susanne Holmgren and Jôrgen Jensen Comparative Neuroscience Unit, Department o f Zoophysiology, University o f Göteborg, Sweden The biochemical identity of putative neurotransmitters in autonomic neurons of different vertebrate groups is discussed. Cholinergic and serotonergic neurons contain the same transmitters in all vertebrate groups: acetylcholine and 5-hydroxytryptamine, respectively. The occurrence of adrenaline and noradrenaline in different proportions in adrenergic neurons is pointed out, and the possibility of purinergic neurons releasing purine derivatives considered. Bombesin, gastrin/CCK/caerulein, neuropeptide Y (NPY), neurotensin, opioids, somatostatin, substance P (tachykinins) and vasoactive intestinal polypeptide (VIP) are all ‘members of different families of peptides’ that occur in neurons and/or endocrine or paracrine cells. Sequence variations of these peptides between vertebrate groups, and the occurrence of several related peptides in one animal are discussed. Available information on the biosynthesis of a neuropeptide in non-mammalian vertebrates has been included. KEY WORDS sequences; acetylcholine; catecholamines; 5-hydroxytryptamine; purines; neuropeptides

INTRODUCTION Over the years, several articles dealing with neurotransmitter functions have discussed criteria to be fulfilled for a substance to be considered a transmitter (e.g. Iversen, 1979; Burnstock, 1981; Nilsson, 1983; Furness and Costa, 1987). Detailed criteria were originally defined based on studies of adrenergic and cholinergic neurons, and, around 1980, involved requirements of the presence of intraneuronal precursors, of synthesis, storage, release and inactivation mechanisms, and of mimicry and pharmacological identity, i.e. exogenous application of the putative transmitter should mimic the response to nerve stimulation, and antagonists should have the same effect on the action of the exogenous putative transmitter as on the response to nerve stimulation. However, with the increasing knowledge of the 69

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diversity of transmitters and their functions and co-existense in neurons, some of these criteria may be considered subsidiary for different reasons. Remaining as central criteria for a neurotransmitter are two concepts: the substance shall be released from a neuron, and it must have an identity of action, i.e. the response to nerve stimulation should be identical to that of the putative transmitter in pharmacological tests (Furness and Costa, 1987). An increasing number of biologically active substances have now been identified as transmitters of autonomic neurons; in particular the enteric nervous system and sympathetic ganglia have proven rich sources for a variety of neurotransmitters, which coexist in the neurons in various pathway-specific combinations (Furness and Costa, 1987; Gibbins et al., 1987; Nilsson and Holmgren, 1989). Comparative studies of the identity of neurotransmitters are in many cases still fragmentary, but available information often suggests an amazing similarity in the chemical structure between the transmitters in different vertebrate groups. The aim of this chapter has been to elucidate differences and similarities in the chemical structure of vertebrate transmitters. Only those substances where research has been carried out on nonmammalian vertebrates are discussed. The amount of words devoted to each transmitter group is in no way correlated to its importance, it merely reflects on the one hand where current research is focused, and on the other hand where the variations in chemical structure of the transmitters occur.

CHOLINERGIC NEURONS Classically, acetylcholine (Figure 2.1) has been considered the transmitter of all preganglionic neurons, and of most postganglionic, parasympathetic neurons of the autonomic nervous system in all vertebrate groups (Nilsson, 1983). Cholinergic neurons also form a part of the enteric nervous system (Furness and Costa, 1987). Although it was originally thought that choline was the transmitter of the cholinergic neurons (hence the name), it was early established that the acetylated form of choline is the active transmitter (Loewi, 1921; Loewi and Navratil, 1926a, b). Acetylcholine is synthesized from acetyl-coenzyme A and choline by the enzyme choline acetyltransferase (choline acetylase, E.C. 2.3.1.6, ChAT) within the cholinergic neuron (Nachmansohn and Machado, 1943). The presence of cholinergic neurons in numerous autonomic pathways has been accepted after a vast number of pharmacological, histochemical and physiological studies (Michelson and Zeimal, 1973). Recently, with the development of immunohistochemical methods, staining of ChAT with specific antisera has proved a useful method to identify cholinergic neurons. With the exception of the primate placenta, ChAT appears to be unique to cholinergic neurons (Nilsson, 1983). The earliest experiments leading to the biochemical identification of acetylcholine were made by Loewi and coworkers (Loewi, 1921; Loewi and Navratil, 1926a, b) on the amphibian heart. The subsequent numerous studies, mainly in mammals, have provided no reason to suspect that the chemical identity of the transmitter in cholinergic nerves is not the same in different vertebrate groups. However, an

COMPARATIVE ASPECTS ON THE BIOCHEMICAL IDENTITY

HgC — C — O — CH2 — CH2— N — CH

I

CH

acetylcholine

71

NH2

w

a

N

ATP

---- N

V?

noradrenaline

NH

I

CHg

5-hydroxytryptamine

adrenaline FIGURE 2.1

The chemical structure of acetylcholine, adrenaline, noradrenaline, ATP and 5-hydroxytryptamine (serotonin).

increasing number of studies demonstrate that other substances occur as cotransmitters or modulators in cholinergic autonomic neurons (Furness and Costa, 1987; Gibbins, 1989).

ADRENERGIC NEURONS Adrenergic neurons use the catecholamines noradrenaline and adrenaline as transmitters (Figure 2.1); the proportion between the two varies between species, and between different organs (Holzbauer and Sharman, 1972). Catecholamines are synthesized along the “Holtz-Blaschko” pathway in adrenergic nerves as in the adrenal medulla (Blaschko, 1939; Holtz, 1939; von Euler, 1972). Noradrenaline is formed from tyrosine by the actions of, in turn, tyrosine hydroxylase, DOPA decarboxylase and dopamine-jS-hydroxylase (DBH). Adrenaline is formed from noradrenaline by the action of the cytoplasmatic enzyme phenylethanol-amine-N-methyl transferase (PNMT).

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PNMT has been demonstrated in chromaffin tissue of representatives of all vertebrate groups (see Santer, chapter 3 this volume), but has also been isolated from tissues innervated by adrenergic nerves in non-mammalian vertebrates (Wurtman et al., 1968; Wooten and Saavedra, 1974; Abrahamsson and Nilsson, 1976; Abrahamsson et al., 1981). This means that adrenaline can be synthesized intraneuronally and can act as a transmitter (along with noradrenaline) in these species. Assays have shown that adrenaline in fact dominates over noradrenaline in many organs of amphibians, holosteans and teleosts (Holtzbauer and Sharman, 1972; Abrahamsson and Nilsson, 1976). On the other hand, it has been proposed that noradrenaline is the sole adrenergic transmitter of mammalian autonomic neurons (von Euler, 1946). Clearly, noradrenaline is the dominating catecholamine in mammalian adrenergic nerves, but analyses of perfusates from the mammalian intestine after stimulation of mesenteric nerves showed that 5-25% of the catecholamines were adrenaline (Mann and West, 1951; Mirkin and Bonnycastle, 1954). Studies performed on the distribution of catecholamines in vertebrates have been reviewed in detail by Holtzbauer and Sharman (1972).

SEROTONERGIC NEURONS 5-Hydroxytryptamine (5-HT, serotonin; Figure 2.1) has only recently been established as a neurotransmitter in mammals, while the presence in some fish species was demonstrated at an earlier stage, possibly depending on differences in transmitter levels in different animals. With the use of the Falck-Hillarp fluorescence histochemical technique, nerve fibres showing the characteristic yellow fluorescence of 5-HT were demonstrated in the gut of cyclostome and teleost fish species (Baumgarten et al., 1973; Goodrich et al., 1980; Watson, 1979; Anderson, 1983). However, 5-HT neurons could not be demonstrated in all teleost species using the Falck-Hillarp method (Anderson, 1983). Subsequent studies, using the more sensitive method of staining with specific antisera, have revealed 5-HT fibres also in these species (Holmgren et al., 1985; Anderson and Campbell, 1988), and it may be speculated that the transmitter storing capacity of 5-HT neurons varies between species. An interesting observation was made by Anderson and Campbell (1988): no 5-HT-storing enterochromaffin cells were found in species possessing mucosal 5-HT fibres showing up with the Falck-Hillarp method (i.e. nerves with a high content of 5-HT), while enterochromaffin cells were common in species with a low content of 5-HT in the mucosal fibres. Similarly, earlier studies in mammals revealed enterochromaffin cells, but not nerve fibres, containing 5-HT while later immunohistochemical studies have demonstrated the additional presence of 5-HT in autonomic neurons (Furness and Costa, 1987; Griffith, 1988). 5-HT neurons appear to be absent in prototherian mammals, reptiles and birds (Adamson and Campbell, 1988), but are present in amphibians (Anderson and Campbell, 1989). A group of ‘amine-handling neurons’ (Furness and Costa, 1987), which take up and process circulating 5-HT, may show up in histochemical investigations, par-

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ticularly after pretreatments aimed to enhance the reactivity. They have, however, no endogenous synthesis of 5-HT, and are therefore not truly 5-HT neurons.

PURINERGIC NERVES ATP (adenosine triphosphate, Figure 2.1) fulfills the criteria for being a neurotransmitter in several mammalian tissues (Burnstock, 1972, 1986), while in other cases the release from autonomic neurons (one of the transmitter criteria) has been questioned (Furness and Costa, 1987). The possibility that ATP acts as a cotransmitter has been raised (Burnstock, 1981; Costa, Furness and Humphreys, 1986), and it has also been proposed that ATP and adenosine constitute a regulatory feedback loop: ATP released from the neuron is degraded to adenosine, which inhibits further release of neurotransmitter (Pelleg and Burnstock, 1990). There are an increasing number of studies reporting an effect of ATP and related adenyl compounds on autonomically innervated organs of non-mammalian vertebrates (e.g. Young 1980 a, b, 1983, 1988; Burnstock and Meghji, 1981; Holmgren, 1983; Meghji and Burnstock, 1984a, b; Hoyle and Burnstock, 1986; Lazou and Beis, 1987; Lennard and Huddard, 1989; Small, MacDonald and Farrell, 1990; Farrell and Davie, 1991). Although several (but not all) of the criteria set for a transmitter are fulfilled in these studies, alternative interpretations have been put forward, and it is at this time premature to conclude that ATP acts as a transmitter in the autonomic nervous system of non-mammalian vertebrates.

NEUROPEPTIDES Peptides were recognized as neurotransmitters of the autonomic and sensory nervous systems at a relatively late stage. It was understood at an early stage that several of the humoral substances released from endocrine cells were peptides; much later, by the use of immunochemical methods, it has become increasingly clear that numerous neuroregulatory substances are peptides and occur in the central nervous system as well as in peripheral nerves (and in endocrine cells). Along with the increased possibilities to determine the nature of the transmitter substances has come an increased interest in studies of autonomic nerves and their functions in mammals as well as in non-mammalian vertebrates. The demonstration of a substance P-like peptide in the intestine of both an elasmobranch, the spiny dogfish, Squalus acanthias, and a teleost, the cod, Gadus morhua, was the first finding of a regulatory peptide in a non-mammalian vertebrate (von Euler and Ostlund, 1956). The study showed a dual location in the brain and gut of the substance P-like peptide, and also indicated its biological activity in a series of mammalian preparations. Two important conclusions were drawn from this study: a. like its mammalian counterpart (Eliasson, Lie and Pernow, 1956) the non-mammalian peptide occurred in both brain and gut, and b.

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several properties of the fish polypeptide were similar to those of the mammalian peptide substance P. It was thus established at an early stage that brain-gut peptides existed in nonmammalian species as well as in mammals. Indeed, neuropeptides occur in the nervous system of all animal groups possessing a nervous system, from the most simple of coelenterates (Thorndyke and Goldsworty, 1988; Nilsson and Holmgren, 1989). However, further studies into the exact identity and function of neuropeptides in non-mammalian animals are relatively recent and still in their first stages. Compared to other known types of transmitters (acetylcholine, amines, nucleotides, amino acids) there is an enormous potential diversity for the individual in using peptides as neuronal messengers, and it has been estimated (and there is increasing evidence for this) that 100 to 200 different peptide molecules may function as neuronal signal substances in any one species (Joosse, 1988). RELATIONSHIPS BETWEEN PEPTIDES Amino acid sequences of related regulatory peptides often vary between species, usually with an increasing degree of exchange of amino acids the more distantly related two species are. Depending on the degree of substitutions, the molecule has been considered either a variant of the same peptide occurring in different species, or a separate but more or less closely related peptide belonging to the same ‘family’. In some cases, separate names have been given to peptides that may be considered variants of the same peptide, e.g. for the simple reason that the amino-acid sequences and thus the relationships were not established before the naming of the peptide. These days relationships between peptides may be accurately determined by gene and precursor studies (see e.g. Conlon, 1989). Usually one part of the sequence is better conserved than the rest of the molecule. Often this is the C-terminal part of the peptide, but e.g. VIP molecules from different animal groups are identical in the N-terminal part and have substitutions in the C-terminal region of the molecule. It is often found that the conserved part of the molecule is most important for the biological action of the peptide. Considering the time span involved, it is amazing how well conserved the biologically active part of the peptide can be. It is 450 to 500 million years since the vertebrates shared a common ancestor (Romer, 1962); still the amino acid sequence of somatostatin 14 appears to be identical in the elasmobranch Torpedo, catfish, anglerfish, pigeon and mammals (Noe et al., 1979; Spiess et al. 1979; Andrews and Dixon, 1981; Conlon et al., 1985a), and there are only six base pair differences between the nucleotide sequences encoding anglerfish somatostatin 14 and rat somatostatin 14 (Montminy et al., 1984; Hobart et al., 1980). Somatostatin is not unique in this respect, there are numerous examples of highly conserved regions in the neuropeptides (see below). There is much more variation in the regions encoding the flanking molecules of the precursor peptides, and even more substitutions in the non-transcribed introns of homologous genes (e.g. Nishizawa et al., 1985; Linder et al.9 1987). This demonstrates the high evolutionary pressure to maintain the amino acid sequences of the biologically active products, while the parts

COMPARATIVE ASPECTS ON THE BIOCHEMICAL IDENTITY

75

of the genes encoding for flanking peptides or spacer peptides have mutated more freely and randomly. When mutations have occurred in the part of the gene encoding the final neuropeptide, these are often single base pair mutations resulting in conservative amino acid substitutions of little importance for the biological properties of the molecule, such as those present in guinea pig VIP compared to porcine VIP (Du et al., 1985). Studies of sequences of genes and their products can give us valuable information about the importance of different neuropeptides. Their degree of conservation throughout the animal kingdom indicates the fundamental physiological importance of the processes they are involved in. BOMBESIN-LIKE PEPTIDES Although first demonstrated in the amphibian skin, bombesin or related peptides have subsequently been demonstrated in autonomic and central neurons as well as in endocrine cells of the gut and lung in a number of non-mammalian species, both vertebrates and invertebrates. It has been agreed that the generic family name “bombesin-like peptides” should be used for all peptides with a C-terminal amino acid sequence similar to bombesin (Erspamer and Go, 1988). Bombesin, the denominator of the group, is a 14 amino acid peptide, which was originally isolated from the skin of the amphibians, Bombina bombina and Bombina variegata variegata (Anastasi et al., 1971). Several bombesin-related peptides have now been identified, most of those are skin peptides which are so far only found in amphibians, but some are also neuropeptides (Erspamer et al., 1984, 1986). Neuromedin B is a peptide resembling ranatensin R and ranatensin C, which has been isolated from mammalian tissues (Minamino et al., 1983). Studies in mammals and birds have also revealed the presence of the so called gastrin releasing peptides (GRPs), named because of their effect in the first bioassay studies. The mammalian GRPs are neuronally contained peptides of 27, 23 or 10 amino acids, with nine of the ten C-terminal residues identical to the bombesins (McDonald et al., 1979; Reeve et al., 1983; Orloff et al., 1984; Table 2.1). GRPs in general show the same biological effects as bombesins and were first considered the mammalian counterpart to the amphibian peptide. However, a frog GRP of 10 amino acid residues (initially named neuromedin C), identical to the Cterminal of chicken GRP, has been isolated from the frog brain and intestine, and it is possible that the neuronally contained bombesin-like peptides in amphibians are closer related to GRP than to the skin peptide bombesin (Conlon et al.9 1991e; Nagalla et al., 1992). This is supported by the fact that nucleotide sequence analyses of cDNAs encoding human GRP and amphibian bombesin suggest that the two peptides are not homologous (Spindel et al., 1984; Richter et al., 1990) Bird (chicken) GRP-27 has been sequenced, and differs from porcine GRP in nine positions, mainly in the N-terminal part (McDonald et al., 1980). To date, bird GRP has only been described in endocrine cells of the gut and not in peripheral nerves (Timson et al.9 1979; Vaillant et al., 1979; own unpublished results from the crested tit, Parus cristatus). The primary structure of a GRP from rainbow trout stomach

76

COMPARATIVE PHYSIOLOGY AND EVOLUTION TABLE 2.1 Amino acid sequences of bombesin-like peptides.

bombesin alytesin porcine GRP human GRP canine GRP chicken GRP shark GRP rainbow trout GRP frog neuromedin C porcine neuromedin C

pE Q R L G N QW A V G H L MN H 2 - G ------- T ----------------------- NH2 A P V S V G G G T V L A KM Y P R - - H -------------------- NH2 V - L P A G G -----------T - MY P R — H -------------------- NH2 ----------- G - Q ------------D - M Y P R — H -------------------- NH2 - - L Q P - - S P A - T - I Y P R - S H -------------------- NH2 ------- E N Q - S F P ♦ * - M F P R - S H ? -------- ? ? ? NH2 S E N T G A I - * * * * - V F P R - - H -------------------- NH2 - S H -------------------- NH2 — H -------------------- NH2

ranatensin ranatensin C ranatensin R porcine neuromedin B litorin glu(OMe)litorin glu(OEt)litorin leu8phyllolitorin phyllolitorin rohdeilitorin

p E V P ---------------- F p E Y P ------- T — F S N T A L - R Y ---------- T — F — L — T — F

NH2 NH2 NH2 - NH2

p E Q ---------------- F p E e ---------------- F p E e ---------------- F p E L ------------- S L p E L ------------- S F pEL — T — F

- NH2 - NH2 - NH2 - NH2 - NH2 - NH2

(-) Amino acids identical to those in the same position in bombesin or porcine GRP 1-13. (*) supposed deletions in fish GRPs. Four amino acids (?) in the C-terminal region of shark GRP have not been determined (Conlon et al., 1978b). (e) indicates O-methylation or O-ethylation of glutamine in litorin

has been determined. This peptide, consisting of 23 amino acids, is identical to mammalian GRP in the C-terminal dodecapeptide (Jensen and Conlon, 1992a). Similarly, a shark GRP, although not fully sequenced, shows a high degree of similarity to other GRPs in the C-terminal region (Conlon et al., 1987b). The combined results of radioimmunoassays, immunohistochemistry and pharmacological studies in the Atlantic cod, Gadus morhua, indicate the presence of more than one bombesin-like peptide with a regional distribution along the gut in this species (Holmgren and Jdnsson, 1988). The close similarity between different GRPs, and between GRPs and other bombesin-like peptides, in the common C-terminal part points to the importance of this region in the biological activity. GASTRIN/CCK-LIKE PEPTIDES Most studies of the occurrence of gastrin/CCK in non-mammalian species have concerned the presence in endocrine cells of the gut. However, recent immunohistochemical studies, mainly in our own laboratory, show the presence of gastrin/ CCK-like immunoreactivity in presumably autonomic nerves of several fish and amphibian species. This is in agreement with studies in mammals, which show the presence of CCK-containing nerves in the gut as well as in the central nervous system (Dockray and Hutchison, 1980; Dockray, 1977, 1980; Larsson and Rehfeld, 1977).

COMPARATIVE ASPECTS ON THE BIOCHEMICAL IDENTITY

77

TABLE 2.2 Amino acid sequences of gastrins, cholecystokinins and related peptides. CCK 39 CCK 33 CCK 8 gastrin 17 gastrin 34 caerulein phyllocaerulein

Y I Q Q A R K A P S G R V S M I K N L Q S L D P S HR I S D RD Y M G W M D F N H 2 --------------------------------------------------------------------------------------- NH2 --------------------------NH2 Q G P W L E E E E E A Y ------------ NH2 p E L G - Q - P P H L V A D P S KKQ - PWL E E E E E A Y ------------ NH2 pE Q - - T ------------- NH2 pE E - - T ------------- NH2

(-) Amino acids identical to those in the same position in cholecystokinin (CCK) 39.

It is possible that further studies will show a general distribution of such nerves amongst all vertebrates. Multiple forms of both gastrins and CCKs have been isolated from tissue extracts (Table 2.2). The gastrins are present as both sulphated and unsulphated variants, the CCKs are usually sulphated. The most obvious common feature of the gastrins, the CCKs and the closely related peptide caerulein, is the common C-terminal pentapeptide sequence; this pentapeptide is also important for the biological activity of the peptides. Therefore, it has been proposed that the gastrins, CCKs and caeruleins have a common ancestor. The gene encoding this common ancestor may have duplicated and formed daughter genes, subsequently evolving independently of each other into genes for gastrins and CCKs respectively (Barrington and Dockray, 1976; Larsson and Rehfeld, 1977). The enteric nerves in mammals have been shown to contain CCK8 rather than a gastrin-like peptide (Dockray, 1977). The first studies of non-mammalian vertebrates, performed in the two amphibians Rana temporaria and Rana esculenta and in the cod, Gadus morhua, led to the conclusion that the amphibian peptide caerulein is more ‘primitive’ than gastrin and CCK, and therefore likely to be the native gastrin/CCK-like peptide in amphibians and fish (Larsson and Rehfeld, 1977). However, subsequent studies indicate a more intricate situation. The gut of the two amphibians Xaenopus laevis and Rana temporaria contains both CCK-and caerulein-like peptides (Dimaline, 1983). In two teleosts, the cod and the rainbow trout, radioimmunological studies indicate the presence of several gastrin/CCK/ caerulein-like peptides in gut extracts: regional differences exist and it is even suggested that the occurrence of gastrin/CCK-like variants may be more extensive than in mammals (Vigna et al.y 1985; Jonsson, Holmgren and Holstein 1987). A similar situation exists in the elasmobranch Squalus acanthias (Aldman et al., 1989). Chicken gastrins show chemical properties similar to CCK, but act more like gastrins (Dimaline and Lee, 1990). Two studies in the protochordate, Ciona intestinalis, give the opposing results that the predominant gastrin/CCK-like immunoreactive material in the neural ganglion resembles C-terminal fragments of mammalian gastrin in one case (Thorndyke and Dockray, 1986) and is closely related to CCK8-S in the other case (Conlon, Schwartz and Rehfeld, 1988b). The hypothesis that ancestral gastrin/ CCK resembles CCK rather than gastrin (Vigna, 1985) is contradicted by studies

78

COMPARATIVE PHYSIOLOGY AND EVOLUTION

indicating that not only CCK-like, but also gastrin-like material is present in invertebrates (Nichols, Schneuwly and Dixon, 1988). Unfortunately, the studies of non-mammalian vertebrate species have not yet determined conclusively (e.g. by sequence analysis) which variants are present, or established their confinement to nerves or endocrine cells. The exceptions are caerulein and related peptides from the amphibian skin, such as phyllocaerulein and [Asn2-Leu5]-caerulein, which have been isolated and sequenced (Erspamer et al., 1984), but their distribution in nerves is not fully clarified. A property of importance for biological activity is the sulphation of the tyrosine residue in position 6 (gastrin) or 7 (CCK, caerulein), at the C-terminal. Whether the native peptides in non-mammalian species are sulphated or not is unknown, but pharmacological experiments demonstrate a much higher sensitivity to sulphated CCK8 (CCK8-S) than non-sulphated CCK8 in fish preparations. Thus, CCK8-S produces contractions of higher amplitude than CCK8 in cod stomach smooth muscle, although there is no difference in the concentration range for the effects of the two (Jonsson, Holmgren and Holstein, 1987). Gallbladder preparations from the rainbow trout are approximately a thousandfold more sensitive to CCK8-S than CCK8 (Aldman and Holmgren, 1987). In the rainbow trout gallbladder, and in the intestine and rectum of the spiny dogfish, Squalus acanthias, the position of the sulphated group appears to be of importance, CCK8-S being more efficient than gastrin 17 II, while gastrin and CCK are equipotent in the coho salmon Oncorhyfichus kisutch (Vigna and Gorbman, 1977; Aldman and Holmgren, 1987; Aldman et al., 1989). Biosynthesis o f gastrin/CCK-like peptides In mammals, the precursor of gastrin, preprogastrin, contains a single copy of G34, extended at both the C-terminal and the N-terminal end to a molecule of about 100 amino acids (depending on species). PreproCCK from rat similarly consists of 115 residues, including one copy of CCK-58, which can be cleaved to all known variants of CCK (Walsh, 1987). Both CCKs and gastrins are subject to post-translational processing such as amidation and sulphation after the proteolytic cleavage of the precursors (Walsh, 1987). Processing of non-mammalian gastrins may involve other types of cleavage than those in mammals. Thus a gastrin 36, a gastrin 30 and a gastrin 26 have been isolated from the chicken antrum, although the presence of a pair of basic residues at positions 12 and 13 in gastrin 36 would indicate tryptic cleavage at this point, rather than cleavage at the positions yielding gastrin 30 and gastrin 26. This cleavage has been proposed to be caused by the action of dipeptidyl aminopeptidase (Dimaline, Young and Gregory, 1986b; Dimaline, 1988). Three types of preprocaerulein cDNA have been found in the amphibian, Xenopus laevis. The most common form contains three copies of caerulein, the other forms contain one and four copies, respectively (Richter, Egger and Kreil 1986).

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79

NEUROPEPTIDE Y (NPY) AND RELATED PEPTIDES NPY is included in the family of peptides comprising the pancreatic polypeptides (PP; of pancreatic origin), the peptide YY (PYY) group originating from the gut, and the NPYs, originally considered exclusively neuronally contained. There are several immunohistochemical reports of the presence of an NPY-like peptide in cardiovascular and gastrointestinal nerves in non-mammalian vertebrates, and also a few reports on PYY immunoreactivity. Unfortunately, the peptides are similar enough to cross-react in immunohistochemistry. In many cases it is therefore impossible to speculate whether the reported immunoreactive material is indeed more closely related to NPY than PP or, especially, PYY. However, in a study in fish, the immunoreaction detected with an antiserum raised to mammalian NPY in perivascular nerves of the elasmobranch, Raja erinacea, and the intestine of the cod, Gadus morhua, is quenched by preincubation with NPY, but unaffected by pretreatment with PYY or PP (Bjenning, Driedzic and Holmgren, 1989). As will be discussed below, pancreatic polypeptides isolated from fish species also resemble mammalian NPY and PYY more than mammalian PP. NPY is a 36 amino acid peptide, first isolated from the porcine brain (Tatemoto, Carlquist and Mutt, 1982). At the same time the closely related PYY from the porcine gut was sequenced (Tatemoto, 1982). There is considerable homology especially in the C-terminal part between the amino acid sequences of NPY and PYY, and only slightly less homology with PP (Table 2.3). NPY appears to be one of the most highly conserved neuroendocrine peptides, maybe the most conserved considering its length (Table 2.3; Larhammar, Soderberg and Blomqvist, 1993). An amphibian NPY differing from porcine NPY in two positions, and from human and rat NPY in one position only, has been independently isolated from two frog species, Rana ridibunda and Rana temporaria (Chartrel et al., 1991; McKay et al., 1992). The peptide occurs exclusively in nerves within the brain, pancreas and gastrointestinal tract (McKay et al., 1992). Using clones encoding NPY, isolated from cDNA libraries of chicken, goldfish (Carassius auratus) and ray (Torpedo marmorata), Blomqvist et al. (1992) demonstrated the presence (in the brain) of true NPY in these species; the amino acid sequence differs from porcine NPY in only two, four and two positions respectively. In contrast to the high conservation of NPY between vertebrate species, there is a considerable variation in the sequence of the pancreatic polypeptides (Table 2.3). A few non-mammalian PPs have been sequenced; amongst these are three avian (chicken, turkey and goose), one reptilian (alligator) and one frog (Rana catesbeiana) peptide. The avian pancreatic polypeptides (APPs) are possibly more related to NPY than the mammalian PPs are, since they show a larger sequence homology with NPY, and often cross-react with NPY antisera in immunohistochemistry (Kimmel, Hayden and Pollock, 1975; Glover et al., 1985). The alligator and frog peptides are unique amongst the so far sequenced related peptides in having a C-terminal amidated phenylalanine instead of an amidated tyrosine (Lance et al., 1984; Pollock et al., 1988). A 37 amino acid peptide isolated from the pancreas of the anglerfish, Lophius americanas, differs from all hitherto sequenced related

(-), amino acids identical to those in the same position in porcine NPY

D — TP - QM- Q DQ-TPDQL-Q D G — V ------- I Q F D ------- V -------- I - F

- A A E - - R ------- M L - - P - - NH2 ------- D - Y Q - - T F -------- P - F NH2 - N D - Q Q - L - V V — P - F NH2 - D N - Q Q - L - V V - - H - - NH2

A-LE-VY A — E-HH T-LQ-KY G - - Q - T Y

porcine PP frog PP alligator PP avian PP (chicken)

— — — - -

- - A - - E A ------------- S P - E - S ---------A S ----------- L - - V ---------------NH2

PYY porcine

P — E ----------------- S P - E Q - K — T ------------------------------------------ NH2 P - - E --------------------P - E - - K - - T ------------------------------------------- NH2 P — E -------------------- P - E ------------------------------------------------------------- NH2 P — E -------------------- P - E — K --------------------------------------------------- NH2 P — E -------------------- P - E — K --------------------------------------------------- NH2 P — E -------- D - - A P - E - - K -------------------------------------------------- NH2 P Q - E S — G N - S P — W - K - H A - V --------V ----------------------- NH2 P — E T — S N - S P — W - S - Q A - V --------V ----------------------- G - P ------------- S P — S P - E - S K - M L - V - N ------------------------------NH2

Y P S K P D N P G E D A P A E D L A R Y Y S A L R H Y I NL I T R Q R YNH2 ---------------------------------------------- M -------------------------------------------------------- NH2 ---------------- S ---------------------------M -------------------------------------------------------- NH2 ---------------------------------------------- M - K -------------------------------------------------- NH2 - - T ------------------- G ----------- E - - K ---------------------------------------------------NH2 ---------------------------- G -------------------- K ---------------------------------------------------NH2 — - — - — — — — M

(porcine) (human) (chicken) (frog) (goldfish) (Torpedo)

Fish peptides o f pancreatic origin eel salmon bowfin gar dogfish skate sculpin PP anglerfish APY lamprey

NPY NPY NPY NPY NPY NPY

TABLE 2.3 Amino acid sequences of neuropeptide Y (NPY), peptide YY (PYY) and pancreatic polypeptide (PP) from different species.

80 COMPARATIVE PHYSIOLOGY AND EVOLUTION

COMPARATIVE ASPECTS ON THE BIOCHEMICAL IDENTITY

81

peptides in having a C-terminal tyrosine-glycine where all the others have an amidated tyrosine (or phenylalanine in the alligator and frog). The peptide resembles NPY and PYY more than PP, and was named Anglerfish Peptide YG (APY or aPY) (Andrews et al., 1985; Noe et al., 1986b). Most fish pancreatic polypeptides similarly resemble NPY more than PP from other vertebrates, but in contrast to APY possess the C-terminal amidated tyrosine characteristic of the main group (Table 2.3; Kimmel et al., 1986; Conlon et al., 1986b, 1991 a, b, c; Cutfield, Carne and Cutfield, 1987; Pollock et al., 1987). Furthermore, these peptides show some ‘substitutions’ characteristic of PYY. Interestingly, studies in anglerfish indicate the presence of a C-terminally amidated PP in addition to APY, and it has been suggested that fish pancreas may contain a mixture of the two peptides, processed slightly differently (Andrews and Dixon, 1985). There are thus great similarities between the NPY-related peptides from fish, while amongst frogs, there are now reports of the presence of true NPYs (isolated from the brain), PYY isolated from the intestine and PP isolated from the pancreas (Table 2.3; Pollock et al., 1988; Chartrel et al., 1991; Conlon, Chartrel and Vaudry, 1992; McKay et al., 1992). This opens interesting possibilities regarding the development of this peptide group (Conlon, Chartrel and Vaudry, 1992; Larhammar, Soderberg and Blomqvist, 1993). Biosynthesis o f NPY-like peptides The structural similarity between NPY, PYY and PP indicates that they all originally come from the same gene, which has undergone successive duplications during evolution. The strong homology of fish pancreatic polypeptides to mammalian NPY and PYY has led to the suggestion that the gene mutations and duplications took place after the time of divergence of the lines of evolution leading to fish and mammals, respectively. The fish peptides are possibly more similar to the common ancestral peptide (Conlon, 1989). However, true NPY as well as NPY-related pancreatic peptides are identified in fish, and the point of divergence may be even earlier (Larhammar, Soderberg and Blomqvist, 1993). Furthermore, the recent demonstrations of true NPY in fish species indicate a strong conservation of this particular peptide, while the PP-and PYY-like peptides have been more susceptible to permutations (Blomqvist et al., 1992; Larhammar, Soderberg and Blomqvist, 1993). THE NEUROTENSIN FAMILY OF PEPTIDES The classical view of neurotensin in mammals, is that of a transmitter in the central nervous system, with peripheral neurotensin-like substances contained in intestinal endocrine cells. However, there are now several reports on the presence of neurotensin in peripheral nerves of the intestine, the cardiovascular system, the pancreas and the kidney. Similarly, there are several immunohistochemical studies reporting the presence of neurotensin-like material in peripheral nerves of non-mammalian species (Carraway and Reinecke, 1989). Radioimmunological studies have demonstrated the presence of neurotensin-like peptides in several vertebrates (and invertebrates). The use of various region-specific

82

COMPARATIVE PHYSIOLOGY AND EVOLUTION TABLE 2.4 Amino acid sequences of neurotensin and related peptides.

Neurotensin (bovine, canine, human) Neurotensin (guinea pig) Neurotensin (chicken) Xenopsin (canine, turkey) Xenopsin (amphibian) LANT-6 (chicken) Neuromedin N (porcine)

pELYENKPRRPY I L -------------- S ---------------- - H V - - A -------------FHPK- -W- pEGK- - W - K N --------K I ---------

Neurotensin-related peptide (NRP), plasma Neurotensin-related peptide (NRP), turkey skin

I A R - H - - FRT-G- -F -

LANT-6, lys8-asn9-neurotensin (8-13) (-), amino acids identical to those in the same position in bovine neurotensin

antisera in a series of representatives of different vertebrate and invertebrate groups has shown a preferential conservation of the C-terminal part of the molecule. This is also the biologically most important part of the neurotensins (Carraway et al., 1982b; Carraway and Reinecke, 1989). Neurotensin-like peptides appear to be present in all vertebrate groups (Reinecke et al., 1980; Reinecke, 1981; Carraway et al., 1982 a, b). The two neurotensin-like peptides isolated from chicken differ very little in amino acid sequence from the mammalian counterparts (Table 2.4). One is a tridecapeptide similar to mammalian neurotensin, except for substitutions in positions 3, 4 and 7 (Carraway and Bhatnagar, 1980). The substitutions are all in the N-terminal half of the peptide and have little effect on the biological properties. The other is a hexadecapeptide, [Lys8-Asn9]-neurotensin 8-13 (LANT 6 ), similar to neuromedin N but for a single substitution of isoleucin for asparagine in position 2 (Carraway and Ferris, 1983). Xenopsin is an octapeptide isolated from the skin of Xenopus, which shares four of the five C-terminal residues with neurotensin (Araki et al., 1973; Table 2.4). Xenopsin-like as well as neurotensin-like material is present in peripheral tissues of Xenopus and other amphibians (Carraway et al., 1982a; Goedert et al., 1984). Information on precursors and their processing is available only for mammalian species. OPIOID PEPTIDES The opioid peptides form three major groups, the enkephalins, the endorphins and the dynorphins. The pentapeptide sequence of met-enkephalin or leu-enkephalin is present in all endogenous mammalian opioid peptides: e.g. /3-endorphin is metenkephalin with a carboxyl extension of 26 amino acids, and /3-neo-endorphin and dynorphin are leu-enkephalin with carboxyl extensions of 4 and 12 amino acids, respectively. Studies of genes and precursors, isolations and immunocytochemical studies indicate that opioid substances have been used by living organisms from a very early evolutionary stage, but the relative proportions of the two enkephalins or their extended molecules differ between species. Met-enkephalins but not leu-enkephalins are found in the amphibian Xenopus laevis (Martens and Herbert, 1984), while both

COMPARATIVE ASPECTS ON THE BIOCHEMICAL IDENTITY

83

met- and leu-enkephalins are reported in Bufo marinus (Kilpatrick et al., 1983). Although all three groups of opioid peptides - the enkephalins, the endorphins and the dynorphins - contain the sequence of leu-or met-enkephalin, they stem from three different precursors. Enkephalins are synthesized from preproenkephalin A. Thus bovine adrenal preproenkephalin A contains four copies of Met-enkephalin and one copy each of Leu-enkephalin, Met-enkephalin-[Arg6-Phe7] and Metenkephalin-[Arg6-Gly7-Leu8]. Preproenkephalin B (from porcine hypothalamus) is processed into /3-neo-endorphin, dynorphin and a Leu-enkephalin with a C-terminal extension. Finally, the endorphins are processed from pro-opiocortin, via ¡3lipotrophin (Corder and Rees, 1981; Kakidani et al., 1982; Rossier, 1982). The preproenkephalin A sequence of reptiles and anuran amphibians differs from the mammalian counterpart in that the Met-enkephalin-[Arg6-Gly7-Leu8] is missing (Lindberg and White, 1986; Martens and Herbert, 1984; Kilpatrick et al., 1983). The amphibian precursor instead contains Met-enkephalin- [Arg6-Gly7-Tyr8], and furthermore five Met-enkephalins rather than four Met-enkephalins and one Leuenkephalin as in mammals, all due to only three base pair substitutions (Martens and Herbert, 1984; Noda et al., 1982). SOMATOSTATIN The somatostatins occur in brain, endocrine cells and in pancreatic tissues in representatives from all vertebrate groups. A number of somatostatin-like peptides have been isolated from non-mammalian vertebrates (Table 2.5): catfish somatostatin which occurs in several forms, salmon somatostatin-25 and anglerfish and sculpin somatostatin-28 are longer forms found in teleosts (Andrews et al., 1984; Morel et al., 1984; Spiess and Noe, 1985; Plisetskaya et al., 1986; Cutfield, Carne and Cutfield, 1987). A shorter form, somatostatin 14, appears to be extremely well preserved during evolution: an identical sequence of the 14 amino acids is reported from the elasmobranch Torpedo marmorata, catfish (Ictalurus sp), anglerfish (.Lophius), pigeon and mammals (Noe et al., 1979; Spiess et al., 1979; Andrews and Dixon 1981; Minth et al., 1982; Conlon, Agoston and Thim, 1985a). The presence in peripheral nerves in nonmammalian vertebrates has been demonstrated by immunohistochemistry in some TABLE 2.5 Amino acid sequences of somatostatins and urotensins. porcine S28 S-14 anglerfish S28 I anglerfish S28 II salmon S25 catfish S22 urotensin carp urotensin II

S A N S N P A M A P R E R K A G C K N F F WKTFT S C A - S G G - L L --------------------------------------------------- V D - T N N L P --------------------------- Y - - G ---------- V D N L P --------------------------- Y - - G ---------D N T V T S K P L N - M - Y ------ S R - A - - T A D C ------ YCV G - G A D C ------ YCV

(-), amino acids identical to those in the same position in porcine somatostatin-28

84 COMPARATIVE PHYSIOLOGY AND EVOLUTION

species, but although several somatostatins of non-mammalian origin have been sequenced, it is not clear which forms are contained in the nerves. However, in the toad, Bufo marinus, the somatostatin-like material from the heart, shown by immunohistochemistry to be contained in nerves, co-elutes with somatostatin 14 (Campbell et al., 1982). The urotensins II from the caudal neurosecretory system in teleosts show partial sequence similarities as well as functional similarities with the somatostatins (Pearson et al., 1980; Ichikawa, 1985). Biosynthesis o f somatostatin In humans and rat a single gene encodes preprosomatostatin. In teleost fish, such as anglerfish, sculpin or salmon, two genes are present. Gene I codes for preprosomatostatin I, while gene II codes for preprosomatostatin II. There is no evidence so far for expression of gene II in any group other than the teleost fish (Conlon, 1989). The subsequent processing of the prosomatostatins in teleost fish follows different pathways. Prosomatostatin I from anglerfish, catfish, and rat have been studied, and are found to be highly conserved in structure (Argos et al., 1983). The prosomatostatins from anglerfish, flounder and daddy sculpin are cleaved as in mammals at an Arg-Lys site to give somatostatin-14 (Noe et al., 1986a; Conlon et al., 1987a). The structure of the prosomatostatin II molecule is similarly well conserved between teleost species (Conlon et al., 1987a), but the further processing to the active substance differs giving somatostatins of different length in different species (Andrews et al., 1984; Morel et al., 1984; Spiess and Noe, 1985; Conlon et al., 1987a; see also above). In mammals, cleavage of preprosomatostatin is tissue-specific, giving different proportions of somatostatin-14 and somatostatin-28 in different tissues. In an elasmobranch, Torpedo marmorata, only somatostatin-14-like immunoreactivity is present in extracts from the brain, stomach, pancreas or gut, suggesting that this is the only form expressed in this species (Conlon, Agoston and Thim, 1985a; Conlon, 1989). An entirely different way of processing somatostatin has been suggested for the cyclostomes. A hagfish somatostatin-34 has been isolated, and the structure of the N-terminal region shows little in common with other vertebrate somatostatins. However, a somatostatin-14 identical to somatostatin-14 from other species is also present, further stressing the high degree of conservation of this particular somatostatin (Conlon, 1989). TACHYKININS Like the bombesins, most of the tachykinins sequenced so far were originally isolated from amphibian skin. However, immunohistochemistry has shown that tachykinins frequently occur in both central and peripheral nerves of vertebrates and invertebrates. It is clear that the tachykinins, as a group, are phylogenetically

COMPARATIVE ASPECTS ON THE BIOCHEMICAL IDENTITY

85

TABLE 2.6 Amino acid sequences of tachykinins. Substance P (mammal) Arg3-substance P (bird) Substance P (rainbow trout) Substance P (Atlantic cod) Physalaemin (amphibian skin) Lys5-Thr6-physalaemin (amphibian) Scyliorhinin I (elasmobranch) Uperolein Eledoisin (mollusc) Phyllomedusin Neuropeptide gamma Neurokinin A (mammal, bird) Neurokinin A (rainbow trout) Neurokinin A (Atlantic cod) Carassin (teleost) Neurokinin B (mammal) Kassinin Glu2-Pro5-kassinin Entero-kassinin Scyliorhinin II (elasmobranch) Hylambatin Entero-hylambatin

R P K P Q Q F FGLMNHj ——R ----------------------- NH2 K - R - H ----------------- NH2 K - R ----------I ---------- NH2 pEAD-NK-Y pEAD-KT-Y A -F D K -Y p E -D -N A -Y p E -S K D A -I pE N - N R - I D A G H G Q I S H K R H - TD S H - TD s H- IN s H- IN s S P ANAQI T RK - H - I N s DMHD F DV — SD DE — - D DE - N SD S P S N s K C P DG - D c

■- V -- V ■- V -- V -- V -V -- V -- V -- I -- V

-------- NH2 -------- NH2 -------NH2 -------NH2 -------NH2 -------NH2 -

-----------

-

NH NR NR NR NR NH: NH; NH; NH: NH:

D P - D - D R - T - M - NH2 D P - N - D R - T - M - NH2

(-), amino acids identical to those in the same position in mammalian substance P

very old. Immunocytochemical studies often indicate the presence of one or several tachykinins in innervated tissues. The common biochemical feature of the tachykinins (except hylambatins) is the C-terminal amino acid sequence Phe-X-Gly-Leu-Met-NH2 (Table 2.6). Substance P was the first tachykinin isolated and sequenced in mammals, and was long considered the only mammalian tachykinin. However, two other tachykinins, neurokinin A (neurokinin a, substance K or neuromedin L) and neurokinin B (neurokinin /3, neuromedin K) have been isolated from the porcine spinal cord (Kangawa et al., 1983; Kimura et al.9 1983), and more recently several bioactive neurokinin A-derived peptides, including neuropeptide K (NPK), neuropeptide gamma (NP7 ) and NKA(3-10) have been discovered (see Helke et al.9 1990). Two tachykinins have been isolated from the elasmobranch Scyliorhinus caniculus (Conlon et al.9 1986a). Scyliorhinin I is a decapeptide, identical to physalaemin in the C-terminal hexapeptide and cross-reacting with antisera directed towards the C-terminal of substance P. Scyliorhinin II is an octadecapeptide with a sequence that is related to the amphibian skin peptide [Glu2-Pro5]-kassinin in its C-terminal part. Scyliorhinin II cross-reacts with antisera directed towards the C-terminal of neurokinin A (Conlon et al.9 1986a). In a receptor binding study it was found that scyliorhinin I binds with higher affinity to NK1/NK2 (SP/NKA) receptors, while scyliorhinin II binds with higher affinity to NK3 (NKB) receptors (Buck and Krstenansky, 1987).

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Neither neurokinin A nor substance P could be demonstrated in Scyliorhinus (Conlon et al., 1986a). In accordance, tachykinin extracts of the central nervous system of the skate Raja batis does not displace the tracer in RIA in parallel with substance P, indicating a different identity (Creagh et al., 1980). A 16 amino acid tachykinin identical to scyliorhinin II (3-18) was isolated from the intestine of the ray, Torpedo marmorata (Conlon and Thim, 1988). Rays and dogfish are earlydeveloped groups and the close similarity, (almost identical), between the corresponding Scyliorhinus and Torpedo peptides indicates the strong evolutionary pressure to conserve this peptide sequence. Isolation of tachykinin-like material from the gastrointestinal tract of whiting, Merlangius merlangus, and carp, Cyprinus carpio, have indicated the presence in teleosts of several different tachykinins (Maule et al., 1989: Kitazawa, Kudo and Ishigami, 1990). None of these appear to be identical to mammalian substance P, but the recent isolation and structural characterization of tachykinins from Atlantic cod and rainbow trout brain have yielded peptides with high structural similarities to substance P (Table 2.6). There are only three amino acid substitutions compared to the mammalian counterpart, but the substance P-related peptide from cod has an unusual substitution in the C-terminal pentapeptide (isoleucine instead of phenylalanine in position 8 ). In addition to the substance P-related peptides, both species have a neurokinin A-related peptide very similar to mammalian neurokinin A (Jensen and Conlon, 1992b). The tachykinins found in the brain of rainbow trout are also present in the intestine of the same species (Jensen, Olson and Conlon, 1993). Another tachykinin, carassin, isolated from the brain of goldfish, Carassius auratus, seems to be related to the mammalian neuropeptide gamma, and has a Cterminal decapeptide identical to rainbow trout and cod neurokinin A (Conlon et al., 1991d). Studies in amphibians generally indicate the presence of more than one tachykinin in extracts of the brain and gut. One of these peptides often shows a close relationship to substance P (Lembeck et al., 1985; Conlon, Ballmann and Lamberts, 1985b; Bowers, Jan and Jan, 1986; Creagh et al., 1980). In the axolotl, Ambystoma mexicanum, one of the peptides in stomach extracts shows characteristics resembling physalaemin (Conlon, Ballmann and Lamberts, 1985b). In the gut of three hylid frog species (Phyllomedusa SPP) and in Kassina senegalensis, kassinin-like immunoreactivity is dominant, while in other species physalaemin-like immunoreactivity is the most obvious besides substance P (Melchiorri and Negri, 1984). In intestinal extracts from a turtle, Testudo kleinmanni, and a lizard, Lacerta 5., several peaks of tachykinin-like material were present, including one eluting close to synthetic substance P (Lembeck et al., 1985). Two tachykinins from the chicken gut have been isolated and sequenced. One is [Arg3]-substance P, which differs only in one position from mammalian substance P, and the other is identical to the mammalian neurokinin A (Conlon et al., 1988a). A third peptide was present in the extracts, but was not structurally characterized. Extracts from the pigeon intestine showed the presence of a tachykinin not resembling substance P (Creagh et al., 1980). The tachykinins are probably the best known neuropeptides from a comparative

COMPARATIVE ASPECTS ON THE BIOCHEMICAL IDENTITY

87

point of view, with several representatives isolated from most vertebrate groups allowing comparative studies. It is striking how well conserved the C-terminal sequence is throughout the animal kingdom - this part of the molecule must be of crucial importance for the physiological actions of the tachykinins. There are no reports on the biosynthesis of tachykinins in non-mammalian species. VASOACTIVE INTESTINAL POLYPEPTIDE (VIP) Reports on VIP-like immunoreactivity in nerves of non-mammalian vertebrates (Nilsson and Holmgren, 1989; Dimaline, 1989; other chapters in this book) are common, and information on the actual sequences of non-mammalian VIP or VIPlike peptides is steadily increasing (Table 2.7). The sequences are amazingly similar, with identical N-terminal regions and only few substitutions in the C-terminal region. VIP from chicken has been sequenced, and differs from porcine VIP in four locations (Nilsson 1975). Helodermin (35 residues) and helospectins (37 and 38 residues), isolated from the Gila monsters, Heloderma suspectum and Heloderma horridum, share 15 residues with VIP and may be the reptilian representatives of VIP (Hoshino et al., 1984; Parker et al., 1984; Vandermeers et al., 1987). A twenty-eight amino acid peptide with a sequence differing from porcine VIP at only five residues has been isolated from the intestine of the elasmobranch Scyliorhinus canicula (Dimaline, Thorndyke and Young, 1986a, 1987). Similarly, VIP from the intestine of the cod, Gadus morhua, has been sequenced, and was found to be identical to avian VIP with the exception of substitutions at position 19 and 28 (Thwaites et al., 1989). Biosynthesis Mammalian preproVIP contains the sequence of VIP and that of the structurally related peptide, Peptide Histidine Isoleucin (PHI) (Itoh et al., 1983; Nishizawa et al., 1985). No non-mammalian VIP precursors have been identified, but it has been suggested that PHI is present in elasmobranch preproVIP on the basis that VIP-containing nerves in Raja radiata and Raja clavata also store a PHI-like peptide (Ekblad et al., 1985).

TABLE 2.7 Amino acid sequences of vasoactive intestinal polypeptide (VIP) and VIP-like peptides. porcine VIP avian VIP dogfish VIP cod VIP

H S D A V F T D N Y T R L R K Q M A V K K Y L N S I L N NH2

helodermin helospectin

- - I — E E - S K - L A K L - L Q -------A -------- G S — T — A E - S K - L A K L - L Q -------E -------- G S

S - I — S - F ------------- A —

(-), amino acids identical to those in the same position in porcine VIP

- - L - A NH2 - - V - A NH2

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COMPARATIVE PHYSIOLOGY AND EVOLUTION

OTHER ‘VERTEBRATE PEPTIDES’ Some more-or-less recently discovered neuropeptides have been structurally described in mammals only. Although immunohistochemical, radioimmunochemical and to some extent physiological and pharmacological studies indicate their presence and function in non-mammalian species, virtually nothing is know yet about their structure and degree of conservation. Amongst these peptides are calcitonin generelated peptide (CGRP), galanin and oxytocin. A pentapeptide with the sequence H-Gly-Phe-Trp-Asn-Lys-OH, termed eel intestinal peptide (EIPP) has been isolated from the eel gut. The peptide is stimulatory on eel gut motility (Uesaka et al., 1991).

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Conlon, J.M., Davis, M.S., Falkmer, S. and Thim, L. (1987a). Structural characterization of peptides derived from prosomatostatins I and II isolated from the pancreatic islets of two species of teleostean: the daddy sculpin and the flounder. Eur. J. Biochem., 168, 647-652. Conlon, J.W, Henderson, I.W. and Thim, L. (1987b). Gastrin-releasing peptide from the intestine of the elasmobranch fish, Scyliorhinus canicula (common dogfish). Gen. Comp. Endocrinol., 68, 415-420. Conlon, J.M., Katsoulis, S., Schmidt, W.E. and Thim, L. (1988a). Arg3-subtance P and neurokinin A from chicken small intestine. Regul. Pept., 20, 171-180. Conlon, J.M., Schwartz, T.W. and Rehfeld, J.F. (1988b). Sulphated CCK-8-like peptides in the neural ganglion of the protochordate Ciona intestinalis. Regul. Pept., 20, 241-250. Conlon, J.M., Balasubramaniam, A. and Hazon, N. (1991a). Structural characterization and biological activity of a neuropeptide Y-related peptide from the dogfish, Scyliorhinus canicula. Endocrinology, 128, 2273-2279. Conlon, J.M., Bjenning, C., Moon, T.W., Youson, J.H. and Thim, L. (1991b). Neuropeptide Y-related peptides from the pancreas of teleostean (eel), holostean (bowfin) and elasmobranch (skate) fish. Peptides, 12, 221-226. Conlon, J.M., Bjôrnholm, B., Jorgensen, F.S., Youson, J.H. and Schwartz, T.W. (1991c). Primary structure and conformational analysis of peptide methioninetyrosine, a peptide related to neuropeptide Y and peptide YY isolated from lamprey intestine. Eur. J. Biochem., 199, 293-298. Conlon, J.M., O’Harte, F., Peter, R.E. and Kah, O. (1991d) Carassin -a tachykinin that is structurally related to neuropeptide-gamma from the brain of the goldfish. J. Neurochem., 56, 1432-1436. Conlon, J.M., O’Harte, F. and Vaudry, H. (1991e). Primary structure of the bombesin-like neuropeptides in frog brain show that bombesin is not the amphibian gastrin-releasing peptide. Biochem. Biophys. Res. Comm., 178, 526-530. Conlon, J.M., Chartrel, N. and Vaudry, H. (1992). Primary structure of frog PYY: Implications for the molecular evolution of the pancreatic polypeptide family. Peptides, 13, 145-149. Corder, R. and Rees, L.H. (1981) Endorphins and enkephalins. In Gut Hormones, edited by S.R. Bloom and J.M. Polak, pp. 228-239. Edinburgh, London, Melbourne, New York: Churchill Livingstone. Costa, M., Furness, J.B. and Humphreys, C.M.S. (1986). Apamin distinguishes two types of relaxation mediated by enteric nerves in the guinea-pig gastrointestinal tract. Naunyn-Schmiedebergs Arch. Pharmacol., 332, 79-88. Creagh, T., Skrabanek, P., Cannom, D., Balfe, A. and Powell, D. (1980). Phylogeny of substance P. Gen. Comp. Endocrinol., 40, 503-506. Cutfield, S.M., Came, A. and Cutfield, J.F. (1987). The amino-acid sequences of sculpin islet somatostatin-28 and peptide YY. FEBS Lett., 214, 57-61. Dimaline, R. (1983). Is caerulein amphibian CCK? Peptides, 4, 457-462. Dimaline, R. (1988). Post-translational modification of peptide messengers in the gut. Q. J. Exp. Physiol., 73, 873-902. Dimaline, R. (1989). Vasoactive intestinal peptide. In The Comparative Physiology o f Regulatory Peptides, edited by S. Holmgren, pp. 150-173. London: Chapman and Hall. Dimaline, R. and Lee, C.M. (1990). Chicken gastrin: a member of the gastrin/CCK family with novel structure-activity relationships. Am. J. Physiol., 259, G882-G888. Dimaline, R., Thorndyke, M.C. and Young, J. (1986a). Isolation and partial sequence of elasmobranch VIP. Regul. Pept., 14, 1-10. Dimaline, R., Young, J. and Gregory, H. (1986b). Isolation from chicken antrum, and primary amino acid sequence of a novel 36-residue peptide from the gastrin/CCK family. FEBS L ett., 205, 318-322. Dimaline, R:, Young, J., Thwaites, D.T., Lee, C.M., Shuttleworth, T.J. and Thorndyke, M.C. (1987). A novel vasoactive intestinal peptide (VIP) from elasmobranch intestine has full affinity for mammalian pancreatic VIP receptors. Biochem. Biophys. Acta, 930, 97-100. Dockray, G.J. (1977). Immunoreactive component resembling cholecystokinin octapeptide in intestine. Nature, 270, 359-361. Dockray, G.J. (1980). Cholecystokinin in rat cerebral cortex: identification, purification and characterization by immunochemical methods. Brain Res., 188, 155-165. Dockray, G.J. and Hutchison, J.B. (1980). Cholecystokinin octapeptide in guinea-pig ileum myenteric plexus: localization and biological action. J. Physiol. (Lond), 300, 28P-29P. Du, B.H., Eng, J., Humes, J.D., Chang, M., Pan, Y.C.-. and Yalow, R.S. (1985). Guinea-pig has a unique mammalian VIP. Biochem. Biophys. Res. Commun., 128, 1093-1098. Ekblad, E., Alumets, J., Ekman, R., Falkmer, S., Hâkanson, R. and Sundler, F. (1985). Two distinct VIP precursors in cartilaginous fish? Peptides, 6 (Suppl 3), 383-388.

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3 Chromaffin Systems Robert M. Santer Department o f Anatomy, University o f Wales College o f Cardiff, UK The term ‘chromaffin* was originally applied to cells of the adrenal medulla which could be stained with chromaffin salts on account of their high catecholamine content. Release of these catecholamines, often as a result of stress, results in elevated plasma concentrations. The contemporary usage of the term ‘chromaffin* extends to cells with similar morphological and histochemical properties whether or not they demonstrate a chromaffin reaction. As well as adrenomedullary chromaffin cells, extra-adrenal chromaffin tissue also exists throughout the vertebrates principally in relation to elements of the autonomic nervous system, the cardiovascular system and in the retro-peritoneum. This chapter recounts the development of our knowledge of the distribution and biology of chromaffin systems in vertebrates with particular emphasis on the adrenomedullary cells, the ‘small* cells of autonomic ganglia and other, dispersed chromaffin elements, with the object of relating their known structural and histochemical properties to their probable functions. KEY WORDS chromaffin cells; adrenomedulla; SIF cells

INTRODUCTION In the mid-nineteenth century Joeston (1864) and Henle (1865) both observed that fixation of the adrenal gland with compounds of chromium resulted in the adrenal medulla staining brown whilst the adrenal cortex remained unstained. It was not until 1902 that the term “chromaffin” was used by Kohn to describe cells that exhibited such coloration following treatment with chromate compounds. The introduction of the word ‘paraganglion’ by Kohn (1903) acknowledged the existence of extra-adrenal chromaffin tissue in the light of the earlier demonstration of a chromaffin reaction in the carotid body (Stilling, 1889). Ogata and Ogata (1923) suggested that the chromaffin reaction could be related to the occurrence of adrenaline in adrenal glands, but the production of a similar coloration in the adrenal medulla after fixation with potassium iodide (Gerard, Cordier and Lison, 1930) threw doubt on this contention. It is now known (Hopwood, 1971) that the production of brown pigments in adrenal medullary cells is due to the oxidation 97

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of the cellular catecholamines adrenaline (A) and noradrenaline (NA), usually by potassium dichromate treatment, to insoluble ring-structured adrenochromes and noradrenochromes. We also know that cells of the ‘chromaffin systems’, the chromaffin cells, do not always show a classical chromaffin reaction as well as not being confined to the adrenal medulla. The term ‘chromaffin systems’ is used nowadays to describe collections of variously located cells in vertebrates which are known to contain high concentrations of catecholamines and/or indolealkylamines such as 5-hydroxytryptamine (5-HT) and which have a fine structural appearance similar to that of adrenal medullary cells, particularly with regard to their content of storage granules. Certain characteristics of chromaffin cells are generally acknowledged: 1) they are derived embryologically from the neural crest from which they migrate to their final destination. 2 ) they have a preganglionic sympathetic innervation which makes cholinergic-type synaptic contacts with the plasma membrane. 3) they contain a multitude of rounded, oval or elongated dense-cored granules of predominantly 100-300 nm diameter but with cores of varying ultrastructural appearance: NAstoring granules have highly electron-dense cores which are irregular in shape whereas A-storing granules are paler and more homogeneous (Figures 3.1 and 3.2). 4) the catecholamines are co-stored with many proteins such as ATPase, dopamine0-hydroxylase and the acidic protein chromogranin A which is the most abundant of the chromogranin family. The adrenal chromogranins appear to have the same molecular properties in all vertebrate classes (Rieker et al., 1988). 5) a variety of neuropeptides are also co-stored within the storage granules. 6 ) chromaffin cells are usually encountered in groups or glomera with an intimate, endocrine-like relationship to fenestrated capillaries but solitary chromaffin cells also exist. 7) the storage granules release their entire contents into the extracellular space by exocytosis which is calcium-dependent. The classical effect of the stress-or stimulus-induced release of catecholamines by exocytosis from the adrenal medulla is to elevate greatly the catecholamine levels in the bloodstream with a consequent increase in heart rate and blood pressure and to stimulate glycogenolysis in liver and skeletal muscle; other effects of catecholamine release from the adrenal medulla include increased metabolic rate, increases in free fatty acids, blood lactate, insulin and glucagon levels. The concomitant release of proteins and peptides into the bloodstream means that the physiological consequences of adrenal medullary secretion are more complex that originally appreciated. The precise functional roles of, and the demonstration of release by exocytosis from extra-adrenal chromaffin cell, is far less well understood and documented than in adrenomedullary chromaffin cells. Among the criteria used by Coupland (1965; 1972) to define chromaffin cells was the fact that they stored sufficient quantities of catecholamine to produce a chromaffin reaction. However, as implied above, there are cell types generally accepted as being members of the chromaffin family in which a classical chromaffin reaction is not detectable. Such cell types are called chromaffin cells on account of other morphological or histochemical properties. In addition to a sufficient quantity of catecholamine being present, the size and subcellular distribution of the cate-

CHROMAFFIN SYSTEMS

FIGURE 3.1 Rat adrenal medulla. Cytoplasm o f a NA-storing cell showing the typical heterogeneous form o f the storage granules with their highly electron-dense cores. Calibration bar 1 /¿m.

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FIGURE 3.2 Rat adrenal medulla. Cytoplasm o f an A-storing cell containing rounded storage granules with cores o f less electron-density than seen in N A granules, Calibration bar 1 fxm.

cholamine storage granules is a critical factor in the production of the classical chromaffin reaction. However, the tissue preservation after direct chromate treatment is often unsatisfactory as it becomes brittle and difficult to section: it is also unsuitable for electron microscopy and does not differentiate between separate NAand A-storing cells in the adrenal medulla. Hillarp and Hokfelt (1953) had demonstrated, by the use of oxidation with potassium iodate, that the two amines were stored in separate cell types in the adrenal medulla, but it was not until the development of the method of fixation with glutaraldehyde (Coupland, Pyper and Hopwood, 1964; Wood and Barrnett, 1964) that the two medullary cell types could be distinguished from one another at the electron microscopical level. The reaction of glutaraldehyde with NA results in the formation of an extremely stable Schiff monobase (Tramezzani, Chiocchio and Wasserman, 1964) which will bind chromium (Nemes, 1974) and produces a strong yellow coloration in adrenal medullary and other NA cells. Other primary catecholamines such as dopamine (DA) and its precursor Dihydroxyphenylalanine (DOPA) and indolealkylamines such as 5-HT react similarly (Hopwood, 1971; Rose, Lever and Santer, 1978).

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Whether secondary catecholamines such as A form an unstable compound with glutaraldehyde or are simply eluted during tissue processing (Coupland and Hopwood, 1966) has not been determined and no reaction product has ever been recorded in A cells. This differential reaction has been confirmed ultrastructurally by quantitative X-ray microanalysis where elemental chromium can be localized not only to the highly electron dense granules characteristic of adrenal medullary NA cells (Wood, 1974; Santer, Lever and Davies, 1978) but in other chromaffin cell types (Lever et al., 1977) and even in the small dense cored vesicles of noradrenergic axon terminals (Santer, Lever and Davies, 1981). Currently the glutaraldehydepotassium dichromate methodology is mainly used in the mapping and subsequent electron microscopy of extra-adrenal chromaffin tissue (Mascorro and Yates, 1977; 1987). Many techniques are in use for studying the location, structure and content of chromaffin cells. The discovery by Erànko (1955) of fluorescing islets of cells within the adrenal medulla after formalin fixation was the vital observation that lead to the development of the formaldehyde-induced fluorescence (FIF) technique for biogenic monoamines in Scandinavia in the early 1960s. By using this technique new extra-adrenal chromaffin cell types were discovered, particularly the small intensely fluorescent (SIF) cells of sympathetic ganglia (Erànko and Hârkônen, 1965), and their distributions mapped. The development of indirect immunofluorescence histochemistry. (Coons, 1958) and the preparation of antibodies to the enzymes of the biosynthetic pathway for catecholamines has produced a highly sensitive method (Hokfelt et al., 1973) for visualising chromaffin cells. The importance of immunohistochemical methodology to our current understanding of chromaffin tissue cannot be understated, particularly with regard to the neuropeptides, and will become evident later in this chapter. In addition, scanning and transmission electron microscopy, enzyme histochemistry and in vitro methods are all currently employed in studies on chromaffin cells.

DISTRIBUTION OF CHROM AFFIN TISSUE IN VERTEBRATES With the exception of the solitary type of SIF cell in autonomic ganglia, chromaffin tissue is found closely related to the vascular system throughout the vertebrates. The most extreme examples of this are the mammalian carotid body which is probably the most highly vascularised organ known, and the widespread distribution of chromaffin tissue in the heart and veins of fish. Comprehensive accounts of the distribution and catecholamine content of chromaffin tissue in the vertebrates can be found in review articles by Coupland (1971; 1972) and Nilsson (1983). In this chapter chromaffin systems will be discussed under the three major headings of: adrenal medulla, small cells of autonomic ganglia and other extra-adrenal chromaffin tissues.

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ADRENAL MEDULLA Separate NA and A cells have been recognized in all vertebrate classes and the two types of storage granules have a similar range of diameters throughout the vertebrates (Coupland, 1971). However, despite this apparent consistency, it is clear that the proportions of NA and A contained in the chromaffin tissue of different species varies widely (Coupland, 1972) as do the stimulus-induced increases in plasma catecholamines. Plasma catecholamine levels are mainly determined by secretion from the adrenal medulla but the morphology and innervation of extraadrenal chromaffin tissue indicates that this tissue may contribute as well. It is therefore inadvisable to link plasma catecholamine levels purely with adrenomedullary secretion. Unfortunately there is insufficient fine structural evidence to be able to state that all groups of extra-adrenal chromaffin cells, not including carotid and aortic bodies, have a preganglionic sympathetic innervation which causes release of their amines into the bloodstream, so that the size of extra-adrenal contribution is difficult to determine. Fish True adrenal glands, in which there is a distinct medulla formed of chromaffin tissue, are only present in birds, reptiles and mammals. In fish, the chromaffin tissue is related mainly to parts of the cardiovascular system but also to the head kidney and sympathetic ganglia. Two types of chromaffin cells have been recognized in Salmo gairdneri (Mastrolia, Gallo and La Marca, 1984) which correspond to NA and A cells but according to biochemical data (Coupland, 1972) there is likely to be a great deal of interspecific variation in the proportions of these two cell types amongst the superorders and orders of the class Pisces. The demonstration that catecholamine release from chromaffin tissue in teleosts is under the control of the sympathetic nervous system (Nilsson, 1976; Wahlqvist and Nilsson, 1977; 1980) provides convincing evidence for an important role of chromaffin tissue in circulatory control. Amphibia In amphibia, the chromaffin tissue is primarily associated with the kidney: in Urodeles as islets either within, between or on the ventral surface of the kidney, but in Anurans as a distinct strip of tissue on the ventral surface of the kidney (Moore, 1964). The position of this ventral strip can be correlated with the evolutionary status of members of the Anura (Grassi, Milano and Accordi, 1983). Substantial aggregations of chromaffin cells are also located in relation to sympathetic chain ganglia (Vogel and Model, 1977; Watanabe, 1977; Weitsen and Weight, 1977) and around the abdominal aorta in amphibia (Hill, Watanabe and Burnstock, 1975). Irrespective of the location of amphibian chromaffin tissue the ultrastructural appearance of the two types of chromaffin cells and their storage granules is consistent (Piezzi, 1967; Grassi, Milano and Accordi, 1983). The effect of adrenomedullary stimulation is a general systemic vasoconstriction (Moore, 1964) as well as the chronotropic and inotropic effects on the heart that were used in the

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classical experiments of Loewi (1936). Exocytosis of adrenal granules has been observed in the Urodele Siren lacertina by electron microscopy where many granules are observed subadjacent to or fusing with the plasma membrane (Accordi, 1988). The amphibians are the most primitive vertebrates in which small granule containing (SGC) cells are encountered in chromaffin tissue, having been identified both in Urodeles (Accordi, 1988) and Anurans (Accordi and Gallo, 1982). 1SGC cells are characterized by their smaller size than the surrounding chromaffin cells and by their content of granular vesicles in the 100-200 nm range. Immunocytochemical methodology has revealed that, in addition to catecholamines, adrenal medullary granules also contain neuropeptides. In the frog Rana ridibunda most of the adrenal chromaffin cells which show tyrosine hydroxylase immunoreactivity (TH-IR) also show vasoactive intestinal polypeptide immunoreactivity (VIP-IR) and, in addition, about 40% of these showed methionine-or leucineenkephalin immunoreactivity (Met-or Leu-enkephalin-IR) (Leboulenger et al., 1983). Serotonin has also been localized in both NA and A cells of the amphibian adrenal medulla (Kuramoto, 1987). Varicose nerve axons are present in the adrenal medulla of the bullfrog Rana catesbeiana in relation to chromaffin cells (Kuramoto, 1987) some of which are thought to be sensory as they are immunoreactive for substance P and calcitonin gene-related peptide (CGRP). Additionally, other nerves immunoreactive for neuropeptide Y (NPY) were localized around small arteries and arterioles in the gland and are likely to have vasoconstrictor effects. Many enzymes have been demonstrated in chromaffin cells by enzyme histochemical techniques (Hopwood, 1971). In contrast to mammals where adrenomedullary acid phosphatase (AP) distribution is highly species-specific, it is widespread in both urodele (Berchtold, 1970) and anuran amphibian adrenomedullary cells. Electron microscopic localization of AP in a number of anura (Mastrolia and Manelli, 1979; Manelli, Mastrolia and Arizzi, 1981) has revealed AP not only in the Golgi apparatus and primary lysosomes of chromaffin cells but also on the membranes of the catecholamine storage granules where it is suggested that the enzyme may be involved in the metabolism of the granule core proteins. The demonstration of acetylcholinesterase (AChE) in the chromaffin cells of Urodeles (Mastrolia, Gallo and Manelli, 1976), as in other vertebrates, is likely to be related to their cholinergic innervation. Reptiles and birds A discrete adrenal gland is present in reptiles and birds although in the former it is closely related to the interrenal tissue and in birds is not divided into a distinct medulla and cortex. In both classes NA and A cells are clearly recognizable on account of their granule morphology (Coupland, 1971; Unsicker 1973a; 1976; Chungsamarnyart, Fujioka and Kitoh, 1981) but the relative amounts of NA and A differ widely between avian and reptilian species (see Nilsson 1983). An analysis of NA and A concentrations in a large number of avian species has led Ghosh (1962) to propose that the presence of phenylethanolamine-N-methyltransferase (PNMT), the enzyme that methylates NA to A, reflects an evolutionary development as it is absent in more primitive avian species. The adrenomedullary cells are innervated

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by cholinergic-type synapses and contain AChE (Gallo and Mastrolia, 1979) though axons with differing vesicle contents have also been observed (Unsicker 1973a, b, 1976). Some of these latter axons have the appearance of peptidergic axons but the lack of immunohistochemical investigations on avian and reptilian adrenal medullae prevents confirmation of the presence of peptidergic nerves. SGC cells and also sympathetic neurons occur in avian (Unsicker, 1973a) and in reptilian (Unsicker, 1976) adrenal medullae but their functional significance is unknown. The physiological effects of released catecholamines are similar in birds and reptiles to other vertebrates but there is a considerable interspecific variation in responses to NA and A particularly with regard to carbohydrate metabolism (Assenmacher, 1973). Neither NA or A release results in thermogenesis in birds (Freeman, 1970a, b). Mammals Separate NA and A storing chromaffin cells can be distinguished in the mammalian adrenal medulla and the relative proportions of the two cell types approximates to the relative concentrations of NA and A in the adrenal gland (Coupland, 1984) which have wide interspecific variations (Holzbauer and Sharman, 1972). During development of the medulla NA can be detected initially and the appearance of A takes place later, coincident with the first detectable expression of PNMT (Verhofstad et al., 1985). Similarly, the first adrenomedullary cell type has the appearance of an NA cell. Only during the perinatal period are NA and A encountered in the same adrenomedullary cells (Elfvin, 1967; Verhofstad et al., 1985) before the distinctive, separate adult cell types are observed. As in lower vertebrates (Jonsson and Nilsson, 1983) PNMT is the rate limiting enzyme in the synthesis of A from NA in mammals and the expression of the enzyme is under the control of adrenal corticosteroids (Wurtman and Axelrod, 1965; Bohn, Goldstein and Black, 1981) which explains the approximation of steroid-producing tissue to medullary chromaffin cells throughout the vertebrates. The study of mammalian adrenomedullary tissue, particularly in the rat, has provided fundamental information with respect to our understanding of secretory mechanisms from endocrine and nervous tissue, of the factors governing differentiation of sympatho-adrenal cell lineages and of the possibility of intracerebral transplants for the alleviation of degenerative brain disorders. Milestones in adrenal medullary research have recently been highlighted by Coupland (1989) and this chapter will therefore concentrate on the current explosion of information concerning the occurrence of neuropeptides in adrenomedullary cells and nerves of the adrenal medulla. The first report, by immunohistochemistry, of the existence of a neuropeptide in adrenomedullary cells was of enkephalin by Schultzberg et al. (1978). Subsequently, enkephalin was shown to be stored in the same granules as catecholamines (Stern et al., 1979) and co-released from them (Livett et al., 1981). Since then many more neuropeptides have been localized in the adrenomedullary cells of many different vertebrate species including man. Amongst those occurring in several species are somatostatin, substance P, VIP, CGRP, enkephalin and other opioid peptides as well as vasopressin and oxytocin. Peptides with an apparently more restricted adrenomedullary distribution are atrial natriuretic polypeptide

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(Inagaki et al., 1986), delta sleep-inducing peptide (Eckman et al., 1987) and adrenocorticotrophic hormone (Ito et al., 1981). The occurrence of these neuropeptides is reviewed by Polak and Bloom (1987) and by Pelto-Huikko (1989) who, in addition, provides data on the occurrence of NPY, neurotensin and galanin in several species. It is clear that there are wide differences between species with regard to a) the occurrence of individual neuropeptides in adrenomedullary cells, b) the co-storage of individual neuropeptides in either NA or A cells, c) the co-storage of more than one neuropeptide in either NA or A cells and d) the proportions of cells demonstrating differing combinations of neuropeptides. Generally, more different neuropeptides are encountered in A than NA cells -for example NPY and CGRP have yet to be reported in NA cells (Pelto-Huikko, 1989; Pelto-Huikko and Salminen 1987). Other proteins such as dopamine and cyclic AMP regulated phosphoprotein-32 (DARRP-32) have been reported in the cow adrenal medulla (Hemmings and Greengard, 1986) and it is also likely that adrenal chromaffin cells produce several neuronotrophic factors which are related to the extrinsic innervation of the medulla (Unsicker et al., 1987). The adrenal medulla has a preganglionic cholinergic innervation via the splanchnic nerves which originates from thoracic spinal cord segments T1-T10 with the most cell bodies occurring at T8-9 (Schramm et al., 1975; Kesse, Parker and Coupland, 1988). The ratio of preganglionic nerves to medullary chromaffin cells in the rat is of the order of 1:1000 (Parker et al., 1988) which suggests that not all medullary cells are directly innervated and that mechanism of stimulus-induced secretion may well involve intracellular coupling. Additionally the medulla is innervated by afferent nerves with cell bodies in dorsal root ganglia throughout the thoracic and upper lumbar levels (Mohamed, Parker and Coupland, 1988) and also in vagal sensory ganglia and the dorsal motor nucleus of the vagus (Coupland et al., 1989). A sparse plexus of nerves containing substance P and CGRP occurs in relation to the medullary chromaffin cells and it is likely that these are sensory afferents (Kuramoto, Kondo and Fijira, 1985; 1987; Pelto-Huikko, 1989). Nerve fibres showing neurotensin-, enkephalin-, galanin-, (Pelto-Huikko, 1989) and NPYimmunoreactivity (IR) (Kuramoto, 1986) have also been observed in the adrenal medullae of several species. It is clear that, despite considerable variation in the arrangement of the adrenal medulla throughout the vertebrates, the concept that it is simply comprised of NAand A-storing cells has to be revised. The discovery of SGC cells, adrenal neurons and the differential expression of a multitude of neuropeptides by adrenomedullary cells now causes one to regard the adrenal medulla as a highly complex tissue which persists into old age and is able to proliferate in response to a variety of stimuli. (Tischler et al., 1989). SMALL CELLS OF AUTONOMIC GANGLIA Chromaffin cells were first reported in sympathetic ganglia by Kohn (1903) in his survey of extra-adrenal ‘paraganglia’. However, contemporary interest in these cells was awakened by Erànkô and Hàrkônen (1963) and Hamberger et al. (1963), with

FIGURE 3.3 Organ o f Zuckerkandl o f a 10 day old rat revealed by formaldehyde-induced catecholamine fluorescence. This mass o f extra-adrenal ‘chromaffin tissue* will almost entirely regress to leave only isolated retroperitoneal clusters o f cells in the adult. Calibration bar 50 pun.

FIGURE 3.4 Atlantic cod coeliac ganglion. Formaldehyde-induced catecholamine fluorescence showing clusters o f small, intensely fluorescent (SIF) cells amongst the larger, less fluorescent ganglionic neurons. Calibration bar 50 /¿m. By courtesy o f S. Nilsson.

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106 COMPARATIVE PHYSIOLOGY AND EVOLUTION

the discovery of ‘small intensely fluorescent’ (SIF) cells interspersed amongst the principal neurons of rat and cat sympathetic ganglia. It was assumed that they were ganglionic chromaffin cells on account of their similar fluorescent appearance to adrenal medullary cells after FIF. Following this, SGC cells were identified amongst the principal neurones in sympathetic ganglia by electron microscopy (Siegrist et al., 1968; Grillo, 1966) and seen to have both afferent and efferent synapses (Williams, 1967; Matthews and Raisman, 1969). Confirmation that SIF and SGC cells were the same cell type visualised by differing methods was achieved by Grillo et al., 1974). The use of glutaraldehyde-potassium dichromate methods by Santer et al., (1975) visualised chromaffin-positive small cells in rat sympathetic ganglia but as the numbers of SIF cells far exceeded the numbers of chromaffin cells it was clear that not all the small cells can be demonstrated by the indirect chromaffin reaction. This underlines the fact that the term ‘chromaffin’ is not a strictly accurate one with respect to the small cells of sympathetic ganglia. Subsequently, SIF cells were discovered in various sympathetic ganglia of vertebrate classes from teleosts (Watson, 1980; Figure 3.4) upwards and the voluminous literature of the distribution of SIF cells has been comprehensively reviewed by Taxi (1979) and Bock (1982). SIF cells have also been found in association with cardiac (Jacobowitz, 1967), ciliary (Ehinger and Falck, 1970) and pterygopalatine (Leblanc and Landis, 1989) parasympathetic ganglia thereby demonstrating that they are not confined exclusively to the sympathetic ganglia. From the mass of literature on ganglionic SIF and SGC cells it emerged that they do not form a homogeneous population of cells as subtypes can be recognized on a number of differing criteria. Firstly, they do not all exhibit the same FIF emission which ranges from an intense pale green through yellow to light brown- the yellow/brown fluorescing SIF cells probably corresponding to those which do display a chromaffin reaction (Kemp et al., 1977). Secondly, the morphology of their granule populations differs with some SGC cells containing granules resembling either adrenomedullary NA or A granules and others containing smaller dense-cored granules like those of adrenal SGC cells. Thirdly, their content of amines and co-stored peptides shows interganglionic and interspecific variation (see below). Fourthly, SIF cells have consistently been shown to occur either as solitary cells or in clusters. The solitary SIF cells have axon-like processes and are identifiable by electron microscopy as being interneuronal in position as they receive cholinergic-type, presumably preganglionic, synapses and themselves make efferent synaptic contact with dendrites or somata (Matthews and Nash, 1970; Polonyi and Kapeller, 1984) of principal ganglionic neurones. Clusters of SIF cells, made up of varying numbers of cells, are found closely related to venules (Abe et al., 1983; Matthews, 1989) or fenestrated capillaries forming glomus-like arrangements reminiscent of endocrine tissue. As fenestrated capillaries are generally uncommon in sympathetic ganglia, much significance has been placed on the close relationship between clustered SIF cells and fenestrated capillaries. As a result of these two arrangements of SIF cells they have been classified as Types I and II respectively (Williams et al., 1976) and putative interneuronal and endocrine functions respectively ascribed to them. A third function, that of chemosensation, has also been suggested for the Type II cells (Lever et al., 1976).

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Since SIF cells were first discovered it has been repeatedly pointed out by many authors that the number of SIF cells differs widely not only between different ganglia and between different species but also between the same ganglion of individual members of a species. Perhaps the most extreme example of this is the range of 40-3332 SIF cells occurring in the paravertebral sympathetic ganglia of bullfrogs (Watanabe and Tonosaki, 1986). Considerable interspecific variation in numbers of SIF cells/volume of tissue has been demonstrated (Chiba and Williams, 1975; Williams et al., 1976; Tomasulo, Sansone and Melsaac, 1982) but accurate counts of the exact numbers of principal neurones and SIF cells in individual ganglia have yet to be made. These numerical differences argue against SIF cells having an essential role in ganglionic transmission, a proposed function that has been forcibly argued against on electro-physiological grounds by Weight and Smith (1980). Also the location of both Types I and II cells is not recognizably consistent with respect to the larger population of principal neurones which makes it difficult to associate the SIF cells with a particular subset of ganglionic neurones associated with a particular function. Recently, however, Reuss and Schroder (1988) have noted that many of the sympathetic neurones of the rat SCG which project to the pineal gland are closely associated with Type I SIF cells and this suggests that this particular pathway might be modulated in some way by the SIF cells. However, to date, the role of SIF cells in ganglionic function is still unclear but there are some important pieces of evidence to consider before dismissing the SIF cells as, perhaps, undifferentiated or vestigial neural crest derivatives located within autonomic ganglia. Matthews (1978) demonstrated the exocytosis of granule cores from SGC cells of the rat and suggested that this process could occur in response to appropriate stimulation, presumably by the preganglionic cholinergic input. The random location of intraganglionic SIF cells has made the testing of this theory by electrophysiological methods very difficult but Dunn and Marshall (1985) discovered that SIF cells in the lumbar sympathetic ganglia of frogs can be seen as small clusters in living ganglia and have successfully made intracellular recordings from them. They found that the SIF cells had a resting membrane potential of —20 to —70 mV, received a functional synaptic input and could be depolarised by acetylcholine, thus establishing their responsiveness to preganglionic stimulation in amphibia. This has now been established in mammals by Yang and Li (1987) who demonstrated that stimulation of the splanchnic nerves of the guinea pig caused a decrease in fluorescence intensity and 30% reductions in granule content of SIF/SGC cells of the coeliac ganglion which was also associated with an increase in monoamine activity. Ganglionic DA metabolism, equated with SIF cell activity, can be activated by muscarinic agonists (Lutold, Karoum and Neff, 1979) and increased by long-term hypoxia (Dalmaz et al., 1988) in the rat SCG. Dalmaz et al. (1988) have also reported that ganglionic NA levels remained unaffected by hypoxia. As 40% of the ganglionic DA has been calculated to be located in SIF cells (Borghini, Dalmaz and Peyrin, 1991) the possibility that the Type II SIF cells are chemoreceptive must receive renewed attention. If this proves to be so, it might explain the frequent occurrence of SIF cell clusters in relation to vascular elements just beneath the

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ganglion capsule. However, careful ultrastructural analyses of such SIF cell clusters by Case and Matthews, (1985) reveal highly complex relationships between the SIF cells and their neuronal and vascular neighbour (Figure 3.4) which, as yet, fails to clarify their functional role(s). The problem of isolating SIF cells in order to investigate their biology was overcome by Doupe, Patterson and Landis, (1985) who established pure SIF cell cultures by growing dissociated rat ganglia in vitro in the absence of Nerve Growth Factor (NGF). These cultured SIF cells had the appearance and biochemical properties of SIF cells in vivo and under appropriate experimental conditions could be transformed into other recognisable neural crest derivatives such as neurones. This led Doupe, Patterson and Landis, (1985) to propose that SIF cells could be undifferentiated precursor cells able to respond to particular signals for differentiation. However the relatively late appearance of SIF cells during ganglionic development has led Leblanc and Landis (1989) to cast doubt on this proposition. The precise nature of the catecholamine (CA) content of SIF cells has proved a difficult problem to solve by microfluorometric methods on account of the very similar emission spectra of DA and NA. The use of mass fragmentography and gas liquid chromatography by Gerold et al. (1982) on isolated groups of SIF cells has confirmed the presence, in varying amounts, of three CAs namely A, NA and DA and it is probable that these exist in differing amounts in different SIF cells as immunohistochemical staining for dopamine-/3-hydroxylase (DBH) and PNMT (Konig and Heym, 1978) and the early observation of several types of SGC cells (Lu et al., 1976) predicted. Sakai et al. (1989) have localised DA by electron microscopic immunohistochemistry in intraganglionic processes which make synaptic contact with principal ganglionic neurones and suggests that this could be the morphological correlate for an interneuronal role of DA in Type I SIF cells. Immunohistochemistry has revealed, as in adrenomedullary cells, the presence of peptides and other biogenic molecules. 5-HT (Verhofstad et al., 1981) is colocalised with histamine and its synthesizing enzyme histidine decarboxylase (Karhula et al., 1990) in SIF cells of guinea-pig prevertebral ganglia and glutamic acid decarboxylase is also present in these cells (Karhula et al., 1988). Peptides such as NPY, substance P, enkephalin, VIP and dynorphin (DYN) (see Polak and Bloom 1987) were located in SIF cells by immunocytochemistry in the late seventies and more recently bombesin/gastrin releasing peptide has been found colocalised with Metenkephalin-Arg-Gly-Leu in rat SIF cells (Helen et al., 1984). Interestingly, in this report, SIF cells were observed that were not immunoreactive for these two peptides but surrounded by immunopositive fibres. In a detailed analysis of the occurrence of five different peptides in SIF cells of the guinea-pig inferior mesenteric ganglia, Chiba and Masuko (1989a) report that 10% of SIF cells contained no detectable peptides whilst the percentage of SIF cells containing one particular peptide or another ranged from 24-46%. Subsequent work by these authors (Chiba and Masuko, 1989b) revealed several sets of peptides in certain SIF cells and have suggested that this might be a basis for considering the existence of groups of functionally distinct SIF cells. Clearly the small cells of autonomic ganglia, whether referred to as chromaffin,

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SIF or SGC cells, have yet to give up all their secrets but it is evident from those instances where it has been possible to investigate them individually, that their potential to perform functional roles has been established. OTHER EXTRA-ADRENAL CHROMAFFIN TISSUE Chromaffin cells are widely distributed and not confined to the adrenal medulla and autonomic ganglia. In fish, where no adrenal medulla is recognisable, chromaffin tissue is found in abundance in such organs as the head kidney, posterior cardinal veins, intercostal arteries, chambers of the heart and, in elasmobranchs, segmentally arranged in association with sympathetic ganglia (Figures 3.5 and 3.6). The capability for local catecholamine synthesis in such tissue has been demonstrated by Jonsson (1983). The anatomical distribution and predominating CA contents vary in different groups of fish and this information has been reviewed by Nilsson (1983). The elevated blood CA levels which result from experimentally-induced stress are derived from this chromaffin tissue which has been shown, at least in teleosts (Nilsson, Abrahamsson and Grove, 1976), to be under preganglionic cholinergic control. Thus in fish a widely dispersed chromaffin system is present and exerts controlling influences over the cardiovascular system. This is of particular significance as a sympathetic cardioexcitatory innervation is not always present as it is in higher vertebrates. Cyclostomes present the most extreme example as an extrinsic sympathetic cardiac innervation is totally absent but the heart and large veins contain much chromaffin tissue separated from the blood by a single layer of endothelial cells. It is likely that extra-adrenal chromaffin tissue may be involved in cardiac function in other vertebrate groups as SIF/SGC cells have been observed in the heart, often juxtapposed to cardiac ganglia, for example in amphibians (Axelsson and Nilsson, 1985; Tay and Wong, 1988) and reptiles (Chiba and Yamauchi, 1973) as well as in higher vertebrates including man (Dail and Palmer, 1973). Extra adrenal chromaffin tissue other than that in autonomic ganglia and in the heart occurs in amphibia, birds and reptiles (see reviews by Coupland, 1972; Taxi, 1979; Nilsson, 1983) especially in the para-aortic regions, but there have been few systematic investigations into the function of this tissue in these groups of vertebrates. Attention to extra-adrenal chromaffin tissue was focused by the classical papers of Zuckerkandl (1901) and Kohn (1902). The organ of Zuckerkandl (Figure 3.3) a mass of retroperitoneal chromaffin tissue located in a pre-and paraaortic position primarily in the upper part of the abdominal cavity was initially regarded as reaching its maximal development in the perinatal period and then degenerating during childhood. By investigating a series of mammals throughout their lifespan Coupland (1965) discovered that remnants of the Organ of Zuckerkandl persisted into adulthood. More recently Hervonen et al. (1978) have demonstrated well vascularised collections of retroperitoneal chromaffin tissue in adult humans and Partanen et al. (1984) have reported an increase in abdominal chromaffin tissue in aged rats. It is suggested that this chromaffin tissue may contribute to the undiminished response to stress (McCarty, 1985) and increased adrenal

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FIGURE 3.5 Axillary body from the spiny dogfish, Squalus acanthias. Cluster o f chromaffin cells revealed by immunohistochemical reaction with DBH antiserum. The strongly reactive chromaffin cells form the anterior part o f the axillary body. Calibration bar 200 /¿m. By courtesy o f S. Holmgren.

FIGURE 3.6

Transverse section o f left posterior cardinal vein (‘azygos vein’) from the African lungfish,

Protopterus aethiopicus. Formaldehyde-induced catecholamine fluorescence showing masses o f chromaffin cells lining the venous lumen. Calibration bar 200 /¿m. Reproduced with permission from Abrahamsson et al. (1979).

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CA secretion (Ito et al., 1986) in aged rats but adrenomedullary hyperplasia (Tomlinson and Coupland, 1990) and hypertrophy (Coupland and Tomlinson, 1989) may also account for this. Further evidence for the maintenance of chromaffin tissue throughout life is provided by the observation of mitoses in extra-adrenal (Bohn, 1987; Mascorro and Yates, 1989) and adrenomedullary (Coupland and Tomlinson, 1989) cells and by the increase that takes place in tyrosine hydroxylase gene expression in the adrenal medulla (Strong et al., 1990). The widespread occurrence of other extra-adrenal chromaffin tissue in mammals can be demonstrated, to cite but a very few examples, by reports of solitary and clustered SIF cells/chromaffin cells/SGC cells in such tissues as the nerve plexus of the guinea-pig gall bladder (Cai and Gabella, 1984), in the pelvic innervation (Furness and Costa, 1976), in nerve trunks and organs of the neck and thorax (McDonald and Blewett, 1981) and other cells of neuroectodermal origin such as the enterochromaffin cells. The widely dispersed arrangement and the difficulty of locating them other than by microscopy makes functional interpretation of such chromaffin tissue highly problematical. On the other hand the function of chemosensation is attributed throughout the vertebrates to the carotid, aortic and subclavian bodies. These organs have long been known to contain chromaffin cells but the precise role of chromaffin cells in the process of chemosensation is still unresolved (see review by Pallot, 1987). As in other chromaffin cells, the discovery that they contain many neuropeptides (Varndell et al., 1982; Heym and Kummer, 1989) and also contain peptidergic nerves (Kummer, Gibbins and Heym, 1989) will further delay our understanding of these important chemosensory organs. It is over one hundred and twenty years since the discovery that the adrenal medulla could be specifically stained by chromium compounds and nearly one hundred years since the introduction of the term ‘chromaffin’. This was a straightforward scientific observation of the time. It is somewhat humbling but nevertheless important to admit that in 1993, even with highly sophisticated methodology, we are still far from a complete understanding of the biology of the chromaffin systems.

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Unsicker, K. (1973b). Fine structure and innervation of the avian adrenal gland II. Cholinergic innervation of adrenal chromaffin cells. Z Zellforsch Mikrosk A nat, 154, 417-442. Unsicker, K. (1976). Chromaffin, small granule-containing and ganglion cells in the adrenal gland of reptiles. Cell Tissue Res., 165, 477-508. Unsicker, K., Lietzke, R., Gehrke, D. and Stogbauer, F. (1987). Chromaffin cells: a novel source for neuronotrophic factors. Exp. Brain Res. Ser., 16, 120-124. Varndell, I.M., Tapia, F.J., De Mey, J., Rush, R.A., Bloom, S.R. and Polak, J.M. (1982). Electron immunocytochemical localization of enkephalin-like material in catecholamine-containing cells of the carotid body, the adrenal medulla and in phaeochromocytomas of man and other animals. J. Histochem. and Cytochem., 30, 682-690. Verhofstad, A.A.J., Steinbusch, H.W.M., Joosten, H.W.M., Colenbrander, B. and MacDonald, A.A. (1981). Development of the noradrenaline- and adrenaline-storing cells in the adrenal medulla and its control by the adrenal cortex. Acta Morphol Neerl-Scand, 19, 230. Verhofstad, A.A.J., Coupland, R.E., Parker, T.L. and Goldstein, M. (1985). Immunohistochemical and biochemical study on the development of the noradrenaline- and adrenaline-storing cells of the adrenal medulla of the rat. Cell Tissue Res., 242, 233-243. Vogel, D.L. and Model, P.G. (1977). Development of the sympathetic system in the Mexican Axolotl Ambystoma mexicanum. Dev. Biol., 56, 76-96. Wahlqvist, I. and Nilsson, S. (1977). The role of sympathetic fibres and circulating catecholamines in controlling the blood pressure in the cod Gadus morhua. Comp. Biochem. Physiol., 57C, 65-67. Wahlqvist, I. and Nilsson, S. (1980). Adrenergic control of the cardio-vascular system of the Atlantic cod, Gadus morhua, during ‘stress’. J. Comp. Physiol., 137, 145-150. Watanabe, H. (1977). Ultrastructure and function of the granule-containing cells in the anuran sympathetic ganglia. Arch Histol Jap, 40, 177-186. Watanabe, H. and Tonosaki, A. (1986). ‘SIF’ cells in the sympathetic ganglia of the bullfrog, Rana catesbeiana: variety in population and innervation. Cell Tissue Res., 245, 413-421. Watson, A.H.D. (1980). The structure of the coeliac ganglion of a teleost fish Myoxocephalus scorpius. Cell Tissue Res., 210, 155-165. Weight, F.F. and Smith, P.A. (1980). Small intensely fluorescent cells and the generation of slow postsynaptic inhibition in sympathetic ganglia. Adv. Biochem. Psychopharmacol., 25, 159-171. Weitsen, H.A. and Weight, F.F. (1977). Synaptic innervation of sympathetic ganglion cells in the bullfrog. Brain Res., 128, 197-211. Williams, T.H. (1967). Electron microscopic evidence for an autonomic interneurone. Nature, 214, 309-310. Williams, T.H., Chiba, T., Black, A.C., Bhalla, R.C. and Jew, J. (1976). Species variation in SIF cells of superior cervical ganglia: Are there two functional types? In SIF cells, Structure and Function o f the Small, Intensely Fluorescent Sympathetic Cells, edited by O. Eranko, pp. 143-162. Washington: DHEW Publication. Wood, J.G. (1974). Positive identification of intracellular biogenic amine reaction product with electron microscope X-ray analysis. J. Histochem. Cytochem., 22, 1060-1063. Wood, J.G. and Barrnett, R.J. (1964). Histochemical demonstration of norepinephrine at a fine structural level. J. Histochem. Cytochem., 12, 197-209. Wurtman, R.J. and Axelrod, J. (1965). Adrenaline synthesis: control by the pituitary gland and adrenal glucocorticoids. Science, 150, 1464-1465. Yang, G. and Li, Z-T. (1987). Histochemical and ultrastructural changes of SIF cells after electrical stimulation of the greater splanchnic nerves. Acta Physiol. Sin., 39, 54-60. Zuckerkandl, E. (1901). Ueber Nebenorgane des Sympathicus in Retroperitonealraum des Menschen. Anat A nz, 15, 97-107.

4 The Gastrointestinal Canal Jörgen Jensen and Susanne Holmgren Department o f Zoophysiology, University o f Göteborg, Göteborg, Sweden The autonomic nervous control of the gastrointestinal canal in vertebrates is reviewed, with special emphasis on the control of gut motility in non-mammalian vertebrates. The presence and function of different established and putative neurotransmitters is discussed. In addition to the presence of adrenergic and cholinergic nerves in the gut, 5-hydroxytryptamine, ATP or a related substance, and several neuropeptides are suggested to act as neurotransmitters in the gastrointestinal canal of vertebrates. Among the neuropeptides demonstrated in the gut of non-mammalian vertebrates by immunohistochemical methods are peptides similar to bombesin, gastrin/cholecystokinin, neuropeptide Y, neurotensin, opioids, somatostatin, substance P (tachykinins) and vasoactive intestinal polypeptide. Functional studies show that many of the substances discussed may well be involved in the nervous control of the gut motility, although the function and mechanism of action varies to some extent between species and between vertebrate groups. KEY WORDS gut; acetylcholine; catecholamines; 5-hydroxytryptamine; purines; neuropeptides

INTRODUCTION The classical view of a nervous regulation of the gut which acts solely through antagonistic cholinergic (excitatory) and adrenergic/noradrenergic (inhibitory) nerves has been considerably changed during the last few decades. Numerous other transmitter substances have been proven or suggested to be involved in the control of the gastrointestinal tract. Among these substances are 5-hydroxytryptamine, adenosine 5'-triphosphate (ATP) or a related substance, and various peptides (neuropeptides, regulatory peptides). This chapter will focus on comparative aspects of the autonomic innervation of the gastrointestinal canal in vertebrates, with particular emphasis on non-mammalian vertebrates. For more detailed information on the morphology and function of the autonomic nervous system of the mammalian gut, the reader is referred to recent excellent reviews by e.g. Furness and Costa (1987), Daniel (1991), and other books in this series. The knowledge of the innervation of the gut in non-mammalian vertebrates by different types of nerves is in many cases meagre. The available information is often 119

120 COMPARATIVE PHYSIOLOGY AND EVOLUTION

limited to histochemical localization of the putative neurotransmitter, while the effects on the target tissue and the mechanisms behind the effects are unknown. Following the development of immunohistochemical methods for the identification of transmitters, numerous studies have described the distribution of neuropeptides in the gastrointestinal canal of vertebrates. Most information on adrenergic and cholinergic innervation in non-mammalian vertebrates refers to older studies, and has been extensively reviewed by e.g. Burnstock (1969), Fänge and Grove (1979), and Nilsson (1983).

MORPHOLOGY OF THE ENTERIC NERVOUS SYSTEM The nervous regulation of the functions of the gut is performed primarily by the enteric nervous system, which is considered a separate part of the autonomic nervous system in addition to the sympathetic and parasympathetic divisions. The enteric nervous system comprises all of the neurons intrinsic to the gut, i.e. all the neurons with their cell bodies within the gut wall, and is largely independent of central nervous control. The activity of the enteric nervous system may be influenced and controlled by extrinsic autonomic nerves, but anatomical and physiological findings indicate that most gastrointestinal functions still work more or less properly after disruption of the extrinsic pathways (Langley, 1898, 1921; Furness and Costa, 1980; Gershon, 1981; Gibbins, chapter 1 this volume). The enteric nervous system, as described in mammals, is arranged as interconnected plexuses, the major parts consisting of the myenteric and the submucous plexuses. The myenteric plexus (Auerbach’s plexus) is situated between the outer longitudinal muscle layer and the circular muscle layer, and the submucous plexus (Meissner’s plexus) within the connective tissue of the submucosa. Neurons from the plexuses connect to each other and to the muscle layers, the submucosa and the mucosa (Furness and Costa, 1980). A ganglionated plexus has been described in the intestinal wall of cyclostomes and, as in mammals, the enteric nervous system of the other vertebrates contains both a myenteric and a submucous plexus. In reptiles, neuronal cell bodies are found in the submucous plexus of the turtle, Pseudemys scripta elegans, while they are absent from the intestinal submucosa of the lizard, Tiliqua rugosa. Similarly, only a few or no neuronal cell bodies are present in the submucous plexus in fish and amphibians, indicating a more simple arrangement of the enteric nervous system in these groups compared to mammals (Kirtisinghe, 1940; Gunn, 1951; Burnstock, 1959, 1969; Read and Burnstock, 1968; Nilsson, 1983; Timmermans et al.9 1991). The avian enteric nervous system comprises ganglionated myenteric and submucous plexuses and, in addition, the control of the gut is influenced by the ganglionated nerve of Remak, which runs along the gut (Bennett, 1974).

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ENDOCRINE A N D PARACRINE CELLS Many of the neurotransmitter substances found in enteric neurons are also suggested to act as hormones in the gastrointestinal canal. In fact, three types of cells that contain biologically active substances have been described, although the demarcation between the cell types may be vague. In addition to the neurocrine cells, which release their transmitter from nerve terminals to act on postsynaptic cells, there are endocrine and paracrine cells. From the endocrine cells the regulatory substance is released into the blood stream and transported to the target cells as a hormone, while from paracrine cells the regulatory substance is released from small cell processes to act on neighbouring cells (Larsson, 1980). The same three cell types have been reported in the non-mammalian gut.

CATECHOLAM INES The noradrenergic neurons supplying the mammalian gut have almost exclusively an extrinsic origin. In the gut wall the catecholamine-containing fibres surround neuronal cell bodies of enteric neurons in the myenteric and submucous plexuses. The main function of the noradrenergic nerves is probably a presynaptic inhibition of the release of acetylcholine from cholinergic nerves (Furness and Costa, 1980, 1987; Gershon, 1981). Although noradrenergic nerves are generally inhibitory in the mammalian gut, it appears that in the non-mammalian vertebrates the effects of catecholamines and the type of receptors involved vary to a great extent between species. Catecholamines produce variable responses, including relaxations and contractions, on the intestine of Myxine. However, the significance of an adrenergic nerve control of the gut motility in cyclostomes is unclear, although the vagus nerve contains small amounts of catecholamines (von Euler and Östlund, 1957; von Euler and Fänge, 1961; Holmgren and Fänge, 1981). In the spiny dogfish, Squalus acanthias, numerous catecholamine-containing axons are observed in the splanchnic nerve and in the stomach wall (Holmgren and Nilsson, 1983a). Adrenaline contracts the stomach of Squalus and electrical stimulation of the splanchnic nerve produces a contraction of the stomach which is blocked by phentolamine, indicating a splanchnic adrenergic excitatory supply to the stomach (Nilsson and Holmgren, 1983). Adrenaline is excitatory on stomach preparations from Scyliorhinus and several species of Raja, but adrenergic nerves are probably not involved in the splanchnic excitatory effects in these species (Young, 1980a, 1983). Excitatory effects of catecholamines have also been described in the intestine of the elasmobranch, while an inhibitory action is found in the rectum (von Euler and Östlund, 1957; Young, 1983, 1988). Adrenergic nerve fibres have been demonstrated in the myenteric plexus and in the smooth muscle layers of the gut in several teleosts, as well as surrounding blood vessels of the gut (Figure 4.1). The action of catecholamines in the stomach is excitatory in some species, while inhibitory as well as both inhibitory and excitatory

122 COMPARATIVE PHYSIOLOGY AND EVOLUTION

ADRENALINE

2 -,

I P R O 1CT6 M

* PH E 10'6 M

kPa

ISOPRENALINE

t

t

t

-7

-7

-7

I P R O 10-6 M

kPa

-7

-7

P H ENYLE PHR IN E kPa

1 min

t

t

I P R O 1CT6 M

* P H E IO'6 M

/W v ^ /W v A -5

t

-5

t

-5

FIGURE 4.1. Catecholamines in the intestine of the Atlantic cod, Gadus morhua. Catecholamine fluorescence histochemistry shows the presence of adrenergic fibres in the muscie layers of the intestine (A) and the surrounding blood vessels in the intestinal submucosa (B). A: x300; B: x240. (C) demonstrates that the inhibitory effect of adrenergic agonists on the motility of the vascularly perfused intestine is mediated by both a-and /3-adrenoceptors. Boluses of 1 ml of agonist (log molar concentration) produced an inhibition of spontaneous contractions. Propranolol (PRO; /3-adrenergic antagonist) abolished the response to isoprénaline (/3-adrenergic agonist) but did not affect the response to adrenaline or phenylephrine (a-adrenergic agonist). Phentolamine (PHE; a-adrenergic antagonist) abolished the effects of adrenaline and phenylephrine. (C) is reproduced with permission from Jensen and Holmgren, 1985.

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123

effects are described in others. The excitatory response is generally «-adrenergic and the inhibitory effects are mediated by ß-adrenoceptors (Fänge and Grove, 1979; Nilsson, 1983, 1984; Kitazawa, Kondo and Temma, 1986). In the intestine, on the other hand, adrenaline is in general solely inhibitory. In a study by Kitazawa, Temma and Kondo (1986) it was suggested that in the intestinal bulb of the carp, the adrenaline-induced inhibition was caused by a decrease in the release of acetylcholine, as in mammals. The types of receptor mediating the effect on the intestine seem to vary between species. Thus, the inhibition has been reported to involve ß-adrenoceptors (e.g. Rhombosolea tapirina, Ammotretis rostrata), «-adrenoceptors (e.g. Cyprinus carpio) and both a- and ß-adrenoceptors (Gadus morhua; Figure 4.1) (Fänge and Grove, 1979; Grove and Campbell, 1979; Jensen and Holmgren, 1985; Kitazawa, Temma and Kondo, 1986). In the toad, Bufo marinus, splanchnic adrenergic fibres produce an inhibition of the motility in both the stomach and the large intestine (Boyd, Burnstock and Rogers, 1964; Campbell, 1969). Similarly, an inhibitory splanchnic adrenergic innervation is present in the intestine of birds, while the splanchnic adrenergic supply to the stomach is excitatory. An adrenergic innervation, including splanchnic adrenergic fibres, is described in reptiles, but the function of these fibres is largely unknown (see Nilsson, 1983). An inhibitory action of catecholamines mediated by ß-adrenoceptors is, however, indicated in the rectum of the rainbow lizard, Agama agama (Inyang and Okpako, 1989). In fish and crocodiles, adrenergic drugs are found to reduce the flow of blood to the gut, as in mammals (Figure 4.2; Axelsson et aU9 1989, 1991; Axelsson and Fritsche, 1991; Holmgren, Axelsson and Farrell, 1992).

ACETYLCHOLINE The view of acetylcholine as a transmitter involved in the excitatory control of gastrointestinal motility has been accepted for a long time. Immunohistochemical identification of cholinergic neurons has so far, especially in non-mammalian vertebrates, only been used in a limited number of studies and a majority of the reports on the distribution of cholinergic neurons are based on pharmacological methods. In mammals, intrinsic cholinergic nerves are involved in the excitatory part of the peristaltic reflex, and they supply the circular and longitudinal muscle layers as well as other enteric neurons. In addition, extrinsic cholinergic neurons act on enteric neurons innervating the smooth muscle of the gut (Furness and Costa, 1980; Gershon, 1981). The origin of cholinergic neurons in the gut of nonmammalian vertebrates is less clear. In general, the effects of cholinergic agonists are excitatory, although exceptions have been reported, such as an inhibitory effect of acetylcholine in the intestine of two lizards (Wacyk, Guerrero and Morello, 1984; Wacyk et al.9 1989). Not much is known about the cholinergic innervation in the gut of cyclostomes and elasmobranchs, but excitatory effects of acetylcholine are reported (von Euler and Ôstlund, 1957; Young, 1980a; Holmgren and Fange, 1981; Nilsson and Holmgren, 1983).

4-1

0J

■r „

t

n

- .

S u b sta n c e P

1 min

I--------------1

Jà

I- - - - - - - - 1

2 min

FIGURE 4.2 The inhibitory effects of adrenaline, and the stimulatory effects of substance P, VIP and bombesin on the flow of blood in the coeliac artery to the gut (FCoA, expressed in kHz dopplershift) in the spiny dogfish, Squalus acanthias, the Atlantic cod, Gadus morhua, the crocodile, Crocodylus porosus (adrenaline, substance P) and the caiman, Caiman crocodylus crocodylus (bombesin), in vivo. The peptides were injected in doses of 0.1 nmol/kg and the doses of adrenaline were 1 nmol/kg in dogfish, 10 nmol/kg in cod and 50 nmol/kg in the crocodile. The tracing showing the effect of adrenaline in the cod is reproduced by courtesy of Dr M. Axelsson, the other traces are reproduced with permission from Holmgren et al., 1989; Axelsson and Fritsche, 1991; Axelsson et al.y 1991; Jensen, Axelsson and Holmgren, 1991; Holmgren, Axelsson and Farrell, 1992.

Crocodilians

Gadus

Squalus

Adrenaline

COMPARATIVE PHYSIOLOGY AND EVOLUTION

THE GASTROINTESTINAL CANAL

125

Several studies have indicated that acetylcholine stimulates muscarinic receptors in the smooth muscle of the gut in teleosts (Grove and Campbell, 1979; Jensen and Holmgren, 1985; Kitazawa, 1989; Burka, Blair and Hogan, 1989). Cholinergic vagal excitatory fibres to the stomach have been demonstrated in some teleosts, although they seem to be absent from others, and the splanchnic nerve is suggested to supply the intestine with cholinergic excitatory fibres (see Fänge and Grove, 1979; Nilsson, 1983). Similarly, the gastrointestinal canal of amphibians, reptiles and birds is innervated by excitatory cholinergic vagal and splanchnic fibres (see Nilsson, 1983). The role of intrinsic cholinergic nerves in these groups of animals is, however, not well understood.

5-HYDROXYTRYPT AM INE 5-Hydroxytryptamine (5-HT) or serotonin is reported to have multiple actions on the mammalian gut, including a stimulatory effect on the gastrointestinal motility. A direct effect on the smooth muscle as well as an indirect action via the stimulation of cholinergic neurons are reported (see Gershon, 1981; Gershon et al., 1990). Amongst non-mammalian vertebrates, the presence and function of a serotonergic innervation has mainly been studied in different groups of fish, while the knowledge concerning amphibians, reptiles and birds is scarce. With the use of different histochemical methods (fluorescence histochemistry, immunohistochemistry, autoradiography), 5-HT or 5-HT-like immunoreactivity (IR) has been demonstrated in neurons and nerve fibres from the gut of cyclostomes, elasmobranchs, teleosts and amphibians. Immunoreactive (IR) nerves are found, with some variation between species, in the myenteric plexus and the muscle layers (Table 4.1). In many of these species, 5-HT has also been demonstrated in enterochromaffin cells (for references see Table 4.1, see also Santer, chapter 3). Adamson and Campbell (1988) were, on the other hand, unable to demonstrate any neurons containing 5-HT-like IR in some reptiles and birds. 5-HT stimulates the gut motility of elasmobranchs and the rectal motility of the lamprey, Lampetra fluviatilis, while no, or only weak, excitatory effects are produced on intestinal preparations from another cyclostome, Myxine glutinosa (von Euler and Ostlund, 1957; Johnels and Ostlund, 1958; Nilsson and Holmgren, 1983; Young, 1983, 1988). A spontaneous release of 5-HT, presumably from enteric neurons, has been demonstrated in the intestine of a teleost, Platycephalus bassensis, and this release is increased by transmural electrical stimulation (Anderson et al., 1991). A contractile effect of 5-HT on the gut is generally found in teleosts. In the stomach and intestine of the rainbow trout, the contractions are mediated by receptors on the smooth muscle (Holmgren, Grove and Nilsson, 1985; Burka, Blair and Hogan, 1989; Kitazawa, 1989), while in the intestine of several other teleosts an indirect pathway for 5-HT is indicated. This indirect pathway is, as in mammals, in several cases suggested to involve the release of acetylcholine from cholinergic nerve

126 COMPARATIVE PHYSIOLOGY AND EVOLUTION TABLE 4.1 Occurrence of 5-HT in nerves in the gut of non-mammalian vertebrates, revealed by formalin-induced fluorescence (FIF), autoradiography (ARG) or immunohistochemistry (IHC). Intestine

Method

Reference

+ a, x b -1-, X +, X

FIF FIF ARG

Sakharov and Salimova, 1980 Baumgarten é ta l., 1973 Goodrich étal., 1980

+

IHC IHC

Cimini et al., 1985 Holmgren and Nilsson, 1983a

HOLOCEPHALI Chimaera monstrosa

+ »X

IHC

Yui, Shimada and Fujita, 1990

DIPNOI Lepidosiren paradoxa

+

IHC

Nilsson and Holmgren, 1992

+

IHC

Burkhardt-Holm and Holmgren, 1992

+, X +, X + »X + +,X +,X

FIF IHC IHC ARG FIF IHC IHC FIF IHC IHC FIF IHC FIF FIF IHC

Anderson, 1983 Anderson, 1990 Anderson and Campbell, 1988 Goodrich et al., 1980 Anderson, 1983 Kiliaan éta l., 1989 Jensen and Holmgren, 1985 Watson, 1979 Holmgren, Grove and Nilsson, 1985 Kiliaan et al., 1989 Anderson, 1983 Anderson and Campbell, 1988 Watson, 1979 Anderson, 1983 Anderson and Campbell, 1988

+,X +, X +

FIF IHC ARG

Anderson and Campbell, 1984 Anderson and Campbell, 1989 Goodrich etal., 1980

Species

Stomach

CYCLOSTOMATA Lampetra pianeri, larval Lampetra fluviatilis Myxine glutinosa ELASMOBRANCHII Scyliorhinus stellaris Squalus acanthias

BRACHIOPTER YGII Polypterus senegalensis

+ +

+

TELEOSTEI Aldrichetta forsteri Anguilla australis Carassius auratus Gadus morhua Myoxocephalus scorpius Oncorhynchus mykiss Oreochromis mossambicus Platycephalus bassensis Pleuronectes platessa Salmo trutta Tetractenos glaber AMPHIBIA Bufo marinus Rana catesbeiana

+,X

+ +

+, +, +, +, +, +, + +

X X X X X X

a+ , nerve fibres; bx , ganglion cells

terminals (Grove, O’Neill and Spillett, 1974; Grove and Campbell, 1979; Jensen and Holmgren, 1985; Kitazawa, Temma and Kondo, 1986; Kiliaan et al., 1989). In the gut of the rainbow trout and the intestine of the cod, 5-HT is thought to be a mediator of the excitatory response on the motility produced by substance P (Holmgren, Grove and Nilsson, 1985; Jensen, Holmgren and Jonsson, 1987; Jensen and Holmgren, 1991; see below). In addition to its effect on motility in fish, 5-HT is also suggested to be involved in intestinal epithelial functions and to affect gastric secretion (Holstein and Cederberg, 1984; Kiliaan et al., 1989). 5-HT initiates a biphasic response on rectal preparations from the fowl, Gallus domesticus. The initial inhibition produced by stimulation of non-adrenergic inhibitory nerves is followed by a contraction involving both a direct effect on the

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smooth muscle and stimulation of excitatory non-cholinergic neurons (Mishra and Raviprakash, 1983).

PURINES ATP and adenosine have been suggested as possible transmitters in inhibitory neurons of the mammalian gastrointestinal tract. In favour of this hypothesis is the ability of the purines to mimic the effect of transmural electrical stimulation in many preparations of the gut. There are, however, several investigations where this is not the case. The lack of specific methods to localize purinergic neurons is also a problem in elucidating the role of purines in gastrointestinal function (see Burnstock, 1972, 1986; White, 1991). Studies concerning the importance of purinergic nerves in the gut of nonmammalian vertebrates is scarce and as in mammals the effect varies with species and the region of the gut. In the stomach of Lophius americanus and the rectum of Raja, ATP induces an inhibition, often followed by a rebound contraction, which mimics the response to vagal and sympathetic stimulation, respectively (Young, 1980b, 1983). Inhibitory responses to purines are also described in other species of fish. However, in several preparations, the primary effect of purines is a stimulation of the muscular activity (Holmgren, 1983; Young, 1983; Jensen and Holmgren, 1985; Lennard and Huddart, 1989). Furthermore, in the intestine of an amphibian and a lizard, it has been indicated that the excitatory non-adrenergic, noncholinergic response to stimulation of the splanchnic nerve may be purinergic (Sneddon et al., 1973). In the study by Lennard and Huddart (1989) it was concluded that two different types of receptors may be present in the ileum of the flounder, Platichtys flesus, where ATP is excitatory and adenosine is inhibitory (P2- and P r purinoceptors, respectively; for the classification of receptors see e.g. White, (1991)). So far, no general pattern can be seen in the function of purines in nonmammalian vertebrates, and it is evident that much more information is needed before conclusions can be drawn concerning a possible purinergic innervation of the gut.

NEUROPEPTIDES BOMBESIN-LIKE PEPTIDES Occurrence The few immunohistochemical studies performed on the avian gut have not been able to detect any nerve fibres containing bombesin-like peptides. Instead, numerous IR endocrine cells are demonstrated in the avian alimentary tract (Timson et al., 1979; Vaillant, Dockray and Walsh, 1979; S. Holmgren, unpublished). In

128

COMPARATIVE PHYSIOLOGY AND EVOLUTION

TABLE 4.2 Occurrence of bombesin-like immunoreactivity in nerves in the gut of non-mammalian vertebrates Species

Stomach

CYCLOSTOMATA Lampetra japónica ELASMOBRANCHII Raja clavata Raja erinacea Raja microocellata Raja montagui Raja naevus Scyliorhinus canicula Scyliorhinus stellaris Squalus acanthias Squatina aculeata

+ + + + + + + + +

HOLOCEPHALI Chimaera monstrosa HOLOSTEI Lepisosteus platyrhincus

+

DIPNOI Protopterus annectens

+

BRACHIOPTERY GII Polypterus senegalensis

+

TELEOSTEI Ciliata mustela Ctenolabrus rupestris Cyprinus carpio Gadus morhua Haplochromis sp. Leuciscus idus Myoxocephalus scorpius Oncorhynchus mykiss Perca fluviatilis Platichtys flesus Poecilia reticulata Pollachius pollachius Raniceps raninus AMPHIBIA Bufo calamita Ambystoma tigrinum Hydromates italicus Necturus maculosus REPTILIA Alligator mississipiensis Caiman crocodylus sp. Caiman latirostris Crocodylus niloticus Pogona barbatus Varanus gouldii Vipera berus

+ +

+ + + + + + + + + +

+ + + + +

Intestine

Reference

+

Yui, Nagata and Fujita, 1988

+ + + + + + + +

Bjenning and Holmgren, 1988 Bjenning, Farrell and Holmgren, 1991 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Tagliafierro et al., 1987 Cimini et al. t 1985; Tagliafierro etal.y 1989 Holmgren and Nilsson, 1983a Tagliafierro et al. y 1987

+

Yui, Shimada and Fujita, 1990 Holmgren and Nilsson, 1983b

+

Tagliafierro et al. y 1987 Burkhardt-Holm and Holmgren, 1992

+ + + + + + + + +

Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988; Holmgren and Jonsson, 1988; Thorndyke and Holmgren, 1990 Langer etal.y 1979 Burkhardt-Holm and Holmgren, 1989 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Burkhardt-Holm and Holmgren, 1989 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988

+ + + -1-

S. Holmgren, unpublished Tagliafierro etal.y 1987 Tagliafierro etal.y 1987 Holmgren etal.y 1985

+ +

Buchan, Lance and Polak, 1983 Holmgren etal.y 1989 Yamada etal.y 1987 S. Holmgren, unpublished S. Holmgren, unpublished S. Holmgren, unpublished S. Holmgren, unpublished

+ + + -1-

THE GASTROINTESTINAL CANAL

129

all other vertebrate groups, bombesin-like IR has been reported in nerves of the gut (Table 4.2), and in mammals, gastrin-releasing peptide (GRP)-and bombesin-like peptides of the gastrointestinal canal are suggested to be restricted to neurons (Dockray, Vaillant and Walsh, 1979; Iwanaga, 1983). Nerves showing bombesin-like IR commonly occur in the myenteric plexus of teleost and elasmobranch fishes (Bjenning and Holmgren, 1988; Table 4.2). In some species, such as Squalus acanthias and Myoxocephalus scorpius, ganglion cells are frequent in the enteric plexuses of the stomach, which indicates an intrinsic nature of the innervation (Holmgren and Nilsson, 1983a; Bjenning and Holmgren, 1988). Perivascular fibres are observed in submucosal arteries and major arteries to the gut, e.g. in the elasmobranchs S. acanthias and Raja erinacea; the presence of varicose fibres in the medio-adventitial border suggests an involvement of a bombesin-like peptide in the control of the vascular resistance (Holmgren and Nilsson, 1983a; Bjenning, Jonsson and Holmgren, 1990; Bjenning, Farrell and Holmgren, 1991). In the cyclostome, Lampetra japonica, a GRP-like peptide is colocalized with serotonin in intestinal neurons (Yui, Nagata and Fujita, 1988). Similarly, perivascular neurons in the gut of the elasmobranchs Scyliorhinus canicula and Squatina aculeata may contain both serotonin and bombesin (Tagliafierro et al., 1988a), but in neither case has the physiological significance of this been further investigated or discussed. No or few IR nerve fibres were detected in the gut of two lungfish species, Lepidosiren paradoxa and Protopterus annectens (Tagliafierro et al., 1987; Nilsson and Holmgren, 1992), and in the bowfin Amia calva (Rajjo, Vigna and Crim, 1989a). Also in the urodele amphibian, Necturus maculosus, and in the anuran amphibian, Bufo calamita, bombesin-IR nerve fibres are most abundant in the myenteric plexus, throughout the gastrointestinal canal (Holmgren et al., 1985; S. Holmgren, unpublished). In crocodilians (Table 4.2) the nerves are more widespread; IR fibres occur in the submucosa, myenteric plexus and circular muscle layer, and in Caiman crocodylus also in the mucosa and around mesenterial vessels (Buchan, Lance and Polak, 1983; Yamada et al., 1987; Holmgren et al., 1989, S. Holmgren unpublished). Bombesin/GRP nerves are evidently a general feature in the innervation of the gut of vertebrates, with the possible exception of birds, and some ancient fish groups. The innervation is not restricted to any special layer of the gut (although it is commonly most abundant in the myenteric plexus), suggesting that bombesin/ GRP nerves could be involved in the control of several functions such as motility, blood flow and secretion. Effects o f bombesin Maybe the most recognized effect of the bombesin-related peptides is the gastrinreleasing effect, leading to an increased secretion of gastric acid in mammals. Indeed, bombesin and GRP stimulate secretion of gastric acid in all vertebrates where they have been tested, but apparently through various mechanisms in nonmammalian species (see Jonsson, chapter 5). Bombesin and related peptides also affect gut motility, both in mammals and

130 COMPARATIVE PHYSIOLOGY AND EVOLUTION TABLE 4.3 Effects of bombesin/GRP/litorin in the non-mammalian gut. Reference

Species

Effect

Proventriculus motility

Pigeon

Inhibition of circular Gascoigne et al., 1988 muscle Gascoigne et al., 1988 Excitation of longitudinal muscle

Stomach motility

Gadus morhua (teleost)

Excitation

Holmgren and Jònsson, 1988; Thorndyke and Holmgren, 1990 Holmgren, 1983 Thorndyke et al., 1984 Holmgren et al., 1985

Excitation Oncorhynchus mykiss (teleost) Myoxocephalus scorpius (teleost) Excitation Necturus maculosus (amphibian) Excitation of longitudinal muscle Inhibition of circular Holmgren et al., 1985 muscle Intestine, rectum Squalus acanthias (elasmobranch) motility Gadus morhua (teleost)

Excitation Inhibition

Lundin, Holmgren and Nilsson, 1984 Jensen and Holmgren, 1985; Holmgren and Jònsson, 1988 Holmgren et al., 1985

Necturus maculosus (amphibian) Excitation of longitudinal muscle Inhibition of circular Holmgren et al., 1985 muscle Excitation Holmgren et al., 1989 Caiman crocodylus (reptile) Falconieri Erspamer Excitation Tortoise (reptile) et al., 1988 Erspamer et al., 1972; Excitation Chicken Falconieri Erspamer et al., 1988 Vascular flow

Squalus acanthias (elasmobranch) Caiman crocodylus (reptile) Crocodylus porosus (reptile)

Increase (stomach) Increase (gut) Increase (gut)

Bjenning, Jònsson and Holmgren, 1990 Holmgren et al., 1989 Holmgren et al., 1989

non-mammalian species (Table 4.3). In mammals, most studies have demonstrated a contractile effect of the bombesin-like peptides (e.g. Erspamer et al., 1972; Mayer, Elashoff and Walsh, 1982; Falconieri Erspamer et al., 1988), but some studies of the canine and human stomach in vivo show an inhibition of motor activity in the gut (Bertaccini et al., 1974; Melchiorri, 1978). The mechanism behind the contraction seems to vary with the species and the region of the gut. In addition to a direct effect on the smooth muscle, an action via cholinergic nerves is observed in several cases (Erspamer et al., 1972; Mayer, Elashoff and Walsh, 1982; Hirning and Burks, 1984; Kantoh et al., 1985). The effects of bombesin-like peptides in the gut of non-mammalian vertebrates have been most studied in fish (Table 4.3). Binding sites for bombesin have been demonstrated in the stomach of the teleost Scorpaeichthys marmoratus (Vigna and Thorndyke, 1989). An excitatory effect is demonstrated on preparations of smooth muscle from the rectum of the spiny dogfish, Squalus acanthias, and from the

THE GASTROINTESTINAL CANAL

131

stomach of several teleosts (Table 4.3). In the stomach of the cod, Gadus morhua, bombesin and litorin (Figure 4.3) are significantly more potent than GRP in producing a contraction, indicating that the receptors react more easily with the amphibian peptides than with the mammalian counterpart (Holmgren and Jônsson, 1988). When bombesin is added before acetylcholine, it potentiates the contraction produced by acetylcholine on preparations of stomach from the rainbow trout (Salmo gairdneri - Oncorhynchus mykiss), Atlantic cod (Gadus morhua) and daddy sculpin (Myoxocephalus scorpius) (Figure 4.3; Thorndyke et al., 1984; Thorndyke and Holmgren, 1990). This effect is much larger than the stimulatory effect produced by bombesin alone. The potentiation is unaffected by treatment with tetrodotoxin, indicating that the interaction between bombesin and acetylcholine is at the synaptic site on the smooth muscle. Bombesin may also induce a release of acetylcholine from cholinergic nerve terminals by a mechanism independent from the sodium channels, similar to the mechanism suggested to be present in the stomach of the guinea-pig by Kantoh et al. (1985). When bombesin is added to vascularly perfused preparations of the intestine of the cod, it produces a weak relaxation of the intestinal wall (Jensen and Holmgren, 1985). Similarly, bombesin and GRP only occasionally produce a relaxation of strips of smooth muscle taken from the intestinal wall, but a consistent inhibitory response is obtained with the related peptide litorin (Figure 4.3; Holmgren and Jônsson, 1988). An inhibitory effect of bombesin is also demonstrated on strips of circular muscle prepared from the stomach and intestine of the amphibian Necturus maculosus and from the proventriculus of the pigeon, while preparations of longitudinal muscle from the stomach of Necturus and the proventriculus of the pigeon respond with a contraction (Figure 4.4; Holmgren et al., 1985; Gascoigne et al., 1988). In thp intestine of the tortoise and the caiman (Caiman crocodylus) and in the intestine and caecum of the chicken, bombesin has a contractile effect, in agreement with mammalian studies in vitro (Erspamer et al., 1972; Falconieri Erspamer et al., 1988; Holmgren et al., 1989). Bombesin may be involved in the control of the distribution of blood to the stomach. Thus, in the caiman, bombesin increases the flow of blood to the gut, bu! decrease the flow through the lung (Holmgren et al., 1989; Figure 4.2). Similarly, in the spiny dogfish, bombesin increases the flow of blood through the vascularly perfused isolated stomach in vitro, and increases the flow to the gut in vivo (Bjenning, Jônsson and Holmgren, 1990; S. Holmgren and M. Axelsson, unpublished). In conclusion, bombesin-like peptides have been isolated from the gut of nonmammalian species (see Holmgren and Jensen, chapter 2) and there is immunohistochemical evidence that the bombesins are contained in neurons (Table 4.2). Pharmacological and physiological studies suggest an involvement in secretory events, and the control of motility and blood flow (Table 4.3). The most generally observed effect of bombesin-like peptides on the gut motility is a contraction, which may be a direct effect on the smooth muscle, and in some species may involve an

LITORIN Stomach

cone Intestine

ACETYLCHOLINE/BOMBESIN

FIGURE 4.3 The bombesin-related peptide litorin is excitatory on stomach smooth muscle (A), but inhibitory on intestinal preparations (B) from the Atlantic cod, Gadus morhua. (C) Bombesin potentiates the response to acetylcholine in strips of smooth muscle from the stomach of the rainbow trout, Oncorhynchus my kiss. Reproduced with permission from Holmgren and Jonsson, 1988; Thorndyke and Holmgren, 1990. Ach, acetylcholine; Bomb, bombesin; w, washout.

FIGURE 4.4

CL

CC

10 min

I

The opposite effects on longitudinal (CL) and circular (CC) smooth muscle from the stomach of the amphibian Necturus maculosus by bombesin (A) and neurotensin (B). The x-axis shows log molar concentrations of agonist.

NEUROTENSIN

BOMBESIN

THE GASTROINTESTINAL CANAL 133

134 COMPARATIVE PHYSIOLOGY AND EVOLUTION

interaction with the cholinergic innervation. Interestingly, there are also indications of antagonistic mechanisms, as in longitudinal versus circular muscle of Necturus and pigeon (Figure 4.4), and stomach versus intestine of the cod (Figure 4.3), possibly due to the presence of more than one type of receptor. The release of bombesin from autonomic nerves may increase the flow of blood to the gut, possibly during the postprandial digestion (Figure 4.2). Bombesin is a potent secretagogue (see Jonsson, chapter 5) and may also be involved in the control of feeding and satiety (Thorndyke, 1989). GASTRIN/CHOLECYSTOKININ-LIKE PEPTIDES Occurrence In the mammalian gut, gastrin/cholecystokinin (CCK)-like IR has been demonstrated in enteric nerves, in addition to endocrine cells (as established earlier). The main form present in the enteric nerves is CCK8 (Dockray, 1977; Schultzberg et al., 1980; see also Holmgren and Jensen, chapter 2). In non-mammalian vertebrates, most immunohistochemical studies of the distribution of gastrin and CCK in the gut have used antisera against their common C-terminal part. In these studies it has therefore not been possible to unequivocally discriminate between different peptides of the gastrin/CCK family. There are comparatively few reports on the occurrence of gastrin/CCK-containing nerves in non-mammalian vertebrates. This may be because interest has been focused on the presence of gastrin/CCK in endocrine cells, but it possibly reflects a sparse innervation. Nerve fibres have been demonstrated in the gut of some of the elasmobranch and teleost fish examined (Table 4.4). The distribution varies between species. In four species of the family Gadidae, IR fibres are abundant in the myenteric plexus of the stomach, but absent in the intestine and rectum (Bjenning and Holmgren, 1988). In Gadus morhua, this follows the levels of IR in the muscle layers measured by radioimmunoassay (Jonsson, Holmgren and Holstein, 1987). In contrast to the Gadidae species, fibres were few in the stomach of the elasmobranch, Squalus acanthias, but more frequent in the intestine and most abundant in the rectum; again this agrees with the levels of IR measured by radioimmunoassay (Holmgren and Nilsson, 1983a; Aldman et al., 1989). In the amphibians, Necturus maculosus and Bufo calamita, and in the reptile, Varanus gouldii, a few IR nerves are detected in the stomach and in Necturus also in the rectum (Holmgren et al., 1985; S. Holmgren, unpublished). There have been no reports to our knowledge on the presence of IR nerves in the gut of birds. Studies on the gut of invertebrates have demonstrated that a gastrin/CCK-like innervation is not restricted to the vertebrates (Grimmelikhuijzen, Sundler and Rehfeld, 1980; Andriés and Tramu, 1985). The available reports taken together give a picture of an innervation without any obvious evolutionary trends, with scattered occurrences in different organs and in different species. Clearly, more studies are needed to establish whether this is a true species difference, or whether the use of unspecific antisera have given this impression.

THE GASTROINTESTINAL CANAL

135

TABLE 4.4 Occurrence of gastrin/CCK-like immunoreactivity in nerves in the gut of non-mammalian vertebrates. Species ELASMOBRANCHII Scyliorhinus stellaris Squalus acanthias HOLOCEPHALI Chimaera monstrosa DIPNOI Lepidosiren paradoxa Protopterus annectens TELEOSTEI Ciliata mustela Cyprinus carpió Gadus morhua Leuciscus idus Myoxocephalus scorpius Pollachius pollachius Raniceps raninus AMPHIBIA Necturus maculosus Bufo calamita REPTILIA Varanus gouldii

Stomach

Intestine

Reference

+ +

-H

Cimini étal., 1985 Holmgren and Nilsson, 1983a

+

Yui, Shimada and Fujita, 1990

+

Nilsson and Holmgren, 1992 Nilsson and Holmgren, 1992

+ + + + + +

+ + +

Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Jonsson, Holmgren and Holstein, 1987; Bjenning and Holmgren, 1988 Burkhardt-Holm and Holmgren, 1989 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988

+ +

Holmgren etal., 1985 S. Holmgren, unpublished

+

S. Holmgren, unpublished

Effects o f gastrin/CCK and related peptides CCK is, in general, excitatory on preparations from the intestine and stomach of mammals in vitro. A contractile effect of CCK is also found on the pyloric sphincter, while the lower oesophageal sphincter is relaxed. Inhibitory effects have been demonstrated in some studies in vivo, e.g. of the canine stomach (see Williams, 1982; Walsh, 1987). Both indirect and direct effects of CCK on the smooth muscle are reported. In the ileum of the guinea-pig, CCK releases acetylcholine from the myenteric plexus (Vizi et al., 1973), and there is some evidence for a release of substance P by CCK (Hutchison and Dockray, 1981). Experiments with the vascularly perfused intestine of the guinea-pig suggest that CCK is involved in the peristaltic reflex, as distension of the intestine, which initiates peristalsis, results in an increased outflow of CCKlike IR material into the perfusate (Donnerer et al., 1985). In the mudpuppy, Necturus maculosus, gastrin-17 and pentagastrin have in most cases weak or inconsistent effects on the smooth muscle of the gut (Table 4.5). Exceptions are the longitudinal intestinal muscle and circular muscle of the cardiac stomach, which respond to pentagastrin with a more consistent increase in tonus and rhythmic activity. Furthermore, gastrin-17 is inhibitory on the circular muscle and excitatory on the longitudinal muscle from the pyloric stomach (Holmgren et al., 1985). Caerulein, CCK and gastrin all have an excitatory effect on gastrointestinal preparations from the cod (Jonsson, Holmgren and Holstein, 1987), and on the intestine and rectum of the spiny dogfish (Aldman et al., 1989). In three

136 COMPARATIVE PHYSIOLOGY AND EVOLUTION TABLE 4.5 Effects of neuronal (?) gastrin/CCK/caerulein on gut motility in non-mammalian species.

Stomach motility

Intestine, rectum motility Gastric emptying Gallbladder motility

Species

Effect

Reference

Raja sp. (elasmobranch)

Andrews and Young, 1988

Gadus morhua (teleost)

Excitation (longitudinal muscle) Excitation

Necturus maculosus (amphibian) Squalus acanthias (elasmobranch) Raja sp. (elasmobranchs) Gadus morhua (teleost)

Excitation Excitation Excitation Excitation

Necturus maculosus (amphibian) Oncorhynchus mykiss (teleost)

Excitation Delay

Oncorhynchus mykiss (teleost)

Excitation

Lepomis macrochirus (teleost) Fundulus heteroclitus (teleost) Amia calva (holostean)

Excitation Excitation Excitation

Jonsson Holmgren and Holstein, 1987 Holmgren etal., 1985 Aldman et al., 1989 Andrews and Young, 1988 Jonsson Holmgren and Holstein, 1987 Holmgren et al., 1985 G. Aldman, S. Holmgren and D.J. Grove, unpublished Aldman and Holmgren, 1987; Aldman, Grove and Holmgren, 1992 Rajjo, Vigna and Crim, 1988 Rajjo, Vigna and Crim, 1988 Rajjo, Vigna and Crim, 1988

species of Raja, pentagastrin produces an increase in tonus and rhythmic activity of longitudinal muscle from the intestine and the cardiac stomach (Andrews and Young, 1988). In the cod stomach, caerulein was the most potent of the peptides, followed by sulphated CCK8, CCK8, gastrin 17 and gastrin 5, and it is possible that the endogenous peptide demonstrated by immunohistochemistry resembles caerulein and CCK more than gastrins (Jonsson, Holmgren and Holstein, 1987). This may be compared with observations in mammals where CCK rather than gastrin is considered the neuronal peptide. In the cod, caerulein produces an increased rhythmic activity in circular preparations from the whole gut (except the cardiac stomach). Longitudinal preparations always respond with an increase in tonus, sometimes with an increased activity superimposed. This could indicate differences in the populations of receptors, but a more likely explanation is that this reflects a difference in the character of circular and longitudinal muscle in response to an excitatory stimulus (Jonsson, Holmgren and Holstein, 1987). In the elasmobranch, Squalus acanthias, the responses to different gastrin/CCKs are much more irregular than in the cod. It is possible that the endogenous gastrin/CCK-like peptide differs more from the mammalian peptides used in the experiments than those in the cod, which could explain the weak responses in the elasmobranch. However, the effect, when present, is always an increase in tonus and/or rhythmic activity as in other vertebrates (Aldman et al., 1989). Preliminary studies show a reduction in the blood flow to the gut after injection of sulphated CCK8 and caerulein in the cod in vivo (J. Gunnarsson, unpublished). The effects of gastrin/CCKs on secretory events in the gut are discussed by Jonsson in chapter 5).

THE GASTROINTESTINAL CANAL

137

As in mammals, the most common effect of gastrin/CCK on the motility of isolated preparations of gut wall from non-mammalian vertebrates is excitatory. However, preliminary studies in the rainbow trout, in vivo, where the transport of food was studied by X-ray, suggest that injections of CCK to the abdominal cavity delay gastric emptying (G. Aldman, S. Holmgren and D.J. Grove, unpublished). This agrees with observations in some mammals in vivo (see Walsh, 1987). An integrated effect of CCK on the processing of food in fish may be discerned: CCK stimulates the gallbladder to release bile once food enters the intestine (Aldman, Grove and Holmgren, 1992); at the same time further release of food from the stomach is delayed, although mechanical mixing of the food within the stomach may well continue. NEUROPEPTIDE Y AND RELATED PEPTIDES Due to their structural similarities, neuropeptide Y (NPY), pancreatic polypeptide and peptide YY (PYY) are treated under the same heading. In the digestive tract of mammals, NPY occurs primarily in nerves, while PYY and pancreatic polypeptide are located predominantly in endocrine cells. In addition to its presence in perivascular neurons, NPY has been identified in non-vascular nerves of the gut in several mammalian species. NPY commonly occurs co-localized with other regulatory peptides and/or adrenergic transmitters (Potter, 1988; Morris, 1989; Taylor and Bywater, 1989). NPY-IR nerve fibres have been observed in the gut of both teleosts and elasmobranchs, notably in Raja species (Table 4.6). As in mammals, the nerve fibres are present in the muscle layers and in the nerve plexuses, as well as surrounding blood vessels (Bjenning and Holmgren, 1988; Bjenning, Driedzic and Holmgren, 1989; Burkhardt-Holm and Holmgren, 1989). Among teleosts, the innervation is never abundant; most fibres are present in the stomach and only occasional fibres are observed in the intestine and rectum. In contrast, in species of Raja there is an even distribution throughout the gut. In the lizard, Varanus gouldii, fibres are abundant in all layers of the gut except the mucosa, while in the viper, Vípera berus, the IR fibres are restricted mainly to the myenteric plexus (S. Holmgren, unpublished). In the frog, Rana temporaria, NPY-IR fibres are common in the circular muscle layer and myenteric plexus of the intestine and rectum, but are absent from the stomach (McKay et al., 1992). Nerve fibres containing a PYY-like peptide are found throughout the gut in the lizard, Lacerta viviparus, and the frog, Rana temporaria. The density is highest in the submucosal layer and, in the frog, the fibres are usually associated with blood vessels (Böttcher et al., 1985). The PYY-IR fibres in Rana are possibly separate from the NPY-IR fibres (cf McKay et al., 1992). PYY-like IR is also reported in the gut of the teleost, Coitus (Myoxocephalus) scorpius, in nerves confined to the pyloric part of the stomach (El-Salhy, 1984a). Not much is known of the effects of NPY in the non-mammalian gut. In the dogfish, Squalus acanthias, injection of NPY decreases the resistance in the vascular bed of the coeliac artery, thereby increasing the flow to the gut (Holmgren, Axelsson

138

COMPARATIVE PHYSIOLOGY AND EVOLUTION

TABLE 4.6 Occurrence of NPY-like immunoreactivity in nerves in the gut of non-mammalian vertebrates. Species ELASMOBRANCHII Raja erinacea Raja radiata Raja clavata Raja microocellata Raja montagui Raja naevus TELEOSTEI Carassius auratus Ciliata mustela Cyprinus carpio Gadus morhua Myoxocephalus scorpius Oncorhynchus mykiss Perca fluviatilis Platichtys flesus Poecilia reticulata Pollachius pollachius Raniceps raninus REPTILIA Varanus gouldii Vipera berus

Stomach

Intestine

Reference

+ + + + + +

+ + + + + -1-

Bjenning, Driedzic and Holmgren, 1989 Bjenning, Driedzic and Holmgren, 1989 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988

+ + + +

+ +

+ + + + + +

A. Kiliaan and S. Holmgren, unpublished Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Burkhardt-Holm and Holmgren, 1989 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988

+ +

+ +

S. Holmgren, unpublished S. Holmgren, unpublished

+ + + + +

and Farrell, 1992). It is notable that this effect is opposite to the effect of adrenaline, which increases the resistance in the coeliac artery. NEUROTENSIN Occurrence In mammals, neurotensin-like IR is reported principally in endocrine cells of the intestine, but the occurrence in enteric nerves has also been confirmed (Schultzberg et al., 1980; Buchan and Barber, 1987; see also Carraway and Reinecke, 1984, 1989). Amongst the non-mammalian vertebrates, neurotensin-like IR is found in nerve fibres in all layers of the gastrointestinal canal of the amphibian, Necturus maculosus, in the gut (especially in the myenteric plexus and the circular muscle layer) of several reptiles and in the crop, oesophagus, proventriculus and gizzard of the chicken. Gastrointestinal nerves that are IR to neurotensin antisera have also been demonstrated in the myenteric plexus and circular muscle of holostean and teleost fishes, while no such nerves have been demonstrated in elasmobranchs (Table 4.7; elasmobranchs: Holmgren and Nilsson, 1983a; Bjenning and Holmgren, 1988). It seems to be a general pattern, that neurotensin-IR nerves are especially common in the myenteric plexus and circular muscle, suggesting a role of a neurotensin-like peptide in the control of motility.

THE GASTROINTESTINAL CANAL

139

TABLE 4.7 Occurrence of neurotensin-like immunoreactivity in nerves in the gut of non-mammalian vertebrates. Species HOLOSTEI Lepisosteus platyrhincus DIPNOI Lepidosiren paradoxa Protopterus annectens TELEOSTEI Barbus conchonius Carassius auratus Centrolabrus exoletus Ctenolabrus rupestris Gillichtys mirabilis Haplochromis sp. Helostoma temminicki Labrus berggylta Labrus mixtus Leuciscus idus Myoxocephalus scorpius Oncorhynchus mykiss Oreochromis mossambicus Perca fluviatilis Platichtys flesus Platypoecilus variatus Poecilia reticulata AMPHIBIA Necturus maculosus REPTILIA Caiman crocodylus sp. Pogona barbatus Varanus gouldii Vipera berus AVES Chicken

Stomach

Intestine

Reference

+

+

Holmgren and Nilsson, 1983b

+ +

+

Nilsson and Holmgren, 1992 Nilsson and Holmgren, 1992

+ + + + + + + + + + +

Rombout and Reinecke, 1984 Kiliaan et al., 1992 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Van Noorden and Patent, 1980 Langer eta l., 1979 Langer et al., 1979 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Burkhardt-Holm and Holmgren, 1989 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Kiliaan et al., 1992 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Langer et al., 1979 Langer éta l., 1979; Burkhardt-Holm and Holmgren, 1989

+

+ + + +

+ + + +

+

+

Holmgren et al., 1985

+ + + -1-

+

S. S. S. S.

+

(+ ) +

Holmgren, Holmgren, Holmgren, Holmgren,

unpublished unpublished unpublished unpublished

Saffreÿ, Polak and Burnstock, 1982

Effects o f neurotensin The effect of neurotensin on the motility of mammalian gut has been studied mainly in rats, guinea-pigs and dogs. Neurotensin produces excitatory, inhibitory or biphasic responses depending on the species, the part of the gut and the type of muscle examined. Direct effects on the smooth muscle as well as effects via other enteric neurons have also been reported (Kitabgi, 1982; Fox et al., 1987; Karaus et al., 1987). In the chicken, neurotensin contracts preparations from the crop, the ileum, the caecum and the rectum (Komori, Fukutome and Ohashi, 1986; Denac and Scharrer, 1987; Rawson, Duke and Brown, 1990) (Table 4.8). In the crop of the chicken, the excitation is unaffected by treatment with atropine, indomethacin, naloxone or tetrodotoxin, indicating a direct effect on the smooth muscle cells (Denac and Scharrer, 1987). It is notable that infusion of avian neurotensin in vivo did not produce the same response as studies in vitro; the ileum was relaxed, and the caecum

140 COMPARATIVE PHYSIOLOGY AND EVOLUTION TABLE 4.8 Effects of neurotensin in non-mammalian tissues.

Crop motility Stomach motility

Intestine, rectum motility

Species

Effect

Reference

Chicken

Excitation

Denac and Sharrer, 1987

Holmgren, 1983 Inhibition + excitation Oncorhynchus mykiss (teleost) Necturus maculosus (amphibian) Excitation of longitudinal Holmgren éta l., 1985 muscle Holmgren étal., 1985 Inhibition of circular muscle Necturus maculosus (amphibian) Excitation of longitudinal Holmgren étal., 1985 muscle Holmgren étal., 1985 Inhibition of circular muscle J. Jensen, unpublished Excitation Caiman crocodylus (reptile) J. Jensen, unpublished Excitation Vipera berus (reptile) Komori, Fukutome and Ohashi, Excitation Chicken, rectum 1986 Rawson, Duke and Brown, Excitation intestine, in vitro 1990 Rawson, Duke and Brown, Relaxation intestine, in vivo 1990)

and colon unaffected (Rawson, Duke and Brown, 1990). The estimated concentration of neurotensin reaching the receptors in the infusion experiments was below the concentration range producing the contraction in vitro in the chicken, but in the same range as that initiating contractile activity in the human gut. The presence of indirect (neurogenic or hormonal) effects of neurotensin, possibly differing from those in mammals, was postulated. Neurotensin is excitatory on the intestine of the caiman, Caiman crocodylus sp. and the viper, Vipera berus, again probably by a direct effect on the smooth muscle because the response is not altered after incubation with tetrodotoxin (J. Jensen, unpublished). In the gut of Necturus maculosus the dominating effect of neurotensin is opposite in the two muscle layers. Longitudinal muscle responds with an increase in tonus and/or rhythmic activity, while the rhythmic activity of preparations of circular muscle is abolished when exposed to neurotensin (Figure 4.4; Holmgren et al., 1985). The excitation of the longitudinal muscle is, as in chicken, caiman and viper, insensitive to tetrodotoxin, and therefore neurotensin probably acts directly on the smooth muscle. The inhibitory effect of neurotensin on the circular muscle layer is, however, not present after pretreatment with tetrodotoxin, indicating that the inhibition is an indirect effect (S. Holmgren, unpublished). In the stomach of the rainbow trout another type of response is observed. Superfused preparations either respond with a weak relaxation, or are unaffected during the exposure to neurotensin, while a contraction, maybe, “rebound” in nature, occurs after the period of exposure (Holmgren, 1983). Thus, the most commonly observed effect of neurotensin in non-mammalian species is a contraction (Table 4.8), probably caused by a direct effect on the smooth

THE GASTROINTESTINAL CANAL

141

TABLE 4.9 Occurrence of opioid peptide-like immunoreactivity in peripheral nerves in non-mammalian vertebrates. Species DIPNOI Lepidosiren paradoxa Protopterus annectens BRACHIOPTER YGII Polypterus senegalensis TELEOSTEI Anguilla anggilla Barbus conchonius Carassius auratus Centrolabrus exoletus Corydoras schultzei Ctenolabrus rupestris Gadus morhua Gillichthys mirabilis Helostoma temminicki Labrus berggylta Labrus mixtus Myoxocephalus scorpius Oreochromis mossambicus Pelmatochromis pulcher Perca fluviatilis Platichtys flesus Poecilia reticulate AMPHIBIA Bufo calamita Bufo marinus Rana temporaria Ambystoma mexicanum Necturus maculosus REPTILIA Caiman crocodylus sp. AVES Parus cristatus Chicken

Stomach

Intestine

Reference

+

Nilsson and Holmgren, 1992 Nilsson and Holmgren, 1992

(+ )

+

Burkhardt-Holm and Holmgren, 1992

+

+

Van Noorden and Falkmer, 1980; Bjenning and Holmgren, 1988 Rombout and Reinecke, 1984 Kiliaan e ta l., 1993 Bjenning and Holmgren, 1988 Langer eta l., 1979 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Van Noorden and Patent, 1980 Langer eta l., 1979 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Kiliaan eta l., 1993 Langer eta l., 1979 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Burkhardt-Holm and Holmgren, 1989

+

+

+ + + + +

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

(+ ) +

+

S. Holmgren, unpublished

+

+ +

S. Holmgren, unpublished Saffrey, Polak and Burnstock, 1982

+

)

S. Holmgren, unpublished Osborne and Gibbins, 1988 Buchan, 1986 Buchan 1986 Holmgren etal., 1985

(+ + + + (+

(B-enda) (B-end) )

B-enda, /3-endorphin

muscle. However, as in mammals, there are examples of other types of responses, such as the indirect effect causing inhibition in Necturus maculosus (Figure 4.4), and the relaxation in vivo of the chicken ileum. The combined effects are compatible with the view that neurotensin may be controlling a simultaneous phase of shortening and widening of the intestine. OPIOID PEPTIDES Occurrence Immunohistochemical studies, in most cases performed with antibodies raised against [Met]enkephalin, have demonstrated IR nerve fibres in the gastrointestinal

142 COMPARATIVE PHYSIOLOGY AND EVOLUTION

canal of the dipnoan, Lepidosiren paradoxa, the holostean, Polypterus senegalensis, and in several species of teleost (Table 4.9). The innervation is particularly prominent in members of the teleost family Labridae, the perch, Perea fluviatilis, and the eel, Anguilla, where numerous IR ganglion cells are also observed in the enteric plexuses. On the other hand, in cyclostomes and elasmobranchs no IR nerves have been found, using the same antiserum that shows IR in teleosts (Bjenning and Holmgren, 1988). The absence of an opioid innervation in the gut of these species may represent an earlier evolutionary stage, but it is also possible that the antiserum used is unable to interact with a cyclostome or elasmobranch opioid. Indeed, studies in invertebrates indicate that the presence of opioid-like neuropeptides in the gut is an early evolutionary feature (Van Noorden et al., 1980; Andries and Tramu, 1985; Leake, Crowe and Burnstock, 1986). Enteric nerves displaying [Met]enkephalin-like IR are present in the gut of both the anuran amphibians, Bufo calamita and Rana temporaria, and the urodele amphibian Necturus maculosus. In addition, the intestine of Rana temporaria and Ambystoma mexicanum contains nerves that react with a /3-endorphin antiserum (Holmgren et al., 1985; Buchan, 1986; S. Holmgren unpublished). [Met]enkephalin-IR nerve fibres have so far been demonstrated in only one reptile, Caiman crocodylus, although several species have been examined. The fibres were confined to the myenteric plexus, where some IR neuronal cell bodies were also observed, and to the submucosal layer (S. Holmgren unpublished). [Met]enkephalin-like IR is also present in enteric nerves of the chicken, especially in the myenteric plexus and in the circular muscle layer of the gizzard (Saffrey, Polak and Burnstock, 1982), and in nerves in the gut of the crested tit, Parus cristatus (Figure 4.5A; S. Holmgren unpublished). More extensive studies have been performed on the mammalian enteric nervous system and IR to several opioid peptides has been detected in the different layers of the gut wall (Taylor and Bywater, 1989). Effects o f opioids Enkephalins hyperpolarize neurons of the myenteric plexus in the ileum of the guinea-pig (North, Katayama and Williams, 1979). The inhibitory action of opioid peptides on gut motility is at least partly due to a reduction in the release of acetylcholine and substance P from enteric neurons (Paton, 1957; Waterfield et al., 1977; Gintzler and Scalisi, 1982; Holzer, 1984). However, excitatory effects of opioids and direct effects on the gastrointestinal muscle cells have also been reported, depending on the species and tissue studied (see Corder and Rees, 1981; Dockray, 1987). Dose-dependent contractions of circular and longitudinal muscle produced by [Met]enkephalin have been obtained in isolated preparations of the gut from the mudpuppy, Necturus maculosus, the stomach of the rainbow trout, Oncorhynchus mykiss, and the intestinal bulb of the carp, Cyprinus carpio (Table 4.10). In the carp, the excitation was found to be mediated through an action on opiate receptors situated on the smooth muscle cells (Kitazawa et al., 1986). It was also found that dynorphin was more potent than [Met]enkephalin, which in turn was more potent

THE GASTROINTESTINAL CANAL

143

FIGURE 4.5 (A) Enkephalin-like immunoreactivity in the myenteric plexus of the duodenum of the crested tit, Parus cristatus. x360. (B) Substance P-like immunoreactivity in the myenteric plexus of the stomach of the Atlantic cod, Gadus morhua. xl25.

than [Leu]enkephalin in producing an effect, and that naloxone selectively reduced the responses (Kitazawa, Hoshi and Chugun, 1990). A more complex response to [Met] enkephalin was obtained in vascularly perfused preparations of the intestine of the cod, involving both inhibitory and excitatory phases (Jensen and Holmgren, 1985). It is impossible to speculate about an evolutionary trend in the effect of opioids on gut motility. The studies in mammals indicate a potentially large diversity in the effects, with the dominating features being the indirect inhibitory action and a direct excitatory action. The information from non-mammalian species is, indeed, fragmentary, but essentially agrees with the effects in mammals. TABLE 4.10 Effects of opioid peptides in non-mammalian tissues.

Stomach motility Intestine motility

Species

Effect

Reference

Oncorhynchus mykiss (teleost) Necturus maculosus (amphibian)

Excitation Excitation

Holmgren, 1983; Holmgren et al., 1985

Gadus morhua (teleost)

Excitation/inhibition

Cyprinus carpio (teleost) Necturus maculosus (amphibian)

Excitation Excitation

Jensen and Holmgren, 1985; Kitazawa et al., 1986 Holmgren et al., 1985

144 COMPARATIVE PHYSIOLOGY AND EVOLUTION

SOMATOSTATIN Occurrence Several studies have shown somatostatin-like IR in both endocrine cells and nerve fibres in the digestive tract of mammals (e.g. Polak and Bloom, 1986; Taylor and Bywater, 1989). Information about the occurrence of somatostatin or somatostatinlike peptides in the enteric nerves of non-mammalian species is, however, restricted to comparatively few investigations (Table 4.11). In the alimentary tract of Squalus acanthias, IR fibres are most frequent in the rectum, and in another elasmobranch, Scyliorhinus stellaris, nerves are present in the stomach, being most numerous in the distal fundus and the pylorus (Holmgren and Nilsson, 1983a; Cimini et al., 1985). A somatostatin-like peptide is present in scattered nerve fibres in the intestine of the teleost, Barbus conchonius, and in the gut of the amphibian, Rana ridibunda, while only weakly IR gastrointestinal nerves are detected in the mudpuppy, Necturus maculosus (Rombout and Reinecke, 1984; Holmgren et al., 1985; Junquera et al., 1986, 1987a,b, 1988). Immunohistochemical evidence also indicates the localization of somatostatin-like IR in enteric nerves of some reptilian species (Buchan, Lance and Polak, 1983; S. Holmgren unpublished). Effects o f somatostatin on gut motility In vitro studies of intestinal preparations from guinea-pig and rabbit have indicated an inhibitory action of somatostatin on spontaneous contractions, or on contractions produced by field stimulation. This inhibition is suggested to be due to a reduction of the release of acetylcholine from enteric nerves, and by an activation of inhibitory neurons (Guillemin, 1976; Cohen et al., 1978; Furness and Costa, 1979).

TABLE 4.11 Occurrence of somatostatin-like immunoreactivity in nerves in the gut of non-mammalian vertebrates. Species ELASMOBRANCHII Scyliorhinus stellaris Squalus acanthias DIPNOI Lepidosiren paradoxa Protopterus annectens TELEOSTEI Barbus conchonius AMPHIBIA Bufo marinus Rana ridibunda Necturus maculosus REPTILIA Alligator mississippiensis Crocodylus niloticus Varanus gouldii

Stomach

Intestine

Reference

+ +

+

Cimini etal.y 1985 Holmgren and Nilsson, 1983a

+

+ - 1-

+

+

Nilsson and Holmgren, 1992 Nilsson and Holmgren, 1992

+

Rombout and Reinecke, 1984; Abad etal.y 1987

+ + +

Osborne and Gibbins, 1988 Junquera etal.y 1986, 1987a,b Holmgren etal.y 1985

+ + +

Buchan, Lance and Polak, 1983 S. Holmgren, unpublished S. Holmgren, unpublished

THE GASTROINTESTINAL CANAL

145

Somatostatin also produces a relaxation, sometimes followed by a contraction, on intestinal preparations from the cod, Gadus morhua, while the effect on preparations of stomach from the rainbow trout, Oncorhynchus mykiss, is excitatory (Holmgren, 1983; Jensen and Holmgren, 1985). However, in the isolated perfused stomach of both cod and rainbow trout, somatostatin reduces the increased activity initiated by distension of the stomach wall in a manner similar to that of atropine, and it was suggested that somatostatin inhibits the excitatory cholinergic neurons of the stomach wall, as in mammals (Grove and Holmgren, 1992a, b). SUBSTANCE P AND OTHER TACHYKININS Occurrence Substance P-like peptides (tachykinins) have been demonstrated in the enteric nervous system of representatives of all the vertebrate groups, except cyclostomes (Table 4.12). In many cases IR ganglion cells are observed in the enteric plexuses, indicating that at least part of the fibres are truly enteric, but an extrinsic origin of other populations of substance P fibres has been established. Thus, in mammals, substance P commonly occurs in sensory neurons, often together with calcitonin gene-related peptide (CGRP), and in populations of extrinsic autonomic neurons (Bartho and Holzer, 1985; Furness and Costa, 1987). Nerve sectioning and tracing studies are, however, few in non-mammalian species, and it is not clear to what degree the observed populations of nerves are truly enteric, sensory or extrinsic autonomic in nature. With only a few exceptions, studies on the occurrence of substance P-IR nerves report a dense innervation, particularly of the enteric plexuses*. Thus, numerous IR nerves are found in all layers throughout the gut of the spiny dogfish, Squalus acanthias (Holmgren, 1985; El-Salhy, 1984b), while only a few nerve fibres, confined to the fundus, are detected in the stomach of Scyliorhinus stellaris (Cimini et al., 1985). A dense substance P-IR innervation of the gut is also present in several teleosts, with some variation in distribution between different species (Figure 4.5b; Table 4.12). Abundant nerve fibres and ganglion cells throughout the gut of the urodele amphibians, Salamandra salamandra and Necturus maculosus, show substance P-like IR, including perivascular nerve fibres (Buchan, Polak and Pearse, 1980; Holmgren et al., 1985). A variable density of IR nerves and ganglion cells is observed in the gastrointestinal tract of anuran species (Table 4.12). In Bufo marinus, there was an almost total disappearance of substance P-IR nerves in the ileum after splanchnic nerve sectioning, indicating an extrinsic origin of the neurons (Osborne and Campbell, 1986). There are few reports on nerve fibres innervating the epithelial layers of the gut; however, in the colon of Bufo, substance P-IR fibres penetrate the mucosa and reach the inner epithelium of the gut. CGRP is colocalized with substance P in these fibres, which were thought to be sensory (Murphy et al., 1990).

146 COMPARATIVE PHYSIOLOGY AND EVOLUTION TABLE 4.12 Occurrence of substance P-like immunoreactivity in nerves in the gut of non-mammalian vertebrates. Species ELASMOBRANCHII Scyliorhinus stellaris Squalus acanthias DIPNOI Protopterus annectens BRACHIOPTER YGII Polypterus senegalensis TELEOSTEI Anguilla anguilla Barbus conchonius Carassius auratus Ciliata mustela Ctenolabrus rupestris Cyprinus carpio Gadus morhua Gillichthys mirabilis Labrus berggylta Leuciscus idus Myoxocephalus scorpius Oncorhynchus mykiss Perca fluviatilis Platichthys flesus Poecilia reticulata Pollachius pollachius Raniceps raninus AMPHIBIA Ambystoma mexicanum Necturus maculosus Salamandra salamandra Bombina bombina Bufo bufo Bufo calamita Bufo marinus Rana ridibunda Rana esculenta Rana temporaria REPTILIA Alligator niississippiensis Caiman crocodylus sp. Crocodylus niloticus Podareis hispánica Pogona barbatus Varanus gouldii Vípera berus AVES Parus cristatus Coturnix coturnix sp. Chicken

Stomach

Intestine

Reference

+ +

+

Cimini et al. y 1985 El-Salhy 1984b; Holmgren, 1985 Nilsson and Holmgren, 1992

+ +

+

Burkhardt-Holm and Holmgren, 1991

+

+ + -1-

Bjenning and Holmgren, 1988 Rombout and Reinecke, 1984 Kiliaan eta l., 1992 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Jensen and Holmgren, 1985; Jensen, Holmgren and Jonsson, 1987; Bjenning and Holmgren, 1988 Van Noorden and Patent, 1980 Bjenning and Holmgren, 1988 Burkhardt-Holm and Holmgren, 1989 Bjenning and Holmgren, 1988 Holmgren, Vaillant and Dimaline 1982; Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Burkhardt-Holm and Holmgren, 1989 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988

+ +

+ + + + + + + + + +

4+ + + + + + + + + + +

+

+ + +

+ + +

+ + +

Buchan, 1986 Holmgren eta l., 1985 Buchan, Polak and Pearse, 1980 Buchan, 1986 Buchan, 1986 S. Holmgren, unpublished Osborne and Campbell, 1986; Murphy eta l., 1990 Junquera eta l., 1986, 1987a,b Gabriel, 1990 Buchan, 1986; McKay eta l., 1990

+ + + + +

+ + + + + + +

Buchan, Lance and Polak, 1983 S. Holmgren, unpublished S. Holmgren, unpublished Burrell eta l., 1991 S. Holmgren, unpublished S. Holmgren, unpublished S. Holmgren, unpublished

+ +

+ +

-I-

- 1-

S. Holmgren, unpublished Fontaine-Perus eta l., 1981 Brodin eta l., 1981; Fontaine-Perus eta l., 1981 Saffrey, Polak and Burnstock, 1982; Aisa e ta l., 1987

+ +

THE GASTROINTESTINAL CANAL

147

In the few reptilian and avian species examined, a similar situation is encountered: frequent IR nerves are present in most layers of the gut wall, and in some species IR ganglion cells are observed (Table 4.12). In mammals, the substance P innervation is present in all parts and in all layers of the gastrointestinal canal. In the guinea-pig, most neurons are thought to be intrinsic, with the exception of nerves around submucous blood vessels and nerves in the submucous ganglia. However, in cat and dog, extrinsic neurons may be more numerous (Bartho and Holzer, 1985). It can be concluded that tachykinin-containing neurons are present in representatives of phylogenetically widely separated groups, often with a widespread distribution within the gut. This indicates that the tachykinins are important in the control of several basic functions in the gut. Effects o f tachykinins A number of studies have shown that substance P affects the gastrointestinal motility of vertebrates. Early experiments on non-mammalian vertebrates showed that substance P, purified from mammalian intestine, had an excitatory effect on the caecum of the chicken and on intestinal preparations from some fish species (Pernow, 1953; Pernow and Rocha e Silva, 1955; von Euler and Ostlund, 1957; Cleugh et al., 1961). More recent results have confirmed this excitatory effect of substance P both in the gut of fish and the caecum of the chicken (Table 4.13). The mechanism of action of substance P may vary between species (Figure 4.6). A direct effect of substance P on the smooth muscle cells of the gut may be a general feature throughout the vertebrates. This may also be the only way of action in the most “primitive” groups of fishes that respond to substance P. Thus, in the gut of elasmobranchs (Sgualus acanthias and Raja species), and in the intestine of the lungfish, Lepidosiren paradoxa, and the bichir, Polypterus senegalensis, substance P-induced contractions are unaffected by treatment with the antagonists atropine, methysergide or tetrodotoxin, indicating a direct action only of substance P on the smooth muscle cells (Holmgren, 1985; Andrews and Young, 1988; Jensen and Holmgren, 1991). In most other species, additional mechanisms of action are indicated, often involving cholinergic and/or serotonergic pathways. Interestingly, the mechanisms involved seem to vary between parts of the gut, as well as between species. In experiments performed on the vascularly perfused stomach of the rainbow trout, low concentrations of substance P caused contractions that could be reduced by methysergide or tetrodotoxin, but not by atropine, suggesting that the effect is partly direct on the smooth muscle and partly via serotonergic neurons. Further support for the assumption that substance P acts partly via serotonergic neurons was gained in experiments where substance P was found to increase the outflow of label into the perfusate from preparations of stomach preloaded with [3H]-5-hydroxytryptamine, and by the fact that both substance P- and 5-HT-like IR are present in nerves of the myenteric plexus (Holmgren, Vaillant and Dimaline, 1982; Holmgren, Grove and Nilsson, 1985). In the stomach of the cod, only a direct effect on the muscle layers could be demonstrated (Jensen etal.9 1993). In contrast, results obtained with vascularly

148

COMPARATIVE PHYSIOLOGY AND EVOLUTION TABLE 4.13 Effects of tachykinins in non-mammalian tissues.

Stomach motility

Intestine motility

Species

Effect

Squalus acanthias (elasmobranch) Raja radiata (elasmobranch) Raja sp. (elasmobranch) Lepidosiren paradoxa (dipnoan) Oncorhynchus mykiss (teleost)

Excitation Excitation Excitation Excitation Excitation

Gadus morhua (teleost) Necturus maculosus (amphibian) Gadus morhua (teleost) Oncorhynchus mykiss (teleost) Cyprinus carpio (teleost) Lepidosiren paradoxa (dipnoan) Polypterus senegalensis (brachiopterygian) Necturus maculosus (amphibian) Bufo marinus (amphibian) Vipera berus (reptile) Caiman crocodylus (reptile) Chicken

Rectum motility

Chicken

Holmgren, 1985 Jensen and Holmgren, 1991 Andrews and Young, 1988 Jensen and Holmgren, 1991 Holmgren, Grove and Nilsson, 1985 Jensen etal.y 1993 Excitation Excitation/inhibition Holmgren et al.y 1985 Jensen and Holmgren, 1985; Excitation Jensen, Holmgren and Jonsson, 1987 Jensen and Holmgren, 1991 Excitation Excitation Kitazawa et al.y 1988 Jensen and Holmgren, 1991 Excitation Excitation Excitation Excitation Excitation No response Excitation Excitation

Vascular beds coeliac artery

Squalus acanthias (elasmobranch) Increased flow

coeliac artery

Gadus rhorhua (teleost)

Increased flow

mesenteric artery Gadus morhua (teleost)

Increased flow

coeliac artery

Increased flow

Crocodylus porosus (reptile)

Reference

Jensen and Holmgren, 1991 Holmgren et al., 1985 Osborne and Campbell, 1986 J. Jensen, unpublished J. Jensen, unpublished Brodin et al.y 1981; Rawson, Duke and Brown, 1990 Komori, Fukutome and Ohashi, 1986 Holmgren, Axelsson and Farrell, 1992 Jensen, Axelsson and Holmgren, 1991 Jensen, Axelsson and Holmgren, 1991 Axelsson, et al.y 1991

perfused intestine from the cod and strips of muscle from the intestine of the rainbow trout suggest that, in addition to the direct stimulatory effect on the smooth muscle, indirect actions of substance P via cholinergic and serotonergic neurons may cause an excitation (Figure 4.6; Jensen, Holmgren and Jonsson, 1987; Jensen and Holmgren, 1991). An indirect effect via enteric cholinergic neurons (and a direct effect) is also proposed in circular muscle from the intestinal bulb of the carp, Cyprinus carpio. However, in this preparation there was no indication of an involvement of serotonergic neurons in the substance P-induced contraction, and in preparations of longitudinal muscle, only a direct effect could be demonstrated (Kitazawa et al., 1988; Kitazawa, Hoshi and Chugun, 1990). The effect of substance P via cholinergic neurons, and the direct effect on the smooth muscle are similar to the mechanisms described for substance P in the ileum of the guinea-pig and dog (Rosell et al., 1977; Yau, 1978; Holzer and Lembeck, 1980; Daniel et al.9 1982).

Q

t kT?

SPINY DOGFISH

CARP FIGURE 4.6 A summary of the possible mechanisms of action for the contractile effect of tachykinins (TK) in the gastrointestinal canal of fish. Circles denote neurons and squares denote endocrine cells. The figure is based on results, by Holmgren, Vaillant and Dimaline, 1982; Holmgren, Grove and Nilsson, 1985; Holmgren, 1985; Jensen, 1990; Jensen and Holmgren, 1985, 1991; Jensen, Holmgren and Jonsson, 1987; Kitazawa et al., 1988; Burkhardt-Holm and Holmgren, 1992. ACh, acetylcholine; 5-HT, 5-hydroxytryptamine.

150 COMPARATIVE PHYSIOLOGY AND EVOLUTION

Mechanical distension of the stomach initiates or increases contractile activity in the vascularly perfused stomach of the rainbow trout in situ; this response is inhibited by tetrodotoxin, suggesting that the effect is nerve mediated (Grove and Holmgren, 1992a). Similar experiments performed in vitro show that the release of substance P-like material from the stomach wall into the vascular perfusate increases during the period of distension, which further indicates that a tachykinin is involved in the excitatory control of the stomach of the rainbow trout (Jensen and Holmgren, 1992). Excitatory effects of substance P have been demonstrated in the gut of amphibians. Strips of muscle from the alimentary canal of Necturus maculosus responded with dose-dependent contractions when exposed to substance P. In addition, an inhibitory response was occasionally obtained in longitudinal muscle of the pyloric stomach (Holmgren et al., 1985). In a study of the ileum from Bufo marinus, Osborne and Campbell (1986) reported a contractile effect of substance P that seemed to be direct on the muscle cells. Studies on the electrical activity of cells in strips of circular muscle from the stomach of the toad show that the effect of substance P is similar to that reported in mammals, an effect on the pacemaker giving a chronotropic response; however, the mechanism of influence of the action potential may vary from that in mammals (Shonnard and Sanders, 1990). Not much is known about the situation in reptiles and birds. A stimulation of the activity of muscle by substance P is observed in the caecum of the chicken, probably by a direct action on the musculature (Brodin et al., 1981). The same is true for the intestine of the viper, Vipera berus, while the intestine of the caiman, Caiman crocodylus, in most cases is unaffected by substance P (J. Jensen, unpublished). Substance P-like IR, like bombesin-like IR, occurs in mucosal and submucosal nerves as well as endocrine cells of many of the species investigated, and effects of the tachykinins, substance P, physalaemin and eledoisin, on the secretion of gastric acid and pepsin have been demonstrated in the cod (Holstein and Cederberg, 1986). This is further discussed by Jonsson in chapter 5 of this volume. Blood vessels in the gut of mammals are richly innervated by nerves containing substance P-like IR (Furness et al., 1982; Barja, Mathison and Huggel, 1983), and numerous studies have shown that tachykinins are vasodilatory in the mammalian gut (Rozsa and Varro, 1987). Among non-mammalian species, substance P-IR neurons have so far only been demonstrated in a limited number of vessels on the surface of the gut of the cod, and it is doubtful whether these nerves can have a general regulatory function on the blood flow in the gut. Still, injections of substance P into cod swimming in running sea water causes a reduced vascular resistance and an increased flow in the coeliac and mesenteric arteries, which supply the gut (Jensen, Axelsson and Holmgren, 1991). Superimposed on the increase in flow in the coeliac artery is a transient decrease, which is abolished by cholinergic blockade, but not by vagotomy; the significance of this enteric, cholinergic “reflex” is not yet understood (Figure 4.2). In the spiny dogfish, Squalus acanthias, and the crocodile, Crocodylus porosus, a prominent decrease in vascular resistance and consequently an increase in flow is obtained in the coeliac artery on injection of

THE GASTROINTESTINAL CANAL

151

substance P (Figure 4.2; Axelsson et al., 1991; Holmgren, Axelsson and Farrell, 1992). A postprandial increase in the flow of blood has been described in mammals (Fara, 1984), crocodiles (Axelsson et al., 1991) and fish (Axelsson et al., 1989; Axelsson and Fritsche, 1991). The importance of tachykinins in the postprandial hyperaemia is not known, but in the dog there is a postprandial increase in the levels of substance P-like IR material in the plasma (Akande et al.9 1981). The presence of a tachykinin innervation, and its excitatory action on the smooth muscle of the gut may be a universal feature in the animal kingdom. Among vertebrates, the direct effect of substance P (or some other tachykinin) seems to be a common mechanism of action. The indirect effects of substance P via cholinergic and/or serotonergic neurons have so far only been demonstrated in fish and in mammals, but it is possible that further studies will show that these mechanisms are present in representatives from other groups of vertebrates as well.

VASOACTIVE INTESTINAL POLYPEPTIDE Occurrence Vasoactive intestinal polypeptide (VIP) or a VIP-like peptide is widely distributed in the alimentary tract of almost all of the vertebrates studied. In mammals it is generally considered that VIP is exclusively located in nerve fibres (Dimaline and Dockray, 1978; Furness and Costa, 1987). The main location of VIP-like IR in the non-mammalian vertebrates is also in nerve fibres, but in some studies the additional occurrence of IR material in endocrine cells is indicated. Of all the neuropeptides so far described in the vertebrate gut, VIP-like peptides probably have the most widespread distribution and are present in the most dense nerve plexuses, penetrating all layers of the gut. The cyclostome, Myxine glutinosa, may look like an exception to this “rule”, with a few VIP-IR endocrine cells found in the mucosa of the intestine, but no nerve fibres reported (Reinecke et al.9 1981; Bjenning and Holmgren, 1988). However, Myxine glutinosa as a species is exceptional; so far no IR gut nerves at all have been demonstrated (Bjenning and Holmgren, 1988). Several studies have demonstrated a dense innervation with nerves containing VIP-like IR in the gut of the elasmobranchs, Squalus acanthias and Scyliorhinus stellaris (Table 4.14). Fibres are most abundant in the myenteric plexus and the submucosa, where IR ganglion cells are also observed. On the other hand, in the gut of five species of Raja, no or only scattered IR nerves could be detected (Bjenning and Holmgren, 1988; Falkmer et al., 1980). VIP-IR nerve fibres are also frequent in the gastrointestinal canal of holostean and teleost fish (Table 4.14), where they are seen in small bundles parallel to the muscle fibres and as a network of nerve fibres in the myenteric and submucous plexuses. As in fish, numerous VIP-IR nerve fibres are present in the gut of amphibian, reptile and avian species (Figure 4.7A, B; Table 4.14).

152 COMPARATIVE PHYSIOLOGY AND EVOLUTION TABLE 4.14 Occurrence of VIP-like immunoreactivity in nerves in the gut of non-mammalian vertebrates. Species ELASMOBRANCHII Raja clavata Raja radiata Scyliorhinus stellaris Squalus acanthias HOLOCEPHALI Chimaera monstrosa HOLOSTEI Lepisosteus platyrhicus Amia calva DIPNOI Lepidosiren paradoxa BRACHIOPTER YGII Polypterus senegalensis TELEOSTEI Anguilla anguilla Barbus conchonius Brachydanio rerio Carassius auratus Centrolabrus exoletus Ciliata mustela Corydoras schultzei Coitus scorpius Ctenolabrus rupestris Cyprinus carpió Gadus morhua Gillichtys mirabilis Gyrinochelius aymonieri Haplochromis sp. Helostoma temminicki Hemigrammus ocellifer Labrus berggylta Labrus mixtus Leuciscus idus Lepomis macrochirus Myoxocephalus scorpius Oncorhynchus mykiss

Stomach Intestine Reference + + + +

Falkmer e ta l., 1980 Falkmer eta l., 1980 Cimini eta l., 1985; Tagliafierro eta l., 1988b, 1989 Reinecke eta l., 1981; Holmgren and Nilsson, 1983a; El-Salhy 1984b

+

Yui, Shimada and Fujita, 1990

+ +

+ +

Holmgren and Nilsson, 1983b Rajjo, Vigna and Crim, 1989b

+

+

Nilsson and Holmgren, 1992

+

+

Burkhardt-Holm and Holmgren, 1991

+

+

Van Noorden and Falkmer, 1980; Bjenning and Holmgren, 1988 Rombout and Reinecke, 1984; Abad eta l., 1987 Langer eta l., 1979 Kiliaan eta l., 1993 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Langer eta l., 1979 Reinecke eta l., 1981 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Van Noorden and Patent, 1980 Langer eta l., 1979 Langer eta l., 1979 Langer eta l., 1979 Langer eta l., 1979 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Burkhardt-Holm and Holmgren, 1989 Rajjo, Vigna and Crim, 1989b Reinecke et al., 1981 Holmgren, Vaillant and Dimaline, 1982; Bjenning and Holmgren, 1988 Kiliaan eta l., 1993 Langer eta l., 1979 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Bjenning and Holmgren, 1988 Burkhardt-Holm and Holmgren, 1989; Langer eta l., 1979 Bjenning and Holmgren, 1988 Reinecke etal., 1981; Bjenning and Holmgren, 1988

+ +

+ + + + + +

+ + +

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

Oreochromis mossambicus Pantodon buchholzi Perca fluviatilis Platichtys flesus Pleuronectes platessa Poecilia reticulata

+ + + +

Pollachius pollachius Raniceps raninus

+ +

+ +

+ + + +

+ + + +

AMPHIBIA Alytes obstetricans Atelops oxyrhyncus Bombino bombino Bufo bufo

+ + + + +

Buchan e ta l., 1981 Buchan, 1986 Buchan, eta l., 1981 Buchan eta l., 1981

THE GASTROINTESTINAL CANAL TABLE 4.14

153

Continued

Species Bufo calamita Bufo marinus Hyla arborea Hyla crepitans Rana esculenta Rana ridibunda Rana temporaria Ambystoma mexicanum Cynops hongkongensis Necturus maculosus Salamandra salamandra Xaenopus laevis REPTILIA Alligator mississippiensis Caiman crocodylus sp. Crocodylus niloticus Lacerta viridis Podareis hispánica Pogona barbatus Varanus gouldii Vípera aspis Vípera berus AVES Turkey Coturnix coturnix sp. Parus cristatus Chicken

Stomach Intestine Reference

+ + + + +

+ + -1+ + + +

+ + + + +

+ + + + +

S. Holmgren, unpublished Osborne and Gibbins, 1988 Buchan e ta l., 1981 Buchan eta l., 1981, Buchan, 1986 Buchan eta l., 1981 Junquera eta l., 1986, 1987a,b Buchan e ta l., 1981, Diaz de Rada, Sesma and Vazquez, 1987 Buchan, 1986 Buchan, 1986 Holmgren eta l., 1985 Buchan, Polak and Pearse, 1980 Reinecke e ta l., 1981

+ +

-1-

+ + + + + + + + +

Buchan, Lance and Polak, 1983 S. Holmgren, unpublished S. Holmgren, unpublished Reinecke eta l., 1981 Burrell eta l., 1991 S. Holmgren, unpublished S. Holmgren, unpublished Masini, 1986 S. Holmgren, unpublished

+ + + +

+ + + +

Vaillant, Dimaline and Dockray, 1980 Fontaine-Perus eta l., 1981; Reinecke eta l., 1981 S. Holmgren, unpublished Fontaine-Perus etal., 1981; Reinecke eta l., 1981; Saffrey, Polak and Burnstock, 1982; Aisa etal., 1987

-H

+ + +

VIP-IR ganglion cells are common in the myenteric plexus of fish and amphibians, but sparse in the submucous plexus. This is different from mammals, where most VIP ganglia occur in the submucosa, and might be explained by the fact that the submucous plexus in fish and amphibians is not well developed, and contains only a few ganglion cells (Burnstock, 1969) In several species, nerves have been found in association with blood vessels of the gut wall. Major systemic arteries may be more densely innervated than the peripheral vascular beds. Moderately dense to dense plexuses of fibres were found in the walls of the coeliac and mesenteric arteries of the spiny dogfish, Squalus acanthias (Holmgren and Nilsson, 1983a) and the Atlantic cod, Gadus morhua (Lundin and Holmgren, 1984). Particularly prominent perivascular plexuses were observed along the small mesenterial branches of the coeliac and mesenteric arteries, and in the branches on the surface of the gut wall (S. Holmgren, unpublished), while the density of perivascular VIP fibres innervating vessels intrinsic to the gut wall was low in the species of fish investigated, such as the holostean, Lepisosteus platyrhincus, (Holmgren and Nilsson, 1983b), the cod (Jensen and Holmgren, 1985) and the rainbow trout (Holmgren, Vaillant and Dimaline, 1982). In a study of the

154 COMPARATIVE PHYSIOLOGY AND EVOLUTION

FIGURE 4.7 (A) VIP-like immunoreactivity in the myenteric plexus of the intestine of the viper, Vipera berus. x250. (B) VIP-like immunoreactivity in nerve fibres and one ganglion cell in the muscle layer of the oesophagus from the viper, Vipera berus. x300. (C) Cardiac stomach of the toad, Bufo calamita. CGRP-like immunoreactivity occurs in nerve fibres of the myenteric plexus and surrounding epithelial blood vessels. x250. (D) Galanin-like immunoreactivity in the myenteric plexus of the cardiac stomach of the cod, Gadus morhua. x300.

myenteric plexus and the muscular layers of 18 species of elasmobranchs and teleosts, it was concluded that VIP-IR fibres follow and surround the vessels of these layers of the gut to some extent only (Bjenning and Holmgren, 1988). Effects o f VIP There are reports on both inhibitory and excitatory effects of VIP on the mammalian gut; the most common effect on gut motility is an inhibition (Fahrenkrug, 1991). The effects of VIP on gut motility in non-mammalian vertebrates has been little investigated, and all of the studies were performed with mammalian VIP which differs from e.g. fish VIP in the C-terminal part (see Holmgren and Jensen, chapter 2, Table 4.15). The prevailing results of these studies do, however, indicate an inhibitory effect of VIP. In the amphibian, Necturus maculosus, VIP produces a decrease in tonus and/or abolishes the spontaneous rhythmic activity in preparations from the whole gastrointestinal canal (Holmgren et al., 1985). VIP is also inhibitory on the muscular activity of the rectum from the spiny dogfish, Squalus acanthias, causing a reduction and abolishment of rhythmic activity similar to that in the gut of Necturus (Lundin, Holmgren and Nilsson, 1984). Studies performed in vitro on teleosts yield inconsistent results. Vascularly per-

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155

TABLE 4.15 Effects of VIP in non-mammalian tissues.

Stomach motility

Intestine, rectum motility Vascular beds Intestine Coeliac artery

Mesenteric artery Gallbladder motility

Species

Effect

Reference

Gadus morhua (teleost)

Inhibition

Oncorhynchus mykiss (teleost)

Inhibition

Necturus maculosus (amphibian) Squalus acanthias (elasmobranch)

Inhibition Inhibition

Necturus maculosus (amphibian)

Inhibition

Jensen, Axelsson and Holmgren, 1991 Grove and Holmgren, 1992a Holmgren et al., 1985 Lundin, Holmgren and Nilsson, 1984 Holmgren et al., 1985

Ictalurus mêlas (teleost) Gadus morhua (teleost)

Dilatation Flow increase

Squalus acanthias (elasmobranch)

Flow decrease

Gadus morhua (teleost)

Flow increase

Oncorhynchus mykiss (teleost)

Inhibition

Holder et al., 1983 Jensen, Axelsson and Holmgren, 1991 Holmgren, Axelsson and Farrell, 1992 Jensen, Axelsson and Holmgren, 1991 Aldman and Holmgren, 1992

fused preparations of the intestine of the cod were insensitive to VIP, and in isolated preparations of the stomach wall of the rainbow trout, VIP produced variable responses (Holmgren, 1983; Jensen and Holmgren, 1985). However, in vivo injections of porcine VIP in cod abolish the rhythmic contractions of the stomach wall (Jensen, Axelsson and Holmgren, 1991), and in rainbow trout, VIP reduces the resistance to distension of the stomach wall (Grove and Holmgren, 1992a), and may thus be involved in the receptive relaxation of the stomach. In the rainbow trout, in vivo, VIP also reduces the amplitude of CCK-induced contractions of the gallbladder, possibly at least partly involving a j3-adrenoceptor-mediated pathway (Aldman and Holmgren, 1992). Electrophysiological studies in isolated strips of circular muscle from the rectum of the chicken show that VIP acts in a manner similar to that of the inhibitory transmitter released from intrinsic nerves: VIP caused a hyperpolarization of the cell membrane with the same properties as the inhibitory junction potential (Komori and Ohashi, 1990). In mammals, VIP is suggested to be the neurotransmitter in the reflex mediating gastric receptive relaxation, which follows distension of the oesophagus. VIP is also a transmitter candidate in the descending inhibition in intestinal peristalsis (see Dockray, 1987 for more information about the situation in mammals). It is possible that VIP is also a neurotransmitter in inhibitory reflexes in non-mammalian vertebrates, as indicated by the inhibitory effects on the gut of fish and Necturus. The dense innervation of vessels to the gut implies a role of VIP in the control of blood flow to the gut. In mammals, VIP has been associated with a dilation of blood vessels, including those of the gut, leading to an increased flow of blood

156 COMPARATIVE PHYSIOLOGY AND EVOLUTION

(Fahrenkrug, 1991). Virtually nothing is known about the effect of VIP on the blood flow to the gut of non-mammalian vertebrates, except for some results obtained from fish. Most studies in fish have been performed with porcine VIP and have produced variable results, but one study in the catfish, Icatalurus melas, using VIP extracted from the gut of the trout and catfish, shows a vasodilation in a perfused intestinal loop (Holder et al., 1983). Surprisingly, in the cod, Gadus morhua, injections of porcine VIP in vivo, causes an increase in flow in the mesenteric artery to the gut, which is only dependent on an increased cardiac output, while the flow in the coeliac artery is further increased by a decrease in vascular resistance due to vasodilation (Jensen, Axelsson and Holmgren, 1991). This could be explained by a more dense innervation by VIP-IR fibres of the coeliac artery than the mesenteric artery, as observed with immunohistochemistry (S. Holmgren, unpublished). However, in the elasmobranch, Squalus acanthias, porcine VIP caused an increased total vascular resistance, including the vascular bed of the coeliac artery, and the: blood flow to the gut was consequently reduced (Holmgren, Axelsson and Farrell, 1992). It is notable that in the rectal gland of Squalus, VIP caused the same effect as that described in mammalian exocrine glands: a vasodilation, combined with an increase in glandular secretion (Solomon et al., 1984; Thorndyke et al., 1989), and it is possible that true species differences exist in the effects of VIP on circulation in the gut.

OTHER TRANSMITTERS There are, in addition to the substances described above, several other candidates for transmitter in the gastrointestinal canal. The current evidence for a transmitter function of these substances in the non-mammalian vertebrates is, however, almost exclusively based on immunochemical findings. Thus, CGRP-like IR is demonstrated in the plexuses and the muscle layers of the intestine in species representing all of the major groups of vertebrates (Figure 4.7C; Yui, Nagata and Fujita, 1988; Ohtani et al., 1989; Scheuermann et al., 1991), and binding sites for CGRP have been demonstrated in the stomach of the rainbow trout (Arlot-Bonnemains et al., 1991). A galanin-like peptide is present in neurons of the myenteric and submucous plexus in the intestine of the mudpuppy, Necturus maculosus (McKeon et al., 1990). In the stomach of the cod, galanin-IR nerve fibres are found in the myenteric plexus (Figure 4.7D), in the muscle layers and surrounding blood vessels and, in addition, approximately 1 0 % of the ganglion cells along the vagal branches to the gut show a positive immunostaining. Galanin has weak excitatory effects on the motility of the muscle of the stomach wall, and slightly increases the tone of isolated preparations of the arteries to the gut (Karila et al., 1993). Finally, 7 -aminobutyrate (GABA)-IR nerve fibres have been reported in the myenteric plexus of the carp, frog and chicken (Gabriel et al., 1990). The physiological functions of these substances in the non-mammalian vertebrates however, remain to be elucidated.

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157

CONCLUDING REMARKS Immunohistochemical studies in non-mammalian vertebrates, particularly on the presence of neuropeptides, have been almost exclusively performed with antisera raised against mammalian peptides. As described by Holmgren and Jensen in Chapter 2, the non-mammalian peptides differ more or less from their mammalian counterparts, and these dissimilarities may cause difficulties in the interaction between the non-mammalian peptide and the “mammalian” antibody. Therefore, a negative result does not exclude the possibility that the native form of the peptide is actually present. Similarly, as the amino acid sequences of the endogenous peptides of non-mammalian vertebrates in most cases are not known, mammalian peptides are generally used in functional studies, and their interaction with the nonmammalian receptors may not be optimal. Furthermore, few specific antagonists of non-mammalian transmitters have been developed. Despite all these complications, the information gathered in this chapter indicates a widespread occurrence in the vertebrate gut of many of the transmitters reviewed. Similarities in the location and function of the transmitters are plentiful, although many deviations from the general view are reported. The ultimate aim of the studies is, of course, to get a view of the integrated functions of all these different types of nerves in the control of the gut. Bearing in mind the difficulties encountered in non-mammalian studies, it is easy to understand why information from this aspect is still fragmentary. One exception may be the stomach of the rainbow trout, which has been studied to some extent. Unlike most other vertebrates, the stomach of the rainbow trout receives no excitatory vagal input. Distension of the stomach initiates muscular activity of the stomach wall. Again,

FIGURE 4.8 Schematic picture indicating the possible neuronal and hormonal control of the activity of smooth muscle in the cardiac stomach of the rainbow trout, Oncorhynchus mykiss. ACh, acetylcholine; BM, bombesin; GAL, galanin; 5-HT, 5-hydroxytryptamine; SP, substance P; SST, somatostatin; VIP, vasoactive intestinal polypeptide.

158

COMPARATIVE PHYSIOLOGY AND EVOLUTION

unlike mammals, this does not involve vagal reflexes. Enteric cholinergic and serotonergic pathways may mediate this excitation, and may in turn be inhibited by hormonal somatostatin and neuronal VIP, respectively. Substance P-like material is released from neurons and/or endocrine cells of the stomach wall on distension, and may act on the stomach either directly or by release of 5-HT (Figure 4.8, Campbell, 1975; Holmgren, Grove and Nilsson, 1985; Grove and Holmgren, 1992a; Jensen and Holmgren, 1992). Further comparative studies will, hopefully, clarify the evolution of the autonomic nervous control of motility, secretion and blood flow of the gut in different groups of vertebrates.

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166 COMPARATIVE PHYSIOLOGY AND EVOLUTION Tagliafierro, G., Rossi, G.G., Bonini, E., Faraldi, G. and Farina, L. (1989). Ontogeny and differentiation of regulatory peptide and serotonin-immunoreactivity in the gastrointestinal tract of an elasmobranch. J. Expt. Zool., suppl. 2, 165-174. Taylor, G.S. and Bywater, R.A.R. (1989). Novel autonomic neurotransmitters and intestinal function. Pharmacol. Ther., 40, 401-438. Thorndyke, M.C. (1989). Peptides in invertebrates. In The Comparative Physiology o f Regulatory Peptides, edited by S. Holmgren, pp. 202-228. London: Chapman and Hall. Thorndyke, M. and Holmgren, S. (1990). Bombesin potentiates the effect of acetylcholine on isolated strips of fish stomach. Regul. Pepi., 30, 125-135. Thorndyke, M.C., Holmgren, S., Nilsson, S. and Falkmer, S. (1984). Bombesin potentiation of the acetylcholine response in isolated strips of fish stomach. Regul. p ept., 9, 350. Thorndyke, M.C., Riddell, J.H., Thwaites, D.T. and Dimaline, R. (1989). Vasoactive intestinal polypeptide and its relatives - biochemistry, distribution, and functions. Biol. Bull., 177, 183-186. Timmermans, J.P., Scheuermann, D.W., Gabriel, R., Adriaensen, D., Fekete, E. and De GroodtLasseel, M.H.A. (1991). The innervation of the gastrointestinal tract of a chelonian reptile, Pseudemys scripta elegans I. Structure and topography of the enteric nerve plexuses using neuronspecific enolase immunohistochemistry. Histochemistry, 95, 397-402. Timson, C.M., Polak, J.M., Wharton, J., Ghatei, M.A., Bloom, S.R., Usellini, L., et al. (1979). Bombesin-like immunoreactivity in the avian gut and its localization to a distinct cell type. Histochemistry, 61, 213-221. Vaillant, C., Dockray, G.J. and Walsh, J.H. (1979). The avian proventriculus is an abundant source of endocrine cells with bombesin-like immunoreactivity. Histochemistry, 64, 307-314. Vaillant, C., Dimaline, R. and Dockray, G.J. (1980). The distribution and cellular origin of vasointestinal polypeptide in the avian gastrointestinal tract and pancreas. Cell Tissue Res., 211, 511-523. Van Noorden, S. and Falkmer, S. (1980). Gut islet endocrinology - some evolutionary aspects. Invest. Cell Pathol., 3, 21-35. Van Noorden, S. and Patent, G.J. (1980). Vasoactive intestinal polypeptide-like immunoreactivity in nerves of the pancreatic islet of teleost fish, Gillichtys mirabilis. Cell Tissue Res., 212, 139-146. Van Noorden, S., Fritsch, H.A.R., Grillo, T.A.I., Polak, J.M. and Pearse, A.G.E. (1980). Immunocytochemical staining for vertebrate peptides in the nervous system of a gastropod mollusc. Gen. Comp. Endocrinol., 40, 375-376. Vigna, S.R. and Thorndyke, M.C. (1989). Bombesin. In The Comparative Physiology o f Regulatory Peptides, edited by S. Holmgren, pp. 34-60. London: Chapman and Hall. Vizi, S.E., Bertaccini, G., Impicciatore, M. and Knoll, J. (1973). Evidence that acetylcholine released by gastrin polypeptides contributes to their effect on gastrointestinal motility. Gastroenterology, 64, 268-277. von Euler, U.S. and Frange, R. (1961). Catecholamines in nerves and organs of Myxine glutinosa, Squalus acanthias, and Gadus callarias. Gen. Comp. Endocrinol., 1, 191-194. von Euler, U.S. and Ostlund, E. (1957). Effects of certain biologically occurring substances on the isolated intestine of fish. Acta Physiol. Scand., 38, 364-372. Wacyk, J., Guerrero, S. and Morello, A. (1984). Inhibitory effects of acetylcholine on muscular tonus of the small intestine of a lizard. J. Exptl. Zool., 230, 297-301. Wacyk, J., Guerro, S., Morello, A. and V&squez, E. (1989). Some characteristics of the inhibitory mechanism of lizard small intestinal muscular tonus. Comp. Biochem. Physiol., 94C, 441-445. Walsh, J.H. (1987). Gastrointestinal hormones. In Physiology o f the Gastrointestinal tract, 2nd edn, edited by R. Johnson, pp. 181-253. New York: Raven Press. Waterfield, A .A., Smokcum, R.W.J., Hughes, J., Kosterlitz, H.W. and Henderson, G. (1977). In vitro pharmacology of the opioid peptides, enkephalins and endorphins. Eur. J. Pharmacol., 43, 107-116. Watson, A.H.D. (1979). Fluorescent histochemistry of the teleost gut: evidence for the presence of serotonergic neurones. Cell Tissue Res., 197, 155-164. White, T.D. (1991). Role of ATP and adenosine in the autonomic nervous system. In Novel Peripheral Neurotransmitters. Sect. 135 o f Int. Encycl. Pharmacol. Ther., edited by C. Bell, pp. 9-64. New York: Pergamon Press. Williams, J.A. (1982). Cholecystokinin: a hormone and a neurotransmitter. Biomed. Res., 3, 107-121. Yamada, J., Campos, V.J.M., Kitamura, N., Pacheco, A.C., Yamashita, T. and Yanaihara, N. (1987). An immunohistochemical study of the endocrine cells in the gastrointestinal mucosa of the Caiman latirostris. Arch. Histol. Jap., 50, 229-241.

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Yau, W.M. (1978). Effect of substance P on intestinal muscle. Gastroenterology, 74, 228-231. Young, J.Z. (1980a). Nervous control of stomach movements in dogfishes and rays. J. Mar. Biol. Assoc. U.K., 60, 1-17. Young, J.Z. (1980b). Nervous control of gut movements in Lophius. J. Mar. Biol. Assoc. U.K., 60, 19-30. Young, J.Z. (1983). Control of movements of the stomach and spiral intestine of Raja and Scyliorhinus. J. Mar. Biol. Assoc. U.K., 63, 557-574. Young, J.Z. (1988). Sympathetic innervation of the rectum and bladder of the skate and parallel effects of ATP adrenalin. Comp. Biochem. Physiol., 89C, 101-107. Yui, R., Nagata, Y. and Fujita, T. (1988). Immunohistochemical studies on the islet and the gut of the Arctic lamprey, Lampetra japonica. Arch. Histol. Cytol., 51, 109-119. Yui, R., Shimada, M. and Fujita, T. (1990). Immunohistochemical studies of peptide-and aminecontaining endocrine cells and nerves in the gut and the rectal gland of the ratfish Chimaera monstrosa. Cell Tissue Res., 260, 193-201.

5 Glands Ann-Cathrine Jônsson Department o f Zoophysiology, University o f Göteborg, Göteborg, Sweden Gastric acid secretion in non-mammalian vertebrates is generally stimulated by acetylcholine (ACh), bombesin/gastrin releasing peptide (GRP) and gastrin, however, in teleosts gastrin is inhibitory. Catecholamines, vasoactive intestinal polypeptide (VIP) and cholecystokinin (CCK) are inhibitory. Tachykinins and 5-hydroxytryptamine (5-HT) are stimulatory in low doses, while high doses are inhibitory. Pepsin secretion is stimulated by ACh, bombesin/GRP, caerulein, gastrin, 5-HT and tachykinins. Bicarbonate secretion has only been studied in amphibians apart from mammals. Cholinergic agents, catecholamines, CCK, and neurotensin are stimulatory, and somatostatin inhibitory. Mucus secretion has been studied in mammals only. Nerves immunoreactive to a number of putative neurotransmitters are found in the non-mammalian pancreas; of these VIP has been found to stimulate exocrine secretion. Salt secretion is stimulated by VIP in both birds and elasmobranchs. ACh stimulates the bird salt secretion. Bombesin/GRP and somatostatin inhibits the elasmobranch salt secretion. KEY WORDS Autonomic nerves; Pancreas; Salt gland; Gastric acid secretion; Bicarbonate secretion

INTRODUCTION This chapter aims to summarize the present knowledge of the autonomic nervous control of secretory processes in non-mammalian species, focusing on secretions of the gut, pancreas and salt glands. Our knowledge about secretory functions in nonmammalian species is fragmentary, and often based on findings from a few species of birds, amphibians and fish. In mammals, secretory processes are regulated in a complex manner both by inhibitory and stimulatory factors, which may involve hormonal, neuronal and paracrine pathways. The processes in non-mammalian species may be better understood by a comparison with the better-known mammals. Below, the control of secretion of gastric acid and pepsin in mammals will be shortly summarized, and the scattered information from birds, fish and amphibians will be compared to this. Aspects of the secretion of bicarbonate and mucus will be given. The secretion of bicarbonate has frequently been studied in amphibians, 169

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while details on the control of gut mucus formation comes mainly from mammals (see Flemstrôm, 1987; Neutra and Forstner, 1987). The vast majority of studies concerning pancreatic secretion in non-mammals have been performed on fish, and deals with hormonal influences (see Epple and Brinn, 1987; Plisetskaya, 1989), but information on neuronal control is increasing. Salt secreting glands with an osmoregulatory function are present in birds, reptiles and elasmobranchs that live in the marine environment. The morphology and secretory functions of these glands were studied in the late 1950s (cf. SchmidtNielsen, 1960). Later studies have been focused on the presence and function of neuronal regulatory peptides. Recent studies provide increasing evidence for an involvement of regulatory peptides in the control of secretory processes, in addition to the ‘classical’ cholinergic and adrenergic neurons and histamine. The majority of the regulatory peptides are found both in neurons and in endocrine cells or paracrine cells, which often makes a clear distinction between the effects of hormones and neurotransmitters impossible. This chapter will, therefore, occasionally include a discussion of peptide actions that may be of hormonal or paracrine, rather than neuronal, origin. For the sake of simplicity the term IR (immunoreactivity, immunoreactive) will be used where immunohistochemical findings are cited.

REGULATORY PEPTIDES A N D AM INES IN THE GUT M UCOSA, SUBM UCOSA, M USCULARIS M UCOSA A N D LAM INA PROPRIA MAMMALS Several types of peptidergic nerves innervate the mammalian gut mucosa, muscularis mucosa and lamina propria and may influence secretion from the gut wall. Among them are gastrin-releasing peptide (GRP), originally studied as the chemically and biologically similar amphibian peptide bombesin which innervates the secretory glands of the stomach mucosa, and the mucosa of the intestine (Dockray, Vaillant and Walsh, 1979). The muscularis mucosa and lamina propria are innervated by substance P, vasoactive intestinal polypeptide (VIP) and enkephalins both in stomach and intestine. In addition, somatostatin and gastrin/ cholecystokinin (CCK) IR nerves are found in small numbers in the lamina propria and muscularis mucosa (Schultzberg et al., 1980). The presence of catecholaminecontaining neurons is indicated by the presence of IR to the catecholaminesynthesizing enzyme dopamine-jS-hydroxylase (Schultzberg et al., 1980). The mammalian fundic region contains endocrine cells that show IR to histamine (Hakanson et al., 1986), glucagon, 5-hydroxytryptamine (5-HT, serotonin) and somatostatin (see Bordi et al., 1989). The somatostatin cells often have basal processes (Larsson et al., 1979), and it has been shown that anterograde transport of somatostatin can occur in these processes, suggesting a paracrine function (Larsson,

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1984). Endocrine cells IR to gastrin are found in the antrum, together with cells IR to enkephalins, somatostatin and glucagon. The same varieties of endocrine cells are also found in the intestine, with varying distribution. In addition, a number of cells IR to secretin, CCK, gastric inhibitory peptide (GIP), motilin, neurotensin and enteroglucagon are found exclusively or mainly in the intestine. NON-MAMMALIAN SPECIES Myelinated nerve fibres with vesicles presumed to contain peptides are present in the mucosa of the anterior chicken gut (Aisa, Parra and Azanza, 1990). Peptide-like IR, catecholamine-, and serotonin fluorescence have been demonstrated in endocrine cells and in nerves of the mucosa, muscularis mucosa, lamina propria, and submucosa along the whole gastrointestinal canal. Thus, bombesin- and substance P-containing endocrine cells and scattered nerves are found in elasmobranchs, teleosts, amphibians and birds (Holmgren, Vaillant and Dimaline, 1982; Jensen and Holmgren, 1985; Holmgren, 1985; Vigna and Thorndyke, 1989; Rajjo, Vigna and Crim, 1989a; Yui, Shimada and Fujita, 1990; Kiliaan et al., 1992). Likewise, 5-HTfluorescence or 5-HT IR have been demonstrated both in nerves and endocrine cells of teleosts and birds (Watson, 1979; Anderson, 1983; Watanabe, et al., 1987; Kiliaan et al., 1989; Yui, Shimada and Fujita, 1990). Calcitonin gene-related peptide (CGRP) IR nerves are present in the mucosa, often along blood vessels, in teleosts,

FIGURE 5.1 Nerve fibres immunoreactive to galanin (GAL) and vasoactive intestinal polypeptide (VIP) in the intestine and cardiac stomach of cod, Gadus morhua. A) Galanin IR fibres in a villus of the intestine. B) Galanin IR fibres in the glandular area of the cardiac stomach. C) Networks of galanin IR in the cardiac stomach. D) Occasional VIP-IR fibres in the submucosa of the intestine. E) Occasional VIP-IR fibres in the glandular region of the cardiac stomach. A-E x 320.

172 COMPARATIVE PHYSIOLOGY AND EVOLUTION

amphibians, reptiles, and birds (Ohtani et al., 1989). In the ratfish, Chimaera monstrosa, some fibres IR to gastrin were found in the lamina propria (Yui, Shimada and Fujita, 1990). The relatively recently discovered peptide galanin is also present in nerves of the stomach mucosa of cod, Gadus morhua, and rainbow trout, Oncorhynchus my kiss (Jonsson, unpublished data), Figure 5.1. VIP-like IR occurs in elasmobranchs, holosteans, teleosts, amphibians, reptiles, and birds (Reinecke et al., 1981; Holmgren, Vaillant and Dimaline, 1982; Holmgren and Nilsson, 1983a; 1983b; Jensen and Holmgren, 1985; Bjenning and Holmgren, 1988; Rajjo, Vigna and Crim, 1989b: Yui, Shimada and Fujita, 1990; Jonsson unpublished data), Figure 5.1. Catecholamine-fluorescent nerves are found in the submucosa, where the fibres mainly innervate blood vessels (Holmgren, 1985). Some regulatory substances have, so far, been found mainly or exclusively in endocrine cells in the mucosa/submucosa of non-mammalian vertebrates. Amongst these are histamine and somatostatin (El-Salhy et al., 1981; Holmgren, Vaillant and Dimaline, 1982; Yoshida, Iwanaga and Fujita, 1983; H&kanson et al., 1986; see also Holmgren, Jonsson and Holstein, 1986), gastrin (Jonsson, 1989), glucagon (El-Salhy et al., 1981; Holmgren, Vaillant and Dimaline, 1982; Yoshida, Iwanaga and Fujita, 1983; see also Holmgren, Jonsson and Holstein, 1986), GIP- and pancreatic polypeptide (PP) (El-Salhy et al.> 1981; see also Holmgren, Jonsson and Holstein, 1986) and neurotensin (Carraway and Reinecke, 1989). For more information about the overall gut innervation see chapter 4 by Holmgren and Jensen.

GASTRIC ACID SECRETION MAMMALS The mechanisms of secretion of gastric acid and pepsin in mammals has earlier been reviewed thoroughly, and a number of recent reviews can be recommended to the reader: Debase, 1987; Forte and Wolosin, 1987; Soil and Berglindh, 1987; Tache, 1987; Walsh, 1987. Gastric acid is secreted from parietal (oxyntic) cells of the mammalian fundus. Acetylcholine (ACh), gastrin, and histamine are the three major stimulators of acid secretion. Inhibition is caused by CCK, enteroglucagon, galanin, GIP, neurotensin, PP, secretin, tachykinins, VIP, and somatostatin. Bombesin and its mammalian counterpart GRP (which shares the C-terminal sequence with bombesin) are potent stimulators of gastrin release. See also Table 5.1. The secretory events can be divided into four phases: 1. Basal secretion, which occurs when no external stimuli are applied. Basal secretion is under vagal control in some species. 2. The cephalic phase is mediated by ACh release from cholinergic nerves, which not only leads to acid secretion, but also induces gastrin release. It was understood earlier that the cephalic phase is triggered by the sight, smell, and taste of food, and that ‘stress’ is an important acid releasing factor (see Tache, 1987).

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3. The onset of the gastric phase is stimulated by the presence of food in the stomach. Acid secretion is triggered by A) distension of the stomach and B) substances in the food, notably amino acids and digested proteins. 4. The intestinal phase starts when the food enters the intestine. This phase is regulated both by inhibitory and stimulatory components. Amino acids, proteins, and peptides stimulate the release of CCK, and triglycerides and glucose stimulate the release of GIP. Acid in the gut induces the release of somatostatin and secretin. Both hormones may be involved in the acidinduced inhibition. Gastrin release is inhibited by a low pH in the antrum, and as a function of this, further acid secretion is inhibited. Acid in the stomach also releases somatostatin. NON-MAMMALIAN SPECIES Gastric acid release in non-mammalian species has not been examined as extensively as it has in mammals. The information available comes mainly from studies performed in birds and teleosts. Most studies in non-mammalian species have been performed with mammalian and amphibian peptides, often with a pharmacological inclination. Therefore, many of the results cited below should be regarded merely as an indication of how acid secretion in non-mammalian vertebrates may be controlled. Basal Secretion Unless the intestine is simultaneously perfused with diluted (50% or 33%) sea water, Atlantic cod (Gadus morhua), become dehydrated and produce only small amounts of gastric acid during experimental conditions (gastric perfusion) (Holstein, 1979). In intestinally perfused fish the basal secretion of acid increased, and shamoperated, intestinally perfused cod had a “basal” acid secretion around 100150pimolkg-1h -1, which remained stable for a week (Holstein and Cederberg, 1980; 1984). The gastric acid output declined by 96% after vagotomy (Holstein and Cederberg, 1980), which shows that basal acid secretion in cod is under vagal control. Gastric acid output also declined after treatment with atropine and histamine antagonists. In chicken, a mean rate of basal gastric acid secretion of 3-5.6 ml h -1 was measured; this equals an acid output of 1.46-3.32 m Eqh-1 (Angelucci and Linari, 1970; Burhol and Hirschowitz, 1970; Sewing and Ruoff, 1972). It has been concluded that acid secretion is under vagal control also in chicken (Burhol, 1973b). Stimulation o f Gastric Acid Secretion A cephalic phase has been documented in birds. Thus, “trained” ducks secreted acid in response to the sound of a bell. The sight of food also stimulated acid secretion, whereas food odours did not (Smit, 1968). Acid secretion in teleosts and frogs is stimulated by stomach distension (Smit, 1968); this probably corresponds to the gastric phase of mammals.

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Gastric acid secretion in elasmobranchs, teleosts, amphibians, reptiles, and birds is stimulated by histamine, ACh and carbachol (Davidson, Lemmi and Thompson, 1966; Kasbekar, 1967; Smit, 1968; Burhol and Hirschowitz, 1970; Sewing and Ruoff, 1972; Holstein, 1975; 1976; 1977). Atropine prevented the increase in acid secretion induced by ACh and carbachol in elasmobranchs and cod (see Smit, 1968; Holstein, 1977) and metiamide, a histamine H2-receptor antagonist, completely inhibited both histamine- and carbachol-stimulated acid secretion in cod (Holstein, 1976). It is thus possible that cholinergic nerves stimulate the release of histamine which in turn triggers the release of acid. This also occurs in bullfrogs; ACh was found to release histamine, which then stimulated acid secretion (Ruiz and Michelangeli, 1984). 5-HT stimulated gastric acid and pepsin secretion in intestinally perfused cod (Holstein and Cederberg, 1984). However, high doses of 5-HT were inhibitory. The 5-HT stimulation may well be caused by a neuronal pathway in addition to a hormonal pathway, as 5-HT IR is contained both in nerve fibres and endocrine cells of the stomach mucosa. In the same type of preparation, low doses of the tachykinins physalaemin and eledoisin stimulated acid release, whereas higher doses were inhibitory (Holstein and Cederberg, 1986). Moderate and high doses of substance P and eledoisin-related peptides tended to stimulate the acid secretion, and the tachykinins were also shown to be effective at stimulating pepsinogen secretion in the cod (Holstein and Cederberg, 1986). Bombesin enhances acid secretion in some non-mammalian species; in chicken, the stimulation by bombesin occurs indirectly via release of gastrin (Linari and Baldieri-Linari, 1976), while in the amphibian, Rana catesbeiana, the stimulatory effect may be direct (Ayalon et al. , 1981). In cod, bombesin also stimulates acid secretion, possibly by an inhibition of the release of VIP (Holstein and Humphrey, 1980). Exogenous porcine VIP is inhibitory on cod acid secretion (see below). The presence of bombesin, a substance P-like tachykinin, and VIP in mucosal nerve fibres, and in endocrine cells, points to a situation with both hormonal and neuronal influences. Gastrins (or related CCK-variants and caerulein), are stimulatory on gastric acid secretion in birds, amphibians and elasmobranch fish, but probably not in teleost fish (see Jonsson and Holmgren, 1989). The gastrin-related peptides have been found in endocrine cells in the mucosa and a recent study demonstrated gastrin-like IR nerves in the lamina propria of ratfish (Yui, Shimada and Fujita, 1990). The possibility therefore exists that gastrin is contained in mucosal nerves of at least some species. However, the release of gastrin from endocrine cells is most likely under the influence of autonomic nerves (see above). Inhibition o f Gastric Acid Secretion Most of the known inhibitors of gastric acid secretion in non-mammalian species are hormones. Thus, insulin dose-dependently inhibited gastric acid secretion in fistula chickens, but did not inhibit pepsin secretion (Burhol, 1973b). In further studies to explain the lack of stimulatory activity of pentagastrin in cod (Holstein,

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FIGURE 5.2 Summarizing figure showing neurotransmitters known to innervate the mucosa and to influence the acid secretion (adapted from Jonsson and Holmgren, 1989). ACh, acetylcholine; BM/GRP, bombesin/gastrin releasing peptide; G/CCK, gastrin/cholecystokinin; H + , gastric acid; Hi, histamine; 5-HT, 5-hydroxytryptamine (serotonin); SP, substance P; VIP, vasoactive intestinal polypeptide. + indicates an increased gastric acid secretion, — indicates a decreased gastric acid secretion, and + / indicates that the effect can be either increased or decreased gastric acid secretion based on results obtained with low or high doses.

1975), it was found that gastrin, the octapeptide form of cholecystokinin (CCK-8), and caerulein all inhibited basal acid secretion (Holstein, 1982). Somatostatin may function via a paracrine action in cod as in mammals, since somatostatin IR endocrine cells of the cod mucosa have varicose basal processes (Jonsson, unpublished). As in mammals, somatostatin inhibits histamine-stimulated acid secretion (see Holmgren, Jonsson and Holstein, 1986). 5-HT and gastric distension inhibited basal-, histamine-, and pentagastrinstimulated acid secretion in chicken (Joyner and Kokas, 1971), an inhibition that may involve enteric neurons and autonomic reflexes as in mammals. In fish, high doses of 5-HT were inhibitory, while lower doses stimulated gastric acid secretion (Holstein and Cederberg, 1984). TABLE 5.1 Summary of “classical” neurotransmitters and putative peptide neurotransmitters that influence gastric acid secretion. + indicates stimulation, - inhibition, and 0 no effect on gastric acid secretion. ACh, acetylcholine and cholinergic agents; CA, catecholamines; GRP, gastrin releasing peptide; CCK, cholecystokinin; VIP, vasoactive intestinal polypeptide. Substance

Mammals

Birds

Reptiles

Amphibians

Teleosts

Elasmobranchs

ACh Bombesin/GRP CA CCK Galanin Gastrin Neurotensin 5-HT(serotonin) Somatostatin Tachykinins VIP

+ +

+ +

+

+ +

+ +

+

+

+

—

+

+

-

+

-

+

0

/ -

-

—

+/—

+/—

+

176 COMPARATIVE PHYSIOLOGY AND EVOLUTION

Administration of the neurotransmitter VIP inhibited acid secretion in cod (Holstein and Humphrey, 1980; Holstein, 1983) and adrenaline inhibits secretion in elasmobranchs (Smit, 1968). For a summary see Figure 5.2 and Table 5.1.

PEPSIN /PEPSIN O G EN SECRETION MAMMALS Pepsin is formed from the inactive precursor pepsinogen after secretion into the lumen of the stomach. The autocatalytical conversion occurs after pepsinogen has entered the stomach lumen as a result of the acidic nature of the stomach contents. Pepsin has a pH optimum around 2 (see Smit, 1968) which, incidently, is the pH in the stomach after ingestion of food in most vertebrates. Pepsinogen is stored in the gastric chief (peptic) cells in mammals. Regulation Pepsinogen secretion in mammals is stimulated by feeding and by the regulatory peptides GRP, bombesin, CCK, secretin, and substance P. These peptides may be released from endocrine cells or from nerve terminals (Hersey, 1987). Muscarinic agents stimulate pepsinogen secretion, whereas adrenergic agents have been suggested to be inhibitory (Hersey, 1987; Bech, 1989). Somatostatin counteracts the stimulatory effect of many substances (Hersey, 1987). Glucagon and high doses of histamine also appear to be inhibitory (Bech, 1989). See also Table 5.2. NON-MAMMALIAN SPECIES In non-mammalian species pepsinogen and acid are stored in and released from the same cells, the oxynticopeptic cells (Smit, 1968; Helander, 1981). In addition, some amphibians, such as frogs and Necturus, have oesophageal pepsinogen-storing cells that lack acid-production (Friedman, 1937; Simpson, Goldenberg and Hirschowitz, TABLE 5.2 Summary of “classical” neurotransmitters and putative peptide neurotransmitters that influence the pepsin secretion. -I- indicates stimulation, — inhibition, and 0 no effect on pepsin secretion. ACh, acetylcholine and cholinergic agents; CA, Catecholamines; GRP, gastrin releasing peptide; CCK, cholecystokinin. Substance

Mammals

Birds

Amphibians

Teleosts

ACh Bombesin/GRP CA CCK/caerulein Gastrin 5-HT (Serotonin) Somatostatin Tachykinins

+ +

+

+ + 0

+

—

0/ + -

-1 -

0/ + +

0

0 +

-

+

GLANDS

177

1980). Antibodies raised against chicken pepsinogen have revealed pepsinogenstoring cells in ascidians, elasmobranchs, teleosts (with stomachs), amphibians, lizards, and birds (Yasugi, 1987; Yasugi, Matsunaga and Mizuno, 1988; 1989). Pepsinogen IR was not found in teleosts lacking a stomach (Yasugi, 1987; Yasugi, Matsunaga and Mizuno, 1988). The structure of pepsinogen appears to have changed little during the course of evolution (Yasugi, 1987). Regulation Although both acid and pepsinogen are formed in the same cells, they seem, to a certain extent, to be controlled by different stimuli. In chicken, pentagastrin stimulated the release of both acid and pepsin, whereas CCK was inactive (Burhol, 1974). However, the pepsin to acid ratio increased after pentagastrin treatment, indicating that pepsin release was stimulated more strongly than acid release (Burhol, 1973a). Caerulein also stimulated pepsin outflow (Angelucci and Linari, 1970). The cholinergic drug bethanechol stimulated pepsinogen secretion from cultured “pure” peptic cells from the oesophagus of bullfrogs, indicating a cholinergic control. Pentagastrin, histamine, isoprenaline (a /3-adrenoceptor agonist) and noradrenaline had no effect (Simpson, Goldenberg and Hirschowitz, 1980). Of the total pepsinogen-content of the stomach 1.33% was secreted during a 150min period (Simpson, Goldenberg and Hirschowitz, 1980). In frogs, secretion from “pure” peptic cells was induced by bombesin (Shirakawa and Hirschowitz, 1984). The immediate effect of bombesin was reported to be relatively independent of Ca2+, whereas the sustained effect was Ca2+ dependent (Hirschowitz et al., 1990.). The pepsin secretion in response to bethanechol from the same “pure” peptic cells was inhibited by somatostatin via a cAMP-independent pathway (Fong, Hong and Wang, 1991). Somatostatin did however not influence the basal pepsin secretion (Fong, Hong and Wang, 1991). Tachykinins effectively stimulated pepsin secretion from gastrically and intestinally perfused cod, whereas their effects on acid secretion were less pronounced (Holstein and Cederberg, 1986). In cod, both acid and pepsinogen secretion, at basal levels and at elevated levels resulting from histamine or carbachol treatment, were stimulated by 5-HT. When administered alone, histamine or carbachol strongly stimulated acid secretion, but only stimulated pepsinogen secretion weakly (Holstein and Cederberg, 1984). For a summary see Table 5.2.

GASTROINTESTINAL BICARBONATE/ALKALI SECRETION Bicarbonate, (H C03~), is secreted from the stomach and intestinal mucosa and from the pancreas. The proximal parts of the intestine secrete more H C 03” than the distal parts. H C 03_ neutralizes the gastric acid secreted from the stomach and has been ascribed a protective role. This protection is associated with the gut mucus and is called the “mucus-bicarbonate barrier”. A low pH in the gut initiates H C 03_ secretion.

178

COMPARATIVE PHYSIOLOGY AND EVOLUTION

MAMMALS Secretion is regulated by neuronal, humoral, and paracrine factors. Intravenous injection of VIP and carbachol stimulated H C 03" output (Lenz, Vale and Rivier, 1989). Intestinal HCOf secretion is influenced by some of the peptides known to occur in the brain. Thus, thyrotropin-releasing hormone (TRH), GRP, bombesin, and corticotropin-releasing factor (CRF) enhance the H C 03" secretion from the rat intestine (Flemstrom and Jedstedt, 1989; Lenz, Vale and Rivier, 1989). The increase after treatment with bombesin or TRH was prevented by vagotomy, suggesting the involvement of a vagal pathway (Flemstrom and Jedstedt, 1989; Lenz, Vale and Rivier, 1989). It has been suggested that the effect of TRH administered centrally results from an increase in vagal outflow, mediated by release of VIP and the activation of a muscarinic pathway (Lenz, Vale and Rivier, 1989). ‘Classical’ neurotransmitters (ACh and catecholamines), as well as opioid peptides, CCK, PP and neurotensin stimulate H C 03~secretion (Flemstrom, 1987). See also Table 5.3. NON-MAMMALIAN SPECIES Ca2+ increased the rate of H C 03_ secretion in the amphibians Necturus and Rana catesbeiana. This effect was abolished by atropine treatment, suggesting that ACh was released from intramucosal nerves (Flemstrom and Garner, 1980). The amphibian proximal duodenum has been used extensively to examine the function of H C 03" release. In vitro preparations have shown that alkali is secreted at a rate of about 1 ¿teqh"1cm-2. HCOf secretion increased after treating the stomach and duodenum with acid (Flemstrom, Heylings and Garner, 1982). Noradrenaline, pancreatic glucagon, neurotensin and GIP all stimulated the secretion of duodenal h c o 3 The fundic and antral parts of the amphibian stomach secrete alkali at a rate of about 0.1-0.2 and 0.35^eqcm-2 h -1, respectively (Flemstrom and Sachs, 1975; TABLE 5.3 Summary of “classical” neurotransmitters and putative peptide neurotransmitters that influence bicarbonate secretion. + indicates stimulation, and 0 no effect on bicarbonate secretion. ACh, acetylcholine and cholinergic agents; CA, catecholamines; GRP, gastrin releasing peptide; CCK, cholecystokinin; VIP, vasoactive intestinal polypeptide. Substance

Mammals

Amphibians

ACh Bombesin/GRP CA CCK Enkephalin/opioids Gastrin Neurotensin 5-HT (serotonin) Somatostatin Tachykinins VIP

+

+

+ + +

+

4-

+

4-

0

4 -/0 0 40 0 0 0

GLANDS

179

Takeuchi et a i , 1983). The gastric secretion of H C 03_ increased after treatment with glucagon or CCK, but these peptides do not influence the intestinal secretion of bicarbonate (Flemström, Heylings and Garner, 1982). Histamine, pentagastrin, tetragastrin, urogastrone, ACTH, bombesin, motilin, secretin, 5-HT, somatostatin, substance P, and VIP had effect on neither gastric nor duodenal H C 03_ secretion (Flemström, Heylings and Garner, 1982). For a summary see Table 5.3.

GASTROINTESTINAL M UCUS Mucus is a complex gel-like secretory product of the epithelial layer of the gastrointestinal canal. It consists mainly of water, electrolytes, immunoglobins, enzymes, and the glycoprotein mucin. Mucin is produced in mucus cells in the salivary glands, oesophagus, intestine, gallbladder, pancreas, and in the cardiac gland area of the stomach. The type of mucin produced varies depending on where it is formed, and on the diet (Neutra and Forstner, 1987). Mucin secretion is stimulated by acetylcholine/cholinergic nerves (Seidler and Sewing, 1989), and CCK is a putative stimulator (Neutra and Forster, 1987). Histamine and pentagastrin had no effect in rabbit (Seidler and Sewing, 1989). The gastric “mucus-bicarbonate barrier” is regarded as providing the gastrointestinal mucosa with effective protection against the hazardous influence of acid and pepsin in the gut. In Rana catesbeiana the luminal pH was 3-3.5, while the pH of the junction mucus-mucosal cells was as high as 6.5 (Takeuchi et a l , 1983). However, the validity of these findings has recently been questioned. The mucosa was not injured even after rat gut with a disrupted “barrier” had been exposed to acid (Wallace, 1989). Nothing is known about the control of mucus secretion in non-mammalian vertebrates.

PANCREAS The pancreatic function ranges in complexity from single endocrine and exocrine cells in the intestinal mucosa of cyclostomes and Amphioxus, to the compact organ found in tetrapods, where the endocrine cells are organized into clusters - the islets of Langerhans - intermingled with the exocrine cells. A compact pancreas of the tetrapod type occurs for the first time in elasmobranchs. In teleosts, the endocrine tissue is arranged as a mass of endocrine cells, often surrounded by a rim of exocrine tissue. These, mainly endocrine, structures are called Brockmann bodies. The endocrine and exocrine parts are considered to have developed independently of each other (Epple and Brinn, 1987). The exocrine pancreas secretes a protein-and bicarbonate-rich fluid. The protein secretions help digest the food and include proteolytic, amylytic, lipolytic, and nucleolytic enzymes. The bicarbonate secretion helps to neutralize the acidic food entering the intestine. The composition of this fluid varies, depending on the stimuli. The endocrine pancreas secretes the metabolic hormones somatostatin, insulin, glucagon, and PP.

180 COMPARATIVE PHYSIOLOGY AND EVOLUTION

THE CONTROL OF PANCREATIC FUNCTION Factors controlling the mammalian pancreas have been studied extensively since the discovery by Bayliss and Starling, 1902 that secretin, released from duodenum, stimulates pancreatic H C 03" secretion. However, information about the nonmammalian pancreas is scarce, with the exception of the endocrine part of the teleost pancreas, the Brockmann bodies. The endocrine and exocrine secretion processes are influenced by both hormonal and neuronal inputs. The pancreas is supplied by nerve fibres via pancreatic nerves, often running along the pancreatic arteries. The nerves can be classified as parasympathetic fibres from the vagal trunks, sympathetic fibres from the splanchnic nerves and visceral afferents (Woods and Porte, 1974). A number of different nerve terminals have been identified ultrastructurally by their type of vesicles (granules). Nerve fibers ultrastructurally classified as adrenergic (mainly small, granular vesicles) and cholinergic (mainly small, agranular vesicles) have been reported in the pancreas of most non-mammalian vertebrates, with the possible exception of cyclostomes and some birds. These nerve fibers may also contain other types of vesicles, often large and dense-cored (Woods and Porte, 1974; Epple and Brinn, 1975). Nerves containing large dense-cored vesicles are considered to contain neuropeptides. The nerve fibres, often located close to endocrine cells and blood capillaries, show ultrastructural signs of exocytosis (Golding and Pow, 1990; Endo, Chiba and Honma, 1991). Glucagon, insulin, somatostatin, PP, neurotensin and the molluscan cardioexcitatory neuropeptide Phenylalanine-Methionine-ArgininePhenylalanine-amide (FMRF) are so far only reported in endocrine cells of the pancreas, and will not be further discussed here. Bombesin-related peptides Bombesin and GRP-like IR has been found in occasional nerves in the pancreas of elasmobranchs, holocephalans, and teleosts (Yui and Fujita, 1986; Jonsson, 1991), Figure 5.3. The fibres innervate the exocrine tissue and to a lesser extent the walls of the blood vessels. In mammals, bombesin stimulates the output of proteins from the pancreas (Herzig, Louie and Owyang, 1988; Varga et al., 1988). Bombesin and GRP also stimulate the overall output of pancreatic juice, trypsin, and amylase, and act as trophic factors (Varga et al., 1988). The innervation of the exocrine tissue and blood vessels of the elasmobranch and holocephalan pancreas indicates that bombesin similarly may exert its effects on the exocrine system of these fish species. Bombesin may also control the blood supply to the endocrine and exocrine cells. Of the teleosts studied, Salmo species lacked bombesin IR nerves in the Brockmann bodies, whereas occasional bombesin IR nerves have been found in the walls of the blood vessels in the Brockmann bodies of cod (Jonsson, 1991). Like in other vertebrates, bombesin may function mainly as an exocrine regulator in teleosts, which would explain the lack of innervation of the Brockmann bodies. Bombesin/GRP also occur in some primary afferent neurones (Holzer, 1988), and it is possible that the bombesin/GRP IR nerves in the fish pancreas have a sensory function.

GLANDS

181

FIGURE 5.3 Nerve fibres immunoreactive to bombesin (BM), galanin (Gal), and vasoactive intestinal polypeptide (VIP) in the pancreas of representatives of elasmobranchs and teleosts. A) A bombesin IR fibre in the spiny dogfish, Squalus acanthias. B) Galanin IR networks in the Brockmann bodies of rainbow trout, Oncorhynchus my kiss. C) Galanin IR networks of fibres in the Brockmann bodies of cod, Gadus morhua. D) Galanin IR fibre in Spiny dogfish, Squalus acanthias. E) VIP-IR fibres in rainbow trout, Salmo gairdneri. A-B and D-E x 320, C x 130.

CGRP CGRP-containing nerves have been demonstrated in the connective tissue of the lamprey, where they were sometimes closely associated with blood vessels and islet cells (Yui, Nagata and Fujita, 1988). CGRP is a comparatively recently discovered neuropeptide, and little is known of its distribution and effect on the pancreas. It has been recently reported that CGRP inhibits CCK-stimulated amylase secretion via an indirect neuronal pathway in rats (Bunnett, Mulvihill and Debas, 1991). CCK CCK-like IR has been detected in nerve fibres of the mammalian pancreas (Larsson, 1979; Larsson and Rehfeld, 1979). Gastrin IR is present in the mammalian pancreas

182 COMPARATIVE PHYSIOLOGY AND EVOLUTION

from early ontogeny, but only in endocrine cells (see Jonsson, 1989). Antisera against the common C-terminal portion of gastrin/CCK reveal scarce nerve fibres in Brockmann bodies of goldfish, and in endocrine cells of Squalus and Raja (Jonsson, 1991). CCK-like IR was also found in pancreatic endocrine cells of elasmobranchs (Tagliafierro, Faraldi and Pestarino, 1985). It is possible that the teleost pancreas is innervated by gastrin/CCK-like containing nerves, whilst the pancreas of ,the more “primitive" elasmobranchs is under a paracrine or hormonal gastrin/CCK-like control (Jonsson, 1991). CCK is a putative sensory transmitter (see Holzer, 1988), and it is conceivable that the gastrin/CCK/caerulein-IR nerves have a sensory function in the fish pancreas. CCK has been reported to stimulate the release of the pancreatic hormones insulin, glucagon, and somatostatin in mammals (see Miller, 1981). Moreover, the peptides of the gastrin/CCK family stimulate pancreatic growth (See Walsh, 1987). These peptides might also have the same functions in fish. Enkephalins Met-enkephalin containing nerve fibres were reported to innervate the pancreas of cats (Larsson and Rehfeld, 1979) and humans (Feurle, Helmstaedter and Weber, 1982). Opioid peptides, in general, have also been found in endocrine cells of the mammalian pancreas, although these findings have been the subject of some controversy (Cetin, 1990). A small number of enkephalin-containing endocrine cells are also present in some holocephalans (Epple and Brinn, 1987). Occasional metenkephalin-Arginine 6-Phenylalanine7 (MERF) nerve fibres were found in cod pancreas, whilst weak IR was regularly detected in endocrine cells of Salmo species and cod (Jonsson, 1991). In mammals, met-enkephalin has been shown to inhibit the release of somatostatin ancl stimulate the secretion of insulin (Hermansen, 1983). In humans metenkephalin stimulates the exocrine secretion (Gullo et al., 1986). The effects of enkephalins on the non-mammalian pancreas are not known. Galanin Neurons containing galanin innervate exocrine and endocrine tissue in mammalian pancreas. Extensive networks of galanin IR nerve fibres are present throughout the Brockmann bodies in rainbow trout, goldfish Carassius auratus, and cod, and in pancreatic tissue of Raja radiata and Squalus acanthias (Jonsson, 1991), Figure 5.3. Galanin inhibits amylase secretion from isolated acini in mammals and influences the secretion of glucagon, insulin, and somatostatin (see Plisetskaya, 1989). The dense innervation of the Brockmann bodies of teleosts (Jonsson, 1991) suggests that galanin is involved in the control of the secretion of all hormones in the islets of these animals. In elasmobranchs, galanin may in addition influence exocrine functions, such as amylase secretion. Galanin has been reported to occur in some sensory nerves in mammals (see Holzer, 1988), and a sensory function in the fish pancreas can not be ruled out.

GLANDS

183

Neuropeptide Y (NPY), oxytocin and substance P Nerves immunoreactive to NPY have been found in the rat pancreas, and the peptide inhibits glucose-stimulated insulin release (see Plisetskaya, 1989). In the anglerfish, Lophius americanus, NPY IR fibres were demonstrated in the islets (Noe et al., 1986), but there are so far no indications of their function. Nerves containing oxytocin-like peptides surround insulin-containing cells in the pancreas of anglerfish, and an influence on the release of insulin has been suggested (McDonald et al., 1987). Nerve fibres immunoreactive to substance P have been found in teleost pancreatic tissue (Van Noorden and Patent, 1980), but there are so far no indications of their function. VIP Nerves IR to VIP have been found in the pancreas of teleosts, elasmobranchs, holocephalans, reptiles, and birds (Van Noorden and Patent, 1980; Yui and Fujita, 1986; Masini 1988; Hiramatsu and Watanabe, 1989; Jonsson, 1991), Figure 5.3. The nerves were observed in the connective tissue, surrounding the islets and innervating blood vessels. Nerve fibres were also found in the exocrine tissue, often surrounding acinar cells and ducts. Neuronal somata labelled with VIP antibodies have been demonstrated in ratfish (Yui and Fujita, 1986), and intrapancreatic ganglion cells labelled with VIP antibodies have been reported from birds (Hiramatsu and Watanabe, 1989). VIP stimulates exocrine secretion in mammals (Lundberg, 1981). In the turkey, porcine and chicken VIP both stimulate exocrine secretion, chicken VIP being the most potent (Dimaline and Dockray 1979). PHI (peptide with N-terminal histidine and C-terminal isoleucine), a peptide originating from the same precursor as VIP, stimulates the turkey pancreas (Dimaline and Dockray, 1980). Fish VIP was shown to stimulate amylase secretion in guinea pigs (see Dimaline, 1989), and dogfish VIP stimulates exocrine secretion in turkey (Dimaline and Thorndyke, 1986). It is very likely that the VIP-like material in fish pancreatic nerves regulates the secretion from the exocrine tissue. VIP is known to increase blood flow and relax vascular smooth muscle in mammals (see Dimaline, 1989), a function that it may also have in the fish pancreas, since the blood vessels are innervated by VIP neurons. Finally, based on the finding that VIP surrounds the islets it is possible that VIP may also control the release of islet hormones, either directly or via changes in the blood flow. Adrenergic nerves The presence of adrenergic nerves in the fish pancreas have been demonstrated lately with antibodies against the catecholamine synthesizing enzyme dopamine-j3hydroxylase (DBH) (Jonsson, 1991; Milgram, McDonald and Noe, 1991) in the Brockmann bodies of cod and anglerfish (Lophius americanus). Of the other two enzymes in the synthesizing chain, phenyl-N-methyl transferase (PNMT) and tyrosine hydroxylase (TH), TH was found in anglerfish (Milgram, McDonald and

184 COMPARATIVE PHYSIOLOGY AND EVOLUTION

Noe, 1991) but not in cod (Jonsson, 1991). PNMT could not be demonstrated (Jonsson, 1991). In mammals, adrenergic agents inhibit the release from the pancreas and induce vasoconstriction (Williams, 1975; Varga, Papp and Vizi, 1990). Noradrenaline and adrenaline inhibit the release of insulin, glucagon, and somatostatin (Miller, 1981). These effects are probably due to a-adrenergic activation, as ^-adrenergic activation appears to stimulate both insulin and glucagon secretion (Woods and Porte, 1974). In the cat adrenergic nerves innervate blood vessels, islets, and ganglion cells (Larsson and Rehfeld, 1979, Larsson, 1979). Occasional fibres have also been found in exocrine tissue. The DBH-IR nerve fibres in the fish pancreas were associated with glucagon and insulin-containing cells, and in the presence of glucose, noradrenaline and isoproterenol (isoprenaline) stimulated glucagon and somatostatin-14-release. Isoproterenol stimulated insulin release. Noradrenaline in low concentrations stimulated, while high concentrations inhibited the insulin secretion (Milgram, McDonald and Nde, 1991). The presence of DBH-IR in perivascular nerves points to a role of catecholamines in the control of the pancreatic blood flow in fish as in mammals. Cholinergic Nerves The acetylcholine synthesizing enzyme choline-acetyltransferase (ChAT) has been demonstrated in the Brockmann body of the anglerfish, indicating the presence of acetylcholine-containing neurones in the fish islet tissue (Milgram, McDonald and Noe, 1991). In mammals, cholinergic agents increase the insulin secretion, whereas the effects on glucagon secretion are mixed (Woods and Porte, 1974). ACh has been shown to induce amylase secretion in mammals (Williams, 1975; Varga, Papp and Vizi, 1990), and PP secretion from the endocrine cells is probably triggered via vagal cholinergic pathways (see Plisetskaya, 1989). Metacholine, a muscarinic agonist stimulated the release of insulin, glucagon and somatostatin-14 from the anglerfish Brockmann body (Milgram, McDonald and Noe, 1991). Ach and VIP are sometimes stored in the same nerves in salivary gland and other exocrine tissues of mammals (Lundberg, 1981), and VIP was shown to potentiate the effect of acetylcholine in cat salivary glands (Lundberg, Angg&rd and Fahrenkrug, 1982). Whether this is the case also in the pancreas is not known.

SALT SECRETING GLANDS Salt secreting glands occur in marine elasmobranchs, reptiles (snakes, lizards, turtles), and birds. These organs, which have an osmoregulatory function, secrete mainly Na+ and Cl", and remain inactive until stimulated by a salt load. Marine birds and turtles have a pair of salt secreting glands in or close to the orbit of the eye (supraorbital glands) (Schmidt-Nielsen and Fange, 1958). Marine lizards have salt glands (nasal glands) that empty their fluids into the nasal cavity, with a ridge

GLANDS

185

that keeps the salty fluid from flowing backwards and being swallowed (SchmidtNielsen and Fange, 1958). In some marine snakes, salivary glands serve in ion secretion. The estuarine crocodile Crocodylus porosus has both nasal glands and supraorbital glands (Schmidt-Nielsen and Fange, 1958). In elasmobranchs a single salt gland, the rectal gland, is situated near the cloaca. Physiological studies of salt glands have been focused on elasmobranchs and birds, and the following information is based on data from these two groups. ELASMOBRANCHS The rectal gland is a cylindrical organ with a central canal that continues as a duct until it empties into the hindgut, posterior to the spiral valves. The blood flow is parallel to that of the secretory fluid. The apical region of the glandular cells, which face the duct, is covered with microvilli (Bulger, 1963). The rectal glands in Squalus acanthias secrete a Na+ and Cl- rich solution with a salt concentration about twice that of blood plasma (Burger and Hess, 1960). VIP-, GRP-, and bombesin-IR nerve fibres form dense plexuses that innervate the capsule surrounding the glandular tissue, and run along the secretory tubules and in the wall of the central excretory duct (Holmgren and Nilsson, 1983a; Chipkin, Stoff and Aronin, 1988; Yui, Shimada and Fujita, 1990). A variety of effects of VIP on the salt-gland secretion have been reported. In Squalus acanthias, porcine VIP stimulated the Cl" secretion via a cAMP-dependent mechanism (Stoff et aL, 1979). However, neither porcine VIP, nor a partially purified Scyliorhinus VIP, had any effect on the secretion of salt from Scyliorhinus canicula or Raja radiata (Shuttleworth and Thorndyke, 1984). Instead, a peptide factor named ‘rectin’, isolated from Scyliorhinus intestine strongly stimulated the rectal gland in Squalus, Scyliorhinus, and Raja (Shuttleworth and Thorndyke, 1984; Thorndyke and Shuttleworth, 1985). Bombesin and somatostatin both inhibit salt secretion (Stoff et al., 1979; Silva, Lear and Epstein, 1988; Silva et al., 1985; 1990). A small number of somatostatinlike fibres innervate the capsule and the tissue surrounding the excretory duct, and a few gastrin/CCK-like fibres have also been demonstrated in the capsule (Holmgren and Nilsson, 1983a). See also Figure 5.4. BIRDS The salt glands in birds are flat and crescent-shaped. The glands consist of two parts, and two ducts run from each side to the beak. The secretory fluid runs through the nares and drips off from the tip of the beak. The glands consists of longitudinal lobes arranged to surround a central canal. The salt glands of birds are highly vascularized with the arterioles forming a counter-current flow system with the secretory fluid (Schmidt-Nielsen, 1960). Nerve fibres containing ACh and VIP have been demonstrated in avian salt glands. The presence of VIP neurons around the secretory tubules and surrounding arterioles has been revealed by immunohistochemistry (Lowy, Schreiber and Ernst,

Glandular cells

BM/GRP -

G/CCK ?

Birds

Capsule

Glandular cells

ACh +

VIP +

FIGURE 5.4 Summarizing figure showing the innervation of the salt glands, and the effect of these neurotransmitters. ACh, acetylcholine; BM/GRP, bombesin/gastrin releasing peptide; G/CCK, gastrin/cholecystokinin; SST, somatostatin; VIP, vasoactive intestinal polypeptide. + indicates an increased salt secretion, — indicates a decreased salt secretion, and ? indicates that the function of these nerves are not known.

Capsule

Elasmobranchs

186 COMPARATIVE PHYSIOLOGY AND EVOLUTION

GLANDS

187

1987; Gerstberger, 1988), and ACh and VIP stimulate both secretion and flow (Gerstberger, Sann and Simon, 1988). Co-infusion of ACh and VIP results in an additional effect on osmolal excretion, while the effect on the capillary blood flow through the glands is non-additive (Gerstberger, Sann and Simon, 1988). Binding sites for VIP have been found at the apical end of the secretory tubules (Gerstberger, 1988). For a summary see Figure 5.4.

SUMMARY The non-mammalian gut mucosa, submucosa, muscularis mucosa and lamina propria are innervated by bombesin, CGRP, substance P, 5-HT, galanin, gastrin/CCK, VIP and catecholamine containing nerves. In non-mammalian species gastric acid and pepsin are secreted from the same cells, the oxynticopeptic* cells, but their release seems to be controlled by partly different mechanisms. The effect of gastrin on pepsin release was stronger than the effect on acid secretion in chickens, and tachykinins also stimulated pepsin release stronger than the acid release in cod. The acid secretion in birds and teleosts are stimulated by the neurotransmitters bombesin, ACh, 5-HT and substance P. Pepsin secretion is stimulated by the neurotransmitters bombesin, cholinergic agents, 5-HT, and tachykinins. The control of alkali secretion from the intestine is different from that of the stomach; thus CCK and glucagon stimulate gastric secretion but not intestinal secretion. The pancreas in non-mammalian species is innervated by bombesin-, CGRP-, DBH-, enkephalin-, galanin-, gastrin/CCK-, substance P-, and VIP-IR nerves. Of these substances, only the effect of VIP has been examined in any of the nonmammalian species, and it has been shown that VIP stimulates the exocrine secretion in birds. Salt secretion from the salt glands in birds and elasmobranchs are under the control of cholinergic and VIP-releasing nerves; both neurotransmitters stimulate salt secretion. Fibres with somatostatin and bombesin IR are also found; these peptides inhibit the salt secretion.

REFERENCES Aisa, J., Parra, P. and Azanza, M.J. (1990). Ultrastructural characteristics of anterior gut innervation of Gallus Galius. Histol. Histopathol., 5, 281-287. Anderson, C. (1983). Evidence for 5-HT-containing intrinsic neurons in the teleost intestine. Cell Tissue Res., 230, 377-386. Angelucci, L. and Linari, G. (1970). The action of caerulein on gastric secretion of the chicken. Eur. J. Pharmacol., 11, 204-216. Ayalon, A., Yazigi, R., Devitt, P.G., Rayford, P.L. and Thompson, J.C. (1981). Direct effect of bombesin on isolated gastric mucosa. Biochem. Biophys. Res. Commun., 99, 1390-1397. Bayliss, W.M. and Starling, E.H. (1902). The mechanism of pancreatic secretion. J. Physiol. (Lond), 28, 325-353.

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Flemström, G. and Garner, A. (1980). Stimulation of gastric acid and bicarbonate secretions by calcium in guinea pig stomach and amphibian isolated mucosa. Acta Physiol. Scand., 110, 419-426. Flemström, G., Heylings, J.R. and Garner, A. (1982). Gastric and duodenal H C 03~ transport in vitro: effects of hormones and local transmitters. Am. J. Physiol., 242, G100-G110. Flemström, G. and Jedstedt, G. (1989). Stimulation of duodenal mucosal bicarbonate secretion in the rat by brain peptides. Gastroenterology, 97, 412-420. Flemström, G. and Sachs, G. (1975). Ion transport by amphibian antrum in vitro. I. General characteristics. Am. J. Physiol., 228, 1188-1198. Fong, J.C., Hong, S.-J. and Wang, F.-F. (1991). Somatostatin inhibits pepsinogen secretion via a cyclic AMP-independent pathway. Regul. Peptides, 31, 351-360. Forte, J.G. and Wolosin, J.M. (1987). HC1 secretion by the gastric oxyntic cell. In Physiology o f the Gastrointestinal Tract. 2nd ed, edited by L.R. Johnson, J. Christensen, E.D. Jacobson, M.J. Jackson and J.H. Walsh, pp. 853-863. New York: Raven Press. Friedman, M.H.F. (1937). Oesophageal and gastric secretion in the frog. J. Cell. Comp. Physiol., 10, 37-50. Gerstberger, R. (1988). Functional vasoactive intestinal polypeptide (VlP)-system in salt glands of the Pekin duck. Cell Tissue Res., 252, 39-48. Gerstberger, R., Sann, H. and Simon, E. (1988). Vasoactive intestinal peptide stimulates blood flow and secretion of avian salt glands. Am. J. Physiol., 255, R575-R582. Golding, D.W. and Pow, D.V. (1990). ‘Neurosecretion’ by synaptic terminals and glandular discharge in the endocrine pancreas. Neuroendocrinology, 51, 369-375. Gullo, L., Priori, P., Baldoni, F. and Mattioli, G. (1986). Stimulation of exocrine pancreatic secretion by met-enkephalin. Gut, 27, 1479-1483. Häkanson, R., Böttcher, G., Ekblad, E., Panula, P., Simonson, M., Dohlsten, M., Hallberg, T. and Sundler, F. (1986). Histamine in endocrine cells in the stomach. A survey of several species using a panel of histamine antibodies. Histochem., 86, 5-17. Helander, H.F. (1981). The cells of the gastric mucosa. Int. Rev. Cytol, 70, 217-289. Hermansen, K. (1983). Enkephalins and the secretion of pancreatic somatostatin and insulin in the dog: Studies in vitro. Endocrinology, 113, 1149-1154. Hersey, S.J. (1987). Pepsinogen secretion. In Physiology o f the Gastrointestinal Tract. 2nd ed., edited by L.R. Johnson, J. Christensen, E.D. Jacobson, M.J. Jackson and J.H. Walsh, pp. 947-957. New York: Raven Press. Herzig, K.-H., Louie, D.S. and Owyang, C. (1988). In vivo action of bombesin on exocrine pancreatic secretion in the rat: Independent of cholecystokinin and cholinergic mediation. Pancreas, 3, 292-296. Hiramatsu, K. and Watanabe, T. (1989). Immunohistochemical study on the distribution of vasoactive intestinal polypeptide (VIP) containing nerve fibers in the chicken pancreas. Z. Mikrosk.-Anat. Forsch., 103, 689-699. Hirschowitz, B.I., Uemura, N., Matsumoto, H. and Dickinson, K.E.J. (1990). The role of Ca2+ in the time-dependent pepsinogen secretion of frog oesophageal peptic glands stimulated by bombesin. Acta Physiol. Scand., 140, 401-412. Holmgren, S. (1985). Substance P in the gastrointestinal tract of Squalus acanthias. Molec. Physiol, 8, 119-130. Holmgren, S., Jönsson, A.-C. and Holstein, B. (1986). Gastrointestinal peptides in fish. In Fish Physiology: Recent advances, edited by S. Nilsson and S. Holmgren, pp. 119-139. London: Croom Helm. Holmgren, S. and Nilsson, S. (1983a). Bombesin-, gastrin/CCK-, 5-hydroxytryptamine-, neurotensin-, somatostatin-, and VIP-like immunoreactivity and catecholamine fluorescence in the gut of the elasmobranch, Squalus acanthias. Cell Tissue Res., 234, 595-618. Holmgren, S. and Nilsson, S. (1983b). VIP-, bombesin-, and neurotensin-like immunoreactivity in neurons of the gut of the holostean fish, Lepisosteus platyrhincus. Acta Zoologica, 64, 25-32. Holmgren, S., Vaillant, C. and Dimaline, R. (1982). VIP-, substance P-, gastrin/CCK-, bombesin-, somatostatin-, and glucagon-like immunoreactivities in the gut of the rainbow trout, Salmo gairdneri. Cell Tissue Res., 223, 141-153. Holstein, B. (1975). Gastric acid secretion in a teleostean fish: A method for the continuous collection of gastric effluence from a swimming fish and its response to histamine and pentagastrin. Acta Physiol. Scand., 95, 417-423. Holstein, B. (1976). Effect of the H2-receptor antagonist metiamide on carbachol-and histamine-induced gastric acid secretion in the Atlantic cod, Gadus morhua. Acta Physiol. Scand., 97, 189-195.

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Masini, M.A. (1988). Immunocytochemical localization of peptides in the endocrine pancreas of the snakes Vipera aspis and Natrix maura. Acta. Histochem., 84, 111-119. McDonald, J.K., Greiner, F., Wood, J.G. and Noe, B.D. (1987). Oxytocin-like immunoreactive nerves are associated with insulin-containing cells in pancreatic islets of anglerfish (Lophius americanus). Cell Tissue Res., 249, 7-12. Milgram, S.L., McDonald, J.K. and Noe, B.D. (1991). Neuronal influence on hormone release from anglerfish islet cells. Am. J. Physiol., 261, E444-E456. Miller, R.E. (1981). Pancreatic neuroendocrinology: Peripheral neural mechanisms in the regulation of the islets of Langerhans. Endocrinology reviews, 2, 471-494. Neutra, M.R. and Forstner, J.F. (1987). Gastrointestinal mucus: synthesis, secretion and function. In Physiology o f the Gastrointestinal Tract. 2nd ed, edited by L.R. Johnson, J. Christensen, E.D. Jacobson, M.J. Jackson and J.H. Walsh, pp. 975-1007. New York: Raven Press. Noe, B.D., McDonald, J.K., Greiner, F., Wood, J.G. and Andrews, P.C. (1986). Anglerfish islets contain NPY immunoreactive nerves and produce the NPY analog aPY. Peptides, 7, 147-154. Ohtani, R., Kaneko, T., Kline, L.W., Labedz, T., Tang, Y. and Pang, P.K.T. (1989). Localization of calcitonin-gene-related peptide in the small intestine of various vertebrate species. Cell Tissue Res., 258, 35-42. Plisetskaya, E.M. (1989). Pancreatic peptides. In The Comparative Physiology o f Regulatory Peptides, edited by S. Holmgren, pp. 174-202. London, New York: Chapman and Hall. Rajjo, I.M., Vigna, S.R. and Crim, J.W. (1989a). Immunohistochemical localization of bombesin-like peptides in the digestive, tract of the bowfin, Amia calva. Comp. Biochem. Physiol., 94C, 405-409. Rajjo, I.M., Vigna, S.R. and Crim. J.W. (1989b). Immunocytochemical localization of vasoactive intestinal polypeptide in the digestive tracts of a holostean and a teleostean fish. Comp. Biochem. Physiol., 94C, 411-418. Reinecke, M., Schlüter, P., Yanaihara, N. and Forssmann, W.G. (1981). VIP immunoreactivity in enteric nerves and endocrine cells of the vertebrate gut. Peptides, 2, suppl. 2, 149-156. Ruiz, M.-C. and Michelangeli, F. (1984). Evidence for a direct action of acetylcholine on the gastric oxyntic cell of the amphibian. Am. J. Physiol., 246, G16-G25. Schmidt-Nielsen, K. (1960). The salt secreting glands of marine birds. Circulation, X X I, 955-966. Schmidt-Nielsen, K. and Fänge, R. (1958). Salt glands in marine reptiles. Nature, 182, 783-785. Schultzberg, M., Hökfelt, T., Nilsson, G., Terenius, L., Rehfeld, J.F., Brown, M., Eide, R., Goldstein, M. and Said, S. (1980). Distribution of peptide-and catecholamine-containing neurons in the gastrointestinal tract of rat and guinea-pig: Immunohistochemical studies with antisera to substance P, vasoactive intestinal polypeptide, enkephalins, somatostatin, gastrin/cholecystokinin, neurotensin and dopamine-/3-hydroxylase. Neuroscience, 5, 689-744. Seidler, U. and Sewing, K.-F. (1989). Ca2+-dependent and -independent secretagogue action on gastric mucus secretion in rabbit mucosal explants. Am. J. Physiol., 256, G739-G746. Sewing, K.-F. and Ruoff, H.J. (1972). Control of gastric acid secretion in chickens. Acta Hepatogastroenterol., 19, 296-300. Shirakawa, T. and Hirschowitz, B.I. (1984). Bombesin induced pepsinogen secretion from frog esophagus peptic glands in vitro. Gastroenterology, 8 6 , 1250. Shuttleworth, T.J. and Thorndyke, M.C. (1984). An endogenous peptide stimulates secretory activity in the elasmobranch rectal gland. Science, 225, 319-321. Silva, P., Lear, S. and Epstein, F.H. (1988). Bombesin inhibits stimulated chloride secretion by the rectal gland of Squalus acanthias. Clin. Res., 36, 598A. Silva, P., Lear, S., Reichlin, S. and Epstein, F.H. (1990). Somatostatin mediates bombesin inhibition of chloride secretion by rectal gland. Am. J. Physiol., 258, R1459-R1463. Silva, P., Stoff, J.S., Leone, D.R. and Epstein, F.H. (1985). Mode of action of somatostatin to inhibit secretion by shark rectal gland. Am. J. Physiol, 249, R329-R334. Simpson, L., Goldenberg, D. and Hirschowitz, B.I. (1980). Pepsinogen secretion by the frog esophagus in vitro. Am. J. Physiol, 238, G79-G84. Smit, H. (1968). Gastric secretion in the lower vertebrates and birds. In Handbook o f Physiology, sect 6: Alimentary canal', volume V: Bile, digestion, ruminal physiology, edited by C.F. Code, pp. 2791-2805. Am. Physiol. Soc. Soll, A.H. and Berglindh, T. (1987). Physiology of isolated gastric glands and parietal cells: receptors and effectors regulating function. In Physiology o f the Gastrointestinal Tract. 2nd ed, edited by L.R. Johnson, J. Christensen, E.D. Jacobson, M.J. Jackson and J.H. Walsh, pp. 883-909. New York: Raven Press. Stoff, J.S., Rosa, R., Hallac, R., Silva, P. and Epstein, F.H. (1979). Hormonal regulation of active chloride transport in the dogfish rectal gland. Am. J. Physiol, 237, F138-F144.

192 COMPARATIVE PHYSIOLOGY AND EVOLUTION Tache, Y. (1987). Central nervous system regulation of gastric acid secretion. In Physiology o f the Gastrointestinal Tract. 2nd ed, edited by L.R. Johnson, J. Christensen, E.D. Jacobson, M.J. Jackson and J.H. Walsh, pp. 911-930. New York: Raven Press. Tagliafierro, G., Faraldi, G. and Pestarino, M. (1985). Interrelationships between somatostatin-like cells and other endocrine cells in the pancreas of some cartilaginous fish. Cell Molec. Biol., 31, 201-207. Takeuchi, K., Magee, D., Critchlow, J., Matthews, J. and Silen, W. (1983). Studies of the pH gradient and thickness of frog gastric mucus gel. Gastroenterology, 84, 331-340. Thorndyke, M.C. and Shuttleworth, T.J. (1985). Biochemical and physiological studies on peptides from the elasmobranch gut. Peptides, 6, suppl. 3, 369-372. Van Noorden, S. and Patent, G.J. (1980). Vasoactive intestinal polypeptide-like immunoreactivity in nerves of the pancreatic islet in the teleost fish, Gillichtys mirabilis. Cell Tissue Res., 212, 139-146. Varga, G., Papp, M. and Vizi, E.S. (1990). Cholinergic and adrenergic control of enzyme secretion in isolated rat pancreas. Dig. Dis. Sci., 35, 501-507. Varga, G., Papp, M., Dobronyi, I. and Scarpignato, C. (1988). Effect of bombesin and its mammalian counterpart, GRP, on exocrine pancreas in the rat. Digestion, 41, 229-236. Vigna, S.R. and Thorndyke, M.C. (1989). Bombesin. In The Comparative Physiology o f Regulatory Peptides, edited by S. Holmgren, pp. 34-60. London: Chapman and Hall. Wallace, J.L. (1989). Gastric resistance to acid: is the “mucus-bicarbonate barrier” functionally redundant? Am. J. Physiol., 256, G31-G38. Walsh, J.H. (1987). Gastrointestinal hormones. In Physiology o f the Gastrointestinal Tract. 2nd ed, edited by L.R. Johnson, J. Christensen, E.D. Jacobson, M.J. Jackson and J.H. Walsh, pp. 181-253. New York: Raven Press. Watanabe, T., Chikazawa, H., Chungsamamyart, N., Fujioka, T. and Yamada, J. (1987). Serotoninstoring cells of the chicken duodenum: light, fluorescence and electron microscopy, and immunohistochemistry. Cell Tissue Res., 247, 25-32. Watson, A.H.D. (1979). Fluorescent histochemistry of the teleost gut: Evidence for the presence of serotonergic neurones. Cell Tissue Res., 197, 155-164. Williams, J.A. (1975). An in vitro evaluation of possible cholinergic and adrenergic receptors affecting pancreatic amylase secretion. Proc. Soc. Exp. Biol. Med., 150, 513-516. Woods, S.C. and Porte, D. (1974). Neural control of the endocrine pancreas. Physiol. Rev, 54, 596-619. Yasugi, S. (1987). Pepsinogen-like immunoreactivity among vertebrates: Occurrence of common antigenicity to an anti-chicken pepsinogen antiserum in stomach gland cells of vertebrates. Comp. Biochem. Physiol., 86B, 675-680. Yasugi, S., Matsunaga, T. and Mizuno, T. (1989). Pepsinogen-like immunoreactivity in ascidian stomach and intestine: Immunohistochemical and biochemical study. Zool. Sci., 6 , 283-288. Yasugi, S., Matsunaga, T. and Mizuno, T. (1988). Presence of pepsinogens immunoreactive to antiembryonic chicken pepsinogen antiserum in fish stomachs: Possible ancestor molecules of chymosin of higher vertebrates. Comp. Biochem. Physiol., 91A, 565-569. Yoshida, K., Iwanaga, T. and Fujita, T. (1983). Gastro-entero-pancreatic (GEP) endocrine system of the flatfish, Paralichtys olivaceus: An immunocytochemical study. Arch. Histol. Jap., 46, 259-266. Yui, R. and Fujita, T. (1986). Immunocytochemical studies on the pancreatic islets of the ratfish Chimaera monstrosa. Arch. Histol. Jap., 49, 369-377. Yui, R., Nagata, Y. and Fujita, T. (1988). Immunocytochemical studies on the islet and the gut of the Arctic lamprey, Lampetra japonica. Arch. Histol. Cytol., 51, 109-119. Yui, R., Shimada, M. and Fujita, T. (1990). Immunohistochemical studies on peptide-and aminecontaining endocrine cells and nerves in the gut and rectal gland of the ratfish Chimaera monstrosa. Cell Tissue Res., 260, 193-201.

6 The Circulatory System Judy L. Morris1 and Stefan Nilsson 2 1Department o f Anatomy & Histology, and Centre fo r Neuroscience, School o f Medicine, Flinders University o f South Australia, Adelaide, South Australia; 2Department o f Zoophysiology, University o f Göteborg, Göteborg, Sweden ABBREVIATIONS ATP adenosine 5 '-triphosphate CCK cholecystokinin IR immunoreactive, immunoreactivity NPY neuropeptide Y VIP vasoactive intestinal polypeptide The autonomic nervous system provides an important mechanism for regulating the cardiovascular system in vertebrates. Some features of the autonomic innervation are common to nearly all vertebrates. These include an inhibitory vagal innervation of the heart and a constrictor innervation of at least some blood vessels by spinal autonomic neurons that synthesize catecholamines. However, there seems to be an evolutionary trend for cardiac excitation to be achieved by catecholamines released from endogenous or nearby chromaffin tissue in cyclostomes, elasmobranchs and dipnoans, but to be mediated primarily by spinal autonomic neurons in teleosts and most tetrapods. All vertebrates have developed reflexes that modify the cardiovascular system in response to stimuli such as hypoxia and exercise. The neuroeffector mechanisms used to achieve autonomic effects seem to vary between parts of the cardiovascular system and between species. Cotransmission involving “classical transmitters”, peptides and purines is widespread, although there are no obvious evolutionary trends in the neurotransmitters used by autonomic neurons controlling homologous structures. KEY WORDS heart; blood vessels; cranial nerves; sympathetic ganglia; chromaffin tissue; cardiovascular reflexes; neurotransmitters

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194 COMPARATIVE PHYSIOLOGY AND EVOLUTION

INTRODUCTION Without doubt, one very strong force in the evolution of the vertebrate organisms is the need to maintain an adequate supply of oxygen to the tissues during different physiological circumstances. Thus, air breathing as a secondary mechanism for acquiring oxygen has evolved independently in several groups of fish, necessitating modifications of the organs responsible for the exchange of gases. Central to the maintenance of adequate oxygenation of the tissues has been the development of effective circulatory systems for the efficient transport of respiratory gases around the body. Furthermore, several mechanisms have developed in the vertebrates to control the flow of blood to respiratory and to non-respiratory organs. These control mechanisms include: (1) the local production of cardioactive or vasoactive substances; (2) the release of endocrine hormones into the circulation and their subsequent cardiovascular actions remote from the site of production; (3) neuronal mechanisms that can produce either brief or long-lasting changes in the flow of blood in discrete regions of the cardiovascular system in response to particular physiological stimuli. The importance of the autonomic nervous system in the regulation of the vertebrate cardiovascular system has been recognized for more than a century (e.g. Gaskell, 1883, 1884, 1886). During this time there has been a wealth of anatomical and physiological studies elucidating the distribution of autonomic nerves throughout the cardiovascular system, and the actions of the nerves on the heart and blood vessels. Many physiological studies have detected autonomic effects by measuring cardiac output, systemic blood pressure or regional blood flow in whole animals, in response to agonist drugs or to stimulation of autonomic pathways. While this information is crucial for establishing the outcomes of the activation of autonomic nerves, it must be supplemented by studies on isolated organs and individual blood vessels in order for us to understand the exact sites and mechanisms of neuroeffector transmission used to achieve specific physiological effects. The development of histochemical methods for the demonstration of monoamines in the 1960s (Falck et al., 1962) enabled significant advancements to be made concerning the sites of cardiovascular regulation by catecholaminergic neurons. Although information on mammals accumulated much faster than that on other vertebrates, several groups, notably Geoffrey Burnstock and colleagues at Melbourne University, applied these new histochemical techniques to studies of the autonomic innervation across the vertebrates. They combined histochemistry with physiological and pharmacological studies, and developed new concepts on the evolution of the autonomic control mechanism (Campbell and Burnstock, 1968; Burnstock, 1969; Campbell, 1970; Berger and Burnstock, 1979). This multidisciplinary approach has also been applied by other research groups, particularly to studies of the circulation in fish (Nilsson, 1983, 1984, 1986; Nilsson and Holmgren, 1988, 1991, 1992). The more recent discovery of neuropeptides in the autonomic nervous system, and the development of immunohistochemical techniques, has lead to a new phase of experimentation on cardiovascular control mechanisms. Again, new information

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has accumulated much faster for mammals than for non-mammalian vertebrates. We are now beginning to understand the roles of neuropeptides in transmission from autonomic neurons to the cardiovascular system in mammals and the physiological significance of having multiple transmitters in the same neurons (see Morris and Gibbins, 1992). Although the functions of many neuropeptides are still not known, particularly in non-mammals, immunohistochemistry of peptides provides a very useful technique for mapping pathways of autonomic neurons that could not be detected with previous histochemical methods (Gibbins, 1989). Indeed, the innervation of some tissues by certain classes of autonomic neurons has been demonstrated for the first time using immunohistochemistry of neuropeptides. This chapter provides a review of the literature on the distribution, histochemistry, pharmacology and physiological functions of autonomic neurons in the cardiovascular systems of vertebrates. In most cases there is not complete information on all aspects of the autonomic innervation of a particular part of the circulation in the same species. Furthermore, the existing information is often restricted to a few well-studied species in a particular class of vertebrates. Consequently, the information presented here is largely restricted to examples that allow comparisons to be made between the innervation of homologous cardiovascular targets across the vertebrates.

AUTONOM IC INNERVATION OF THE HEART The heart of cyclostomes, elasmobranchs and actinopterygians (including the ganoids and teleosts) consists of three or four chambers connected in series: the sinus venosus, the atrium, the ventricle and, in elasmobranchs and some primitive bony fish, the bulbus cordis (sometimes also called the conus arteriosus). In teleost fish the pulse-smoothing (“wind-kessel”) function of the bulbus cordis is carried out by the elastic proximal portion of the ventral aorta, the bulbus arteriosus (Satchell, 1991). In the cyclostomes, myxinoids (hagfish) and lampetroids (lampreys), the circulation of blood is facilitated by “branchial hearts”, and by the action of a pump in the caudal venous sinus, “caudal heart”, both of which consist of non-vascular tissue. In myxinoids, the action of the systemic heart is further aided by the pulsatile “portal heart”, which is situated in the hepatic portal vein, and pumps blood into the liver (Fänge, Johnels and Engers 1963; Satchell, 1984, 1986; Davie et al., 1987; Forster et al., 1991). The heart of some fish, notably elasmobranchs, is encased in a rigid pericardium, which causes a negative pressure in the atrium during ventricular contraction. Blood from the sinus venosus thus enters the atrium due to suction of the blood (vis a fronte), a process different from that in the tetrapods where the central venous pressure determines the atrial filling (vis a tergo). In fish, but also in amphibians and to some extent in reptiles, ventricular filling (and thus cardiac stroke volume), depends very directly on the atrial filling. As will be seen later, it is clear that the

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autonomic nervous control of the stroke volume is chiefly aimed at atrial control (Johansen, 1971; Johansen and Burggren, 1980). The heart of lungfish shows anatomical characteristics not present in other fish. Thus the pulmonary vein enters the atrium directly, and there is a high degree of separation between the oxygenated and deoxygenated blood during the passage through the heart, despite the anatomically single ventricle (see Johansen and Burggren, 1980; Nilsson and Holmgren, 1991). In amphibians and all groups of reptiles except the crocodilians, the heart possesses two atria and a single ventricle. Despite the single ventricle, blood flpw from the right and left atria remain remarkably separate due to the trabeculae within the ventricle and the organization of the outflow vessels. In varanid lizards (goannas, monitors), the ventricular compartments are isolated by a muscular ridge during systole, allowing different pulmonary and systemic blood pressures despite the anatomically continuous (at diastole) ventricle (Johansen, 1971; Johansen and Burggren, 1980; Nilsson, 1992). In crocodilians, the heart has four chambers, just as in birds and mammals. However, a peculiar arrangement of the outflow tract and the aortic arches, including a foramen between the bases of the left and right aorta (foramen Panizzae), allows right-to-left blood shunting when the pulmonary vascular resistance increases (see, e.g., Axelsson, Holm and Nilsson, 1989; Grigg, 1989; Nilsson, 1994). CYCLOSTOME HEART Innervation by cranial autonomic nerves The heart of lampetroids receives a functional vagal innervation, which runs along the jugular vein to the sinus venosus, or enters along the bulbus cordis (Carlson, 1906; Tretjakoff, 1927; Augustinsson et al., 1956; Johnels, 1956; Beringer and Hadek, 1972). In contrast to the pattern in all other vertebrates, this vagal innervation is mainly excitatory, acting via nicotinic cholinoceptors (Carlson, 1906; Zwaardemaker, 1924; Otorii, 1953; Augustinsson et al., 1956; Falck et al., 1966; Lukomskaya and Michelson, 1972). A small effect of the agonists of the muscarinic cholinoceptor, pilocarpine and choline muscarine, was observed by Otorii (1953) on the heart of Endoshenus japonicus, but not by Falck et al. (1966) on the heart of Lampetra. The systemic heart of myxinoids (Myxine, Eptatretus) appears to lack a functional external autonomic innervation, although some work suggests the presence of neurons within the heart of Eptatretus (Greene, 1902; Augustinsson et al., 1956; Hirsch, Jellinek and Cooper, 1964). The isolated heart of Myxine is insensitive to many exogenously applied drugs, including acetylcholine and the catecholamines (Fange and Ostlund, 1954; Ostlund, 1954; Augustinsson et al., 1956). Chromaffin tissue One striking feature of the systemic (and portal) heart of cyclostomes is the storage of large quantities of adrenaline and noradrenaline within the heart. The cate-

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cholamines are stored within specialized cells containing myocardial granules, and there is a weak chromaffin reaction of the myocardia of Myxine and Lampetra (Augustinsson et al., 1956). Formaldehyde-induced fluorescence histochemistry (Falck-Hillarp technique) reveals intensely fluorescent cells embedded in the myocardium (Dahl et al., 1971; Shibata and Yamamoto, 1976). The catecholamine-storing issue of the lampetroids appears to receive no extrinsic innervation (Caravita and Coscia, 1966; Beringer and Hadek, 1972), and the mode of control of these cells is not known. Catecholamines and tyramine stimulate the lampetroid heart, and experiments with pronethalol suggest that the effect is mediated by /3-adrenoceptors (Otorii, 1953; Augustinsson et al., 1956; Nayler and Howells, 1965; Falck et al. 1966). The isolated heart of Myxine is remarkably insensitive to exogenously applied catecholamines. However, reserpine and dihydroergotamine produce negative inotropic and chronotropic effects, possibly by disturbing the stimulating effects of catecholamines released from the endogenous chromaffin tissue (Fange and Ôstlund, 1954; Ôstlund, 1954; Augustinsson et al., 1956; Bloomed al., 1961). More recent experiments with intact, unanaesthetized Myxine showed marked cardiostimulatory effects of injected catecholamines, and the /3-adrenoceptor antagonist, sotalol, produced a strong negative chronotropic effect. This further supports the idea that endogenously released catecholamines from the chromaffin stores within the heart are of importance for normal cardiac function (Axelsson, Farrell and Nilsson, 1990). ELASMOBRANCH HEART Innervation by cranial autonomic nerves The elasmobranch heart receives a well-developed inhibitory vagal innervation, which acts via muscarinic cholinoceptors, as in all vertebrates except the cyclostomes. The vagal innervation reaches the heart in two distinct pairs, which run from the visceral and posterior branchial branches, respectively (Norris and Hughes, 1920). There is a wealth of studies that demonstrate that the effect of acetylcholine mimics that of vagal stimulation, and that both effects are blocked by the antagonist of the muscarinic cholinoceptor, atropine (for references, see Laurent, Holmgren and Nilsson, 1983; Nilsson, 1983; Nilsson and Holmgren, 1988). Studies of the heart of the dogfish, Scyliorhinus canicula, demonstrate the presence of P r purinoceptors, which mediate negative inotropic and chronotropic effects in the atrium but not in the ventricle (Meghji and Burnstock, 1984a). The presence and function of neuropeptides in autonomic nerves of the elasmobranch heart are not yet well understood, and the true origin of the fibres (cranial or spinal autonomic system) is not at all clear. A sparse innervation by bombesinimmunoreactive (IR) fibres has been demonstrated in the heart and coronary vessels of the little skate, Raja erinacea (Bjenning, Farrell and Holmgren, 1991). In Raja erinacea and Raja radiata, the sinus venosus and the atrium receive a sparse

198 COMPARATIVE PHYSIOLOGY AND EVOLUTION

innervation by neuropeptide Y (NPY)-IR nerve fibres, and in R. erinacea there is also a moderately dense innervation of the ventricle (Bjenning, Driedzic and Holmgren, 1989). Innervation by spinal autonomic nerves There are, with one possible exception (Mustelus; Pick, 1970), no visible connections between the spinal autonomic system and the vagi in elasmobranchs, and there is no functional adrenergic innervation of the elasmobranch heart (see Laurent, Holmgren and Nilsson, 1983; Nilsson and Holmgren, 1988). However, in Scyllium, the sinoatrial conduction time decreased upon electrical stimulation of the anterior paravertebral ganglia after atropine (Izquierdo, 1930), and sparse adrenergic fibres in the sinus venosus have been described in the Port Jackson shark, Heterodontus portusjacksoni (Gannon, Campbell and Satchell, 1972). If an adrenergic control of the elasmobranch heart does occur, it must be due chiefly to circulating catecholamines, released from the chromaffin tissue within the posterior cardinal sinuses (Satchell, 1970, 1971; Gannon, Campbell and Satchell, 1972; Abrahamsson, 1979). Positive inotropic and to some extent also chronotropic effects of catecholamines, mediated via ß-adrenoceptors, have been reported. A transient, atropine-sensitive bradycardia in response to catecholamines has also been reported, which suggests a cholinergic element in the response (Lutz, 1930a-c; Hiatt, 1943; Fänge and Östlund, 1954; Capra and Satchell, 1977; Nilsson and Holmgren, 1988). TELEOST HEART Innervation by cranial autonomic nerves The teleost heart is innervated by one pair of cardiac branches from the vagus, which enter the sinus venosus. A ganglion which consists mainly or solely of nonadrenergic cell bodies lies close to the sinoatrial border, and postganglionic vagal fibres innervate the sinus and atrium. The density of nerves is by far highest in the sinoatrial nerve plexus (Laurent, 1962; Yamauchi and Burnstock, 1968; Gannon and Burnstock, 1969; Sanier, 1972; Santer and Cobb, 1972; Holmgren, 1977, 1981). The innervation of the ventricle varies betweeen species, being most dense in the atrioventricular border and more sparse in the ventricular myocardium (Laurent, 1962; Yamauchi and Burnstock, 1968; Santer, 1972). Numerous studies have shown that the vagal fibres are inhibitory, producing negative chronotropic and inotropic effects mediated by muscarinic cholinoceptors associated with the pacemaker and atrium (for references, see Cameron, 1979; Wood, Pieprzak and Trott, 1979; Laurent, Holmgren and Nilsson, 1983; Nilsson, 1983; Nilsson and Holmgren, 1988). It has been suggested that the vagal cholinergic control of the heart is most important at lower temperatures, while adrenergic mechanisms become more important at higher temperatures (Laffont and Labat, 1966; Priede, 1974; Wood, Pieprzak and Trott, 1979). Acetylcholine and related drugs produce negative inotropic effects on atrial, but generally not on ventricular strip preparations in vitro. As stated above, however, a vagal control of atrial

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contractility would be sufficient to affect ventricular filling and thus the cardiac stroke volume (Jones and Randall, 1978; Johansen and Burggren, 1980). Studies of the heart of the rainbow trout (Salmo gairdneri) demonstrated negative inotropic and positive chronotropic effects of adenosine and adenosine 5 '-triphosphate (ATP) on the atrium. The effects were insensitive to a Pi-antagonist, suggesting that the purinoceptor of the heart of the rainbow trout differs from the type found in other vertebrates (Meghji and Burnstock, 1984b). The presence, origin and functions of peptidergic nerves in the heart of the teleost are not well known, as in elasmobranchs. Injection of vasoactive intestinal polypeptide (VIP) in vivo has been shown to produce an increase in the cardiac stroke volume in the cod, Gadus morhua (Jensen, Axelsson and Holmgren, 1991). Innervation by spinal autonomic nerves An adrenergic innervation of the teleost heart occurs in most species studied (Govyrin and Leont’eva, 1965; Gannon, 1969, 1971; Gannon and Burnstock, 1969; Holmgren, 1977; Cameron, 1979; Donald and Campbell, 1982), but appears to be absent in pleuronectids (Falck et al., 1966; Santer, 1972; Cobb and Santer, 1973; Donald and Campbell, 1982). An adrenergic innervation of the heart is similarly absent in the sturgeon (Huso huso), but is present in another ganoid, Lepisosteus platyrhincus (Balashov et al., 1981; Nilsson, 1981). When present, the adrenergic nerve fibres can enter the heart along three routes: (1) by entering the vagi via the connections with the cephalic sympathetic chains (to form “vagosympathetic trunks”); (2) along the anterior pair of spinal nerves; or (3) along the coronary arteries (Gannon and Burnstock, 1969; Holmgren, 1977). The adrenergic innervation is excitatory, producing positive inotropic and chronotropic effects mediated by /3-adrenoceptors (Fänge and Östlund, 1954; Östlund, 1954; Randall and Stevens, 1967; Gannon and Burnstock, 1969; Forster, 1976; Holmgren, 1977; Cameron and Brown, 1981; Axelsson et al., 1989b). Adrenergic fibres, in some cases possibly enhanced by circulating catecholamines, exert a tonic influence on the heart of most vertebrates studied, including teleosts (Cameron, 1979; Cameron and Brown, 1981; Axelsson, 1988; Axelsson et al., 1989b). DIPNOAN HEART The heart of the lungfish receives a vagal innervation which is inhibitory and cholinergic, as in the majority of vertebrates. However, a contribution of autonomic fibres of spinal origin seems unlikely in view of the poorly developed sympathetic chains in this group (Jenkin, 1928; Abrahamsson et al., 1979b). Carbachol produces negative inotropic effects on the atrium, but not the ventricle, of Protopterus (Abrahamsson et al., 1979b), but a cholinergic tonus on the heart of Protopterus in vivo appears to be lacking, as atropine was without effect on the heart rate in the intact animal (Johansen and Reite, 1968). In the South American lungfish, Lepidosiren paradoxa, there is a cholinergic cardiac tonus, which varies with the breathing cycles allowing an increased cardiac output in relation to a breath of air (Axelsson et al., 1989a). A small cholinergic vagal tonus has been demonstrated in

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the Australian lungfish, Neoceratodus forsteri, and there is also a similarly small adrenergic tonus affecting the heart at rest (R. Fritsche, M. Axelsson, G.C. Grigg, C.E. Franklin and S. Nilsson, unpublished). A remarkable feature of the heart in all three dipnoan genera (Protopterus, Lepidosiren, Neoceratodus) is the presence of large quantities of catecholamines in endothelial cells of the atrium (Abrahamsson et al., 1979a; Scheuermann, 1979; Scheuermann et al. 1981; Axelsson et al. 1989a; S. Holmgren, unpublished). The role of these cells in cardiac control remains obscure, but an adrenergic tonus on the heart has been demonstrated in Lepidosiren (Axelsson et al. 1989a). AMPHIBIAN HEART The heart in most amphibian species, like that in the other tetrapods, is innervated both by cranial autonomic neurons in vagal pathways (Figure 6.1), and by spinal autonomic neurons originating in paravertebral sympathetic ganglia (Figure 6.3). The postganglionic sympathetic nerve fibres join the vagal preganglionic nerve fibres bilaterally to form vagosympathetic nerve trunks which enter the heart at the junction of the venae cavae and the sinus venosus. Varicose processes of smalldiameter sensory neurons are also abundant in the hearts of tetrapods (Figure 6.3). Careful anatomical and functional studies are required to unequivocally distinguish between sensory and autonomic nerve fibres within the heart (see Gibbins, Furness and Costa, 1987). Innervation by cranial autonomic nerves Until recently, there has been no anatomical method that has clearly demonstrated the distribution of postganglionic vagal neurons within the heart. The recent immunohistochemical localization of neuropeptides in intracardiac neurons in an anuran (JBufo marinus) and in a urodele (Necturus maculosus), has enabled precise mapping of the postganglionic vagal neurons in these species. Somatostatin-IR occurs in nearly all intracardiac neuronal cell bodies in the toad Bufo marinus (Campbell et al., 1982; Morris, Gibbins and Osborne, 1989; Figure 6.1a). Varicose nerve fibres with somatostatin-IR project from the neuronal cell bodies in the sinus venosus and intra-atrial septum, to all chambers of the heart and to the pericardium. Many somatostatin-IR axons are found associated with each cardiac muscle bundle. As somatostatin-IR has not been found in sympathetic or sensory neurons in Bufo marinus, and the density of somatostatin-IR nerve fibres in the heart was not altered after sectioning the vagosympathetic nerve trunks, it can be concluded that somatostatin-IR provides a good marker for postganglionic vagal neurons in this species. Immunoreactivity to the peptides VIP or galanin occurs in subpopulations of intracardiac neuronal cell bodies in Bufo marinus. Some neurons contain IR to both somatostatin and galanin (82% total neurons; Morris, Gibbins and Osborne, 1989; Figure 6.1), or to both somatostatin and VIP (25% total neurons; Gibbins et al., 1987), while all three peptides may coexist in a small proportion of intracardiac neurons. Galanin-IR is also found in many somatostatin-IR varicose nerve fibres

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FIGURES 6.1 and 6.2 Cell bodies of autonomic neurons innervating the heart, la, lb: Whole-mount of intracardiac vagal neurons in the inter-atrial septum of the toad (Bufo marinus), double-labelled to show immunoreactivity for somatostatin (Som) (la) and galanin (Gal) (lb). Many neurons have immunoreactivity for both peptides (arrows). Figure courtesy of I. Gibbins and J. Morris. 2a, 2b: Section of the stellate ganglion from the bearded dragon (Pogona vitticeps), double labelled to show immunoreactivity for tyrosine hydroxylase (TH) (2a) and galanin (2b). A subpopulation of catecholamine-synthesizing neurons contains galanin (arrows). Figure courtesy of I. Gibbins, J. Morris and S. Holmgren. Scale bar = 20 nm for all figures.

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throughout the heart, but VIP-IR has not been detected in the axons of intracardiac neurons (Campbell et al., 1982; Gibbins et al.f 1987). Somatostatin has not been found in intracardiac vagal neurons of all amphibians. Although it is present in Xenopus laevis, it is absent from several species of Australian frogs (Campbell, 1987), and from the mudpuppy (Necturus maculosus; Holmgren, cit in Axelsson and Nilsson, 1985). Instead, all intracardiac neuronal cell bodies in the mudpuppy contain galanin-IR (Parsons et al., 1989; McKeon and Parsons, 1990). Processes of the galanin-IR intracardiac neurons project to muscle bundles in all regions of the heart and to the epicardium (Parsons et al., 1989). Furthermore, a small proportion of intracardiac neuronal cell bodies seems to receive galanin-IR collaterals from other intracardiac neurons. About 40% of the small intrinsic neurons also contain galanin-IR and 50% of these have processes closely associated with the cell bodies of intracardiac neurons (McKeon and Parsons, 1990). The vagal neurons have negative chronotropic and inotropic effects in anurans and urodeles, and the inotropic effect is present in both the atria and the ventricle (Bidder, 1868; Gaskell, 1884; McWilliam, 1885; Elliott, 1905; Loewi, 1921; Kirby and Burnstock, 1969b; Woods, 1970; Campbell et al., 1982; Axelson and Nilsson, 1985). Detailed pharmacological studies in both of these species have demonstrated clearly that a large component of the inhibitory responses is mediated by acetylcholine acting on muscarinic receptors on the cardiac muscle cells. However, it has been found that the muscarinic receptors mediating vagal inhibitory responses in B. marinus are different from the muscarinic receptors mediating inhibitory effects of exogenous acetylcholine (Bywater et al., 1989). Furthermore, components of the vagal inhibition of the sinus venosus and atria of the toad, and of the heart of the mudpuppy, are mediated by non-cholinergic mechanisms (Campbell et al., 1982; Axelsson and Nilsson, 1985). Although both somatostatin and galanin are contained in intracardiac neurons in B. marinus, only somatostatin has negative inotropic and chronotropic actions in the sinus venosus and atria of this species (Campbell et al., 1982, G.D.S. Hirst, personal communication). Indeed, pharmacological studies have provided compelling evidence that somatostatin mediates the non-cholinergic vagal inhibition in the atria and sinus venosus of the toad that occurs on stimulation at frequencies above 2 Hz (Campbell et al., 1982). Galanin has negative inotropic and chronotropic effects on the heart of the mudpuppy, and hyperpolarizes the cardiac muscle (Parsons et al., 1989). Therefore, galanin is a candidate for mediating the non-cholinergic vagal inhibition of the heart of the mudpuppy. Furthermore, galanin produces hyperpolarization or depolarization of intracardiac postganglionic neuronal cell bodies (Konopka and Parsons, 1989; Konopka, McKeon and Parsons, 1989; Parsons et al., 1989). Thus, any galanin released from processes of small intrinsic neurons or postganglionic neurons in the vicinity of postganglionic cell bodies, may play a role in regulating the activity of the vagal postganglionic neurons.

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Innervation by spinal autonomic nerves Histochemical studies have shown a dense plexus of catecholamine-containing nerve fibres projecting from the paravertebral sympathetic ganglia to all chambers of the heart in anuran amphibians (Falck, Hággendal and Owman, 1963; Morris, Gibbins and Clevers, 1981). However, the myocardium of at least one urodele amphibian, the mudpuppy N. maculosus, is not innervated by catecholamine-synthesizing neurons (Axelsson and Nilsson, 1985). In this species, intracardiac fibres with catecholamine fluorescence are restricted to the vagal nerve trunks, some small blood vessels, and the atrioventricular border. It is not known whether the fibres at the atrioventricular border project from the small cells in the vicinity with intense catecholamine fluorescence (small intrinsic neurons), or from paravertebral sympathetic ganglia. The small intrinsic neurons have long processes that are closely associated with the cell bodies of the vagal postganglionic neurons (McMahon and Purves, 1976; Parsons and Neel, 1987), and have been suggested to act as interneurons (McMahon and Purves, 1976). These small neurons contain dopamine as their predominant catecholamine (McMahon and Purves, 1976), but many also contáin 5-hydroxytryptamine (Parsons and Neel, 1987). Adrenaline is the predominant catecholamine synthesized in, and released from, spinal autonomic neurons innervating the heart of anuran amphibians (Loewi, 1921, 1922, 1936; Azuma, Binia and Visscher, 1965). The majority of these neurons in B . marinus do not appear to contain any of the neuropeptides normally found in autonomic neurons. One exception is adrenergic nerves terminating within the intracardiac vagosympathetic nerve trunks, some of which contain NPY (Morris et al., 1986a). Stimulation of the spinal autonomic neurons innervating the anuran heart has positive inotropic and chronotropic effects. These excitatory responses can be abolished by the drug bretylium, which blocks adrenergic neurons (Morris, Gibbins and Clevers, 1981), and are reduced by high concentrations of ergot alkaloids (Bramich, Edwards and Hirst, 1990) or by the purine analogue, a,/3-methylene ATP (Hoyle and Burnstock, 1986; Bramich, Edwards and Hirst, 1990). /3-Adrenoceptors are widespread in the anuran heart, and mediate excitatory effects (Stene-Larson and Helle, 1978; Morris, Gibbins and Clevers, 1981; O’Donnell and Wanstall, 1982; Ask, 1983). However, the positive inotropic and chronotropic effects of sympathetic nerve stimulation can only partly be attributed to adrenaline acting on /3-adrenoceptors (Morris, Gibbins and Clevers, 1981; Hoyle and Burnstock 1986). The remaining sympathetic effects seem to have different causes in R. esculenta and in B. marinus. In the frog, ATP released from adrenergic nerve terminals is likely to mediate the faster excitatory events that are resistant to /3-adrenoceptor antagonists (Hoyle and Burnstock, 1986). However, in the toad, the fast component of the sympathetic excitation seems to be due to neuronally released adrenaline acting on specialized adrenoceptors located close to the sympathetic nerve terminals (Bramich, Edwards and Hirst, 1990).

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REPTILIAN HEART Innervation by cranial autonomic nerves The neuron cell bodies that lie along the intracardiac branches of the vagus nerve in the ratsnake Elaphe obsoleta, which presumably are postganglionic vagal neurons, are somatostatin-IR (Donald, O’Shea and Lillywhite, 1990a). Processes of these somatostatin-IR neurons project to the sinus venosus, atria and ventricle. Similarly, electron microscopic studies of the heart of the turtle indicate that nonadrenergic nerve terminals, presumed to be cholinergic (cf. Gibbins, 1982) and therefore of vagal origin, occurred in all of the cardiac chambers (Yamauchi, 1969; Yamauchi and Chiba, 1973). Functional studies have demonstrated that the vagal neurons have negative inotropic and chronotropic effects in all of the major groups of reptiles (Gaskell, 1883; Khalil and Malek, 1952; Berger, 1971; Hedberg and Nilsson, 1975; Berger and Burnstock, 1979; Donald, O’Shea and Lillywhite, 1990a). Indeed, continuous vagal stimulation of the heart of the turtle maintains cardiac arrest for long periods (Mills, 1885). The negative inotropic responses are largely restricted to the atria (Gaskell, 1883; Knowlton, 1942; Hedberg and Nilsson, 1976; Berger and Burnstock, 1979). Furthermore, blockade of the conduction of impulses from the pacemaker to the atria (Berger and Burnstock, 1979), and from the atria to the ventricle (Burggren, 1978), contributes to the cardioinhibitory action of the vagus. In all of the reptiles tested, the vagal effects on the heart can be mimicked by exogenous acetylcholine and are blocked by antagonists for muscarinic cholinergic receptors (de la Lande, Tyler and Pridmore, 1962; Berger, 1971; Hedberg and Nilsson, 1976; Berger and Burnstock, 1979; Donald, O’Shea and Lillywhite, 1990a). Thus, even though exogenous somatostatin has negative inotropic and chronotropic actions in the heart of the ratsnake, no acute role for endogenous somatostatin in vagal neurotransmission has been revealed (Donald, O’Shea and Lillywhite, 1990a). Innervation by spinal autonomic nerves Catecholamine-synthesizing autonomic neurons project from the paravertebral sympathetic ganglia to all chambers of the reptilian heart (Govyrin and Leont’eva, 1965; Furness and Moore, 1970; Hedberg and Nilsson, 1975; Berger and Burnstock, 1979; Donald, O’Shea and Lillywhite, 1990a). Catecholamine-synthesizing neurons in the stellate ganglion of some Australian lizards contain galanin (Figure 6.2), sometimes in addition to NPY (I. Gibbins, J. Morris and S. Holmgren, unpublished). The nerve fibres either run together with the vagus in a vagosympathetic nerve trunk, or project directly from the sympathetic chain to the heart (see Nilsson, 1983). In the ratsnake, some of these cardiac nerve fibres contain NPY-IR (Donald, O’Shea and Lillywhite, 1990a). Nerve fibres with catecholamine fluorescence may also surround the cell bodies of intracardiac neurons (Nilsson, 1983). The spinal autonomic neurons have positive inotropic and chronotropic effects on the reptilian heart, which are mediated by ^-adrenoceptors (van Han, Emaus and Meester, 1973; Hedberg and Nilsson, 1975, 1976; Berger and Burnstock, 1979).

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No purinoceptors were found in the heart of the turtle (Emys orbicularis; Meghji and Burnstock, 1983), hence any ATP released from sympathetic nerve terminals cannot contribute to cardioexcitation in this species (cf. R. esculenta). AVIAN HEART Innervation by cranial autonomic nerves The vagus nerves provide a dense innervation of all chambers of the avian heart (Bolton, 1971a; Bennett, 1974). They form cardiac plexuses in the region of the sinoatrial and atrioventricular nodes, where postganglionic neuronal cell bodies are located (Malinovsky, 1962). Some vagal postganglionic neuronal cell bodies are surrounded by axons with catecholamine fluorescence (Bennett and Malmfors, 1970). The vagus exerts a tonic inhibition of heart rate (Tummons and Sturkie, 1969), although the degree of vagal tone is variable between species. In addition to negative chronotropic effects, there are prominent negative inotropic responses in birds that can be attributed to the dense vagal innervation of the ventricles (Paton, 1912; Johansen and Reite, 1964; Bolton and Raper, 1966; Tummons and Sturkie, 1968, 1969, 1970; Yamauchi, 1969; Cohen et al., 1970). These negative chronotropic and inotropic responses, which are mediated by acetylcholine, are particularly prominent in diving birds (see Butler and Jones, 1982). It has been claimed that one or other of the vagal nerve trunks mediates diving bradycardia at any particular time (Butler and Jones, 1968), but this laterality could not be confirmed in later studies (see Butler and Jones, 1982). Innervation by spinal autonomic nerves Autonomic neurons providing an excitatory innervation to all chambers of the avian heart (Bennett and Malmfors, 1970) leave the spinal cord at the level of the last cervical ganglion (pigeon: MacDonald and Cohen, 1970), or the first thoracic ganglion (chicken: Tummons and Sturkie, 1969). The spinal autonomic neurons also contribute to the intracardiac plexuses formed by the vagal neurons. The cardiac neurons of spinal origin mediate positive chronotropic and inotropic responses via /3-adrenoceptors (Johansen and Reite, 1964; Enemar, Falck and H&kanson, 1965; Govyrin and Leont’eva, 1965; Bolton and Raper, 1966; Tummons and Sturkie, 1968, 1969, 1970; Akester, Akester and Mann, 1969; Yamauchi, 1969; Bennett and Malmfors, 1970; Bolton, 1971b). These excitatory neurons are also tonically active in the resting bird (Tummons and Sturkie, 1969), and thus, together with the vagal innervation, they determine the resting heart rate (Johansen and Reite, 1964). MAMMALIAN HEART Innervation by cranial autonomic nerves The cell bodies of vagal postganglionic neurons innervating the heart are located in an extensive cardiac nerve plexus, which is partly contained in the walls of the

ACh Spinal autonomic NA NPY Enk Dyn Cranial autonomic postganglionic ACh SST-28 VIP NPY SP CGRP

ACh

Spinal autonomic

Adr NPY

Cranial autonomic postganglionic

ACh SST-14 Gal (VIP)

Guinea-pig

FIGURE 6.3 Diagram showing the sources and distributions of autonomic and peptide-containing sensory nerve fibres in the heart in an amphibian (Bufo marinus) and a mammal (guinea-pig). Known or putative neurotransmitters are listed for each main class of neurons. ACh, acetylcholine; Adr, adrenaline; CGRP, calcitonin gene-related peptide; DRG, dorsal root ganglia; Dyn, dynorphins; Enk, enkephalins; Gal, galanin; NA, noradrenaline; NKA, neurokinin A; NPY, neuropeptide Y; SP, substance P; SST-14, somatostatin 14; SST-28, somatostatin 28; VIP, vasoactive intestinal polypeptide. Adapted from Morris, 1989.

Cranial autonomic preganglionic

Cranial autonomic preganglionic

Sensory (vagal+DRG) CGRP SP NKA

Toad

SP (CGRP) (NPY)

Sensoi 206 COMPARATIVE PHYSIOLOGY AND EVOLUTION

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atria, and which extends outside the heart near the origins of the aorta and the pulmonary vessels (Gabella, 1976). In some species, cardiac ganglia are also found in the walls of the ventricles (Davies, Francis and King, 1952; Smith, 1971). Aggregations of vagal postganglionic neurons in different locations within the cardiac plexus appear to innervate discrete regions of the heart (Randall et al., 1986, 1987; Pardini et al., 1987). Recent immunohistochemical studies have found a variety of neuropeptides within intracardiac neuronal cell bodies. These include somatostatin, NPY, VIP, substance P and calcitonin gene-related peptide (Weihe and Reinecke, 1981; Weihe, Reinecke and Forssmann, 1984; Hassall and Burnstock, 1984, 1987; Day et al., 1985; Dalsgaard et al., 1986; Franco-Cereceda, Lundberg and Hokfelt, 1986; Gerstheimer and Metz, 1986; Baluk and Gabella, 1989; J. Morris and I. Gibbins, unpublished; Figure 6.3). These studies confirm the heterogeneity of vagal postganglionic neurons, and raise the possibility that there are intrinsic circuits involving several different populations of intracardiac vagal neurons (see Gibbins and Morris, 1991). Postganglionic vagal neurons project to the atria, the sinoatrial node, the atrioventricular node and the conducting tissue in all mammals, where they mediate negative inotropic, chronotropic and dromotropic actions. The cardioinhibitory actions of the vagus have been attributed to acetylcholine acting on specialized populations of muscarinic receptors located close to the vagal postganglionic nerve terminals (Campbell et al., 1989). Although many of the peptides contained in vagal neurons have acute cardiac effects (see Morris, 1989), as yet there have been no studies performed that have demonstrated roles for neuropeptides in vagal neurotransmission to the mammalian heart. A vagal innervation of the ventricles has been found in some mammals. However, in most species the negative inotropic action of the vagus on the ventricles is only apparent at times of sympathetic excitation, and not in the resting heart (Levy, 1971, 1991). A marked exception to this pattern has been found in the ventricles of a hibernating mammal, the bat Miniopterus schreibersii (O’Shea and Evans, 1985). In this species, unilateral stimulation of the vagus nerve produces up to 90% inhibition of the force of contraction in the paced ventricle, even in the absence of sympathetic drive. Indeed, application of single current pulses to the vagus can produce significant reductions in the force of ventricular contraction. The ventricle of the bat is also inhibited by exogenous acetylcholine, and the atria and ventricles have similar sensitivities to the negative inotropic action of acetylcholine (O’Shea and Evans, 1985). The potency of exogeous acetylcholine as a cardioinhibitor is maintained at low temperatures when the basal heart rate is reduced, conditions that occur during hibernation (O’Shea, 1987, cf. Chatfield and Lyman, 1950; Lyman and O’Brien, 1963). The prominence of the vagal innervation of the ventricles in the bat, compared with other mammals, is likely to be related to the extreme cardiovascular requirements of initiating and terminating periods of hibernation. Furthermore, the vagal innervation may help to prevent ventricular arrhythmias during prolonged periods of low body temperature (O’Shea and Evans, 1985).

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Innervation by spinal autonomic nerves The cell bodies of catecholamine-synthesizing neurons innervating the mammalian heart are located in the stellate ganglia, the superior or middle cervical ganglia and the mediastinal ganglia (Nielsen, Owman and Santini, 1969; Gabella, 1976; Armour and Hopkins, 1984; Hopkins and Armour, 1984). The nerve fibres travel to the heart either in the cardiac nerves or by joining with the vagal nerve trunks. Spinal autonomic neurons innervate all of the chambers of the heart, although there is considerable species variation in the extent of the innervation of the ventricles (Dahlstrom et al., 1965; Nielsen and Owman, 1968; O’Shea and Evans, 1985). Nerve fibres with catecholamine fluorescence are also located among the intracardiac ganglion cells (Dahlstrom et al., 1965). In most mammalian species the catecholamine-synthesizing neurons innervating the heart contain NPY (Lundberg et al., 1983; Gu et al., 1983, 1984; Sternini and Brecha, 1985; Dalsgaard et al., 1986; Morris et al., 1986b; Wharton et al., 1988). An exception is found in some species of Australian marsupials, whose thoracic sympathetic ganglia lack NPY (Morris, Gibbins and Murphy, 1986). Instead, many spinal autonomic neurons in these species contain the peptide galanin (Morris, Gibbins and Holmgren, 1992). Neurons in the sympathetic chain ganglia of cats and dogs, which must include the cardiac neurons, contain both galanin and NPY (Kummer, 1987; Lindh, Lundberg and Hokfelt, 1989; Gibbins, unpublished). In guinea-pigs, the cardiac neurons containing noradrenaline and NPY lack galanin, but contain low levels of dynorphin-related opioid peptides (Lang et al., 1983; Weihe et al., 1985; Kummer et al., 1988). The spinal autonomic neurons mediate excitation of the mammalian heart, mainly via noradrenaline acting on ^-adrenoceptors in the region of the pacemaker and the myocardium (Hedberg, Minneman and Molinoff, 1980). However, NPY, which is released from cardiac neurons on electrical stimulation (Rudehill et al., 1986; Haas et al., 1989), may mediate non-noradrenergic excitatory effects on the heart of the pig, seen after prolonged stimulation of the stellate ganglion (Rudehill et al., 1986). Furthermore, stimulation of the spinal autonomic neurons supplying the heart results in a long-lasting, non-noradrenergic inhibition of the cardiac vagal nerves. This effect, which seems to be due to inhibition of the release of transmitter from the vagal nerves (Lundberg, Hua and Franco-Cereceda, 1984), may be mediated by NPY (Potter, 1985, 1987a, b; Warner and Levy, 1989) and/or opioid peptides (Koyanagawa et al., 1989) released from noradrenergic neurons.

AUTONOMIC INNERVATION OF THE VASCULATURE The vasculature of all vertebrates commonly receives a vasoconstrictor innervation by neurons whose cell bodies lie in spinal autonomic ganglia. These neurons characteristically synthesize catecholamines, and most also synthesize one or more neuropeptides. Catecholamine-synthesizing neurons may produce vasodilatation, in addition to, or instead of, vasoconstriction. Neurons with cell bodies in cranial or

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spinal autonomic ganglia, which do not synthesize catecholamines, but which synthesize acetylcholine and/or neuropeptides, generally have a more restricted distribution in the vasculature. Many of these neurons produce vasodilatation, while some produce vasoconstriction. Microganglia containing neurons with catecholamine fluorescence, or with acetylcholinesterase activity, are commonly found associated with large systemic vessels, and appear to contribute to their innervation (see Gibbins et al.9 1988). The catecholamine-synthesizing neurons in these ganglia are likely to be remnants of prevertebral spinal autonomic ganglia. However, it is not known whether the noncatecholamine-synthesizing neurons are in cranial or spinal autonomic pathways. There is good evidence in the cat and the guinea-pig intestinal mucosa that enteric neurons innervate submucosal arterioles. They mediate vasodilatation in response to local irritation of the mucosa (see Furness and Costa, 1987). Similar enteric reflexes may well occur in other vertebrates, but so far they have not been demonstrated. In addition to these autonomic neurons, processes of sensory neurons of small diameter often form a dense plexus around blood vessels. Antidromic stimulation of these sensory nerve endings can release transmitters, particularly the neuropeptides substance P and related tachykinins, and calcitonin gene-related peptide, which are often potent vasodilators. In many studies it is difficult to distinguish between the presence or actions of these sensory nerves, and those of autonomic nerves. This has lead to some uncertainty about the reported presence of an autonomic vasodilator innervation of some parts of the vasculature. CYCLOSTOME VASCULATURE Spinal autonomic nerve fibres innervate blood vessels in lampreys and histochemical studies have demonstrated that this innervation contains adrenergic elements (Tretjakoff, 1927; Johnels, 1956; Leont’eva, 1966; Govyrin, 1977). Both catecholamines and acetylcholine produce an increase of systemic and branchial vascular resistance in Myxine (Reite, 1969; Axelsson, Farrell and Nilsson, 1990), but little is known about the origin and nature of vasomotor nerves in the cyclostomes. There is no evidence for vasomotor innervation of the branchial vasculature of cyclostomes.

ELASMOBRANCH VASCULATURE Innervation by cranial autonomic nerves There is no conclusive evidence for an innervation of the branchial vasculature in elasmobranchs via the cranial nerves (Nicol, 1952). However, the flow of blood in the gills can be altered during electrical stimulation of the branchial nerves due to contractions of the striated muscle of the gill arch (Metcalfe and Butler, 1984).

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Innervation by spinal autonomic nerves Spinal autonomic pathways do not reach the branchial nerves in elasmobranchs, and there is no evidence for branchial vasomotor innervation in these fish (Nilsson, 1983). Early attempts to induce vasomotor responses with adrenaline in the branchial vasculature of the spiny dogfish, Squalus acanthias, were unsuccessful (Ôstlund and Fange, 1962). Later studies have, however, revealed a branchial vasodilation in response to this drug in Scyliorhinus canicula, and the concentration of catecholamines in blood plasma from Scyliorhinus was high enough to affect the branchial vascular resistance (Davies and Rankin, 1973). An adrenergic control via circulating catecholamines is therefore possible, compensating for the lack of an adrenergic innervation of the gills (Satchell, 1971; Gannon, Campbell and Satchell, 1972; Nilsson, 1984). There are several studies that demonstrate neuropeptides in nerves associated with blood vessels in elasmobranch fishes. However, the origin of these fibres remains unclear, and conclusions about the possible localization of the cell bodies of the paravertebral ganglia must await further analysis. Moderately dense plexuses of VIP- and somatostatin-IR fibres were demonstrated in the coeliac and mesenteric arteries of Squalus acanthias (Holmgren and Nilsson, 1983a). In contrast to the common vasodilator effect of VIP, an injection of VIP into unanaesthetized Squalus in vivo increased the vascular resistance of the gut, thus reducing the flow of blood in the gut (Holmgren, Axelsson and Farrell, 1992). Bombesin-IR nerve fibres provide a sparse innervation of the coronary arteries of the skate, Raja rhina, and in vessels in the gut of several species of elasmobranch (Tagliafierro et al., 1988; Bjenning, Driedzic and Holmgren, 1989; Bjenning, Jônsson and Holmgren, 1990). Bombesin contracts coronary arteries and generally increases the systemic vascular résistance (Bjenning, Jônsson and Holmgren, 1990; Bjenning, Farrell and Holmgren, 1991; S. Holmgren, M. Axelsson and A.P. Farrell, unpublished). NPY occurs in perivascular nerves in several organs, including branchial arteries, in skates (Raja spp.) (Bjenning, Driedzic and Holmgren, 1989). Injections of the native peptide into Squalus caused a dose-dependent increase in the arterial blood pressure (Conlon, Balasubramaniam and Hazon, 1991). It is possible that NPY coexists with adrenaline/noradrenaline as in other vertebrates (Gibbins et al., 1988; Morris, 1989), but unequivocal evidence is lacking. FIGURE 6.4 Diagram showing innervation of the branchial vasculature in a teleost fish. Blood flows from the afferent branchial artery (ABA) via the afferent filamental artery (AFA) and lamellar arterioles (ALa) into the lamellae, where exchange of gases takes place. The oxygenated blood passes through the efferent lamellar arterioles (ELa) and efferent filamental artery (EFA) to the efferent branchial artery (EBA). Spinal autonomic pathways with postganglionic adrenergic nerve fibres (Adr; broken line) innervate both afferent and efferent effectors, and run to the central vascular sinus (CVS). Cranial autonomic, vagal pathways include both cholinergic (ACh, solid line) and serotonergic (5-hydroxytryptamine containing; 5-HT, dash/dot line) postganglionic neurons. Cholinergic fibres are found chiefly in the sphincter at the base of the efferent filamental artery (Sph), and this sphincter also receives substantial innervation by serotonergic and adrenergic fibres. In addition, serotonergic fibres are found in most other efferent vascular elements and in the central venous sinus. Other abbreviations: FC, filamental cartilage; CNS, central nervous system.

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figure 6.4

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TELEOST VASCULATURE Innervation by cranial autonomic nerves Cranial autonomic (vagal) nerves carry vasomotor fibres to the gills of teleosts. Ultrastructural studies of the complex vasculature of the gills of teleosts reveal cholinergic-type nerve profiles innervating the sphincter at the base of the efferent filamental arteries, and histochemical studies demonstrate high levels of cholinesterase in this region (Bailly and Dunel-Erb, 1986; Figure 6.4). Physiological studies corroborate the structural findings, showing a functional cholinergic vasoconstrictor control of the filamental sphincters (Smith, 1977, 1978a; Pettersson and Nilsson, 1979; Nilsson and Pettersson, 1981; Nilsson, 1984). In addition, serotonergic (5-hydroxytryptamine-containing) nerves of vagal origin occur in the efferent vasculature of the gills of teleosts (Dunel-Erb, Bailly and Laurent, 1982, 1989; Bailly et al., 1989; Figure 6.4). Innervation by spinal autonomic nerves The sympathetic chains of teleosts and other actinopterygian fish continue into the head bearing ganglia in contact with the cranial nerves (see Nilsson, 1983; Gibbins, Chapter 1, this volume). Grey rami communicantes enter the spinal and cranial nerves, and the vagus may thus be regarded as a “vagosympathetic trunk” of the type described for amphibians and reptiles. There is ample evidence that the spinal autonomic system is the origin for an adrenergic vasomotor innervation of the branchial and systemic vascular beds in teleosts and other actinopterygians. Adrenergic fibres join the branchial nerves and run to the vasculature of the gill, where fibres run to both afferent and efferent arteries, to the nutritive vasculature of the gill and to the central venous sinus (for references, see Nilsson and Holmgren, 1992; Figure 6.4). Spinal autonomic pathways also run in the anterior splanchnic nerve to the anterior viscera, and in the posterior splanchnic nerve (“vesicular nerve”) to the posterior viscera, notably to the urinogenital organs where blood vessels receive an adrenergic innervation (Nilsson, 1970, 1973, 1976; Uematsu, Holmgren and Nilsson, 1989). In contrast to elasmobranchs, the spinal nerves of teleost fish possess grey rami communicantes and spinal autonomic pathways may thus join the spinal nerves and run to the somatic vasculature. There are several studies that conclude that adrenergic nerves are involved in the regulation of blood pressure in teleosts both at rest and during various stimuli, such as exercise and hypoxia (Wood and Shelton, 1975; Smith, 1978a; Axelsson and Nilsson, 1986; Axelsson, 1988; Axelsson et al., 1989b; Fritsche and Nilsson, 1989, 1990, 1992). The presence of neuropeptides in vasomotor nerves in teleost fish is well established. A sparse innervation by VIP-IR nerves has been demonstrated in small blood vessels of the swimbladder, the urinary bladder, the gonads and the gallbladder of the cod (Lundin and Holmgren, 1984, 1986, 1989; Aldman and Holmgren, 1987; Uematsu, Holmgren and Nilsson, 1989), and there is also a sparse innervation by such fibres of blood vessels in the gut of the holostean, Lepisosteus platyrhincus,

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(Holmgren and Nilsson, 1983b), the cod (Jensen and Holmgren, 1985) and the rainbow trout (Holmgren, Vaillant and Dimaline, 1982). The vascular resistance of the perfused swimbladder and gas gland of the cod decreased in response to porcine VIP (Lundin and Holmgren, 1984), and the arterioarterial branchial vascular resistance of the brown trout, Salmo trutta, was decreased by VIP in a dosedependent manner (Bolis et al., 1984). Injections of porcine VIP into cod in vivo caused an increase in the flow of blood in the coeliac and mesenteric arteries, as well as an increased cardiac stroke volume. The increase in flow in the mesenteric artery simply reflected the increase in cardiac output, while in the coeliac artery there was an additional decrease of vascular resistance (Jensen, Axelsson and Holmgren, 1991). The difference in the responses to VIP between the two vascular beds is consistent with the observation of a more dense VIP innervation of the coeliac artery than the mesenteric artery (S. Holmgren, unpublished). Innervation of the blood vessels of teleosts by fibres containing substance P is sparse, although endocrine cells of the gut have been shown to store tachykinin-like peptides (Jensen, 1989; Jensen and Holmgren, 1992). Substance P causes vasodilation in the cod and injection in vivo produces complex vascular responses that include a cholinergic component (Jensen, Axelsson and Holmgren, 1991). In the cod, galanin-like IR occurs in perivascular nerves of arterial branches to the gut, and galanin has excitatory effects on isolated arterial preparations from the cod (P. Karila and S. Holmgren, unpublished). Preliminary experiments with the cod in vivo, demonstrate a reduction in the flow of blood in the gut after injection of sulphated cholecystokinin 8 (CCK8-S) and caerulein (J. Gunnarsson, S. Holmgren and S. Nilsson, unpublished), and there is a dramatic elevation in the ventral aortic blood pressure in response to these peptides (J. Jensen, M. Axelsson and S. Holmgren, unpublished). Part of the general cardiovascular effect of CCK8-S is explained by the marked vasoconstriction of the branchial vasculature (Sundin and Nilsson, 1992). Gastrin/CCK-IR neurons have also been demonstrated in the branchial vasculature of the cod, Gadus morhua, and both CCK8-S and caerulein produce strong vasoconstrictor effects on the branchial vasculature. However, the origin of these fibres is not clear (Sundin and Nilsson, 1992; S. Holmgren, unpublished). AMPHIBIAN VASCULATURE Innervation by cranial autonomic nerves A prominent innervation by cranial autonomic neurons has been demonstrated both morphologically and physiologically in only two vascular beds in anuran amphibians: the lingual and the pulmonary vascular beds (Figure 6.5). Postganglionic neurons lying in glossopharyngeal pathways contain IR to VIP and galanin, and innervate the vasculature of the tongue (Baecker, Yanaihara and Forssmann, 1983; Cousins and Campbell, 1988; Morris, Gibbins and Osborne, 1989). These neurons produce dilatation of the lingual vasculature, which, in the

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frog retrolingual membrane seems to be cholinergic (Berman and Siggins, 1968), and in the toad appears to be mostly non-cholinergic (Cousins and Campbell, 1988). It is possible that VIP or galanin are involved in vasodilatation in the tongue of the toad, but this has not yet been specifically tested. Postganglionic vagal neurons innervating the pulmonary artery of the toad contain somatostatin- and galanin-IR, and their cell bodies lie along the vagosympathetic nerve trunk (Gibbins et al., 1987; Morris, Gibbins and Osborne, 1989). The terminals of these neurons form a circularly oriented band around the pulmonary artery just distal to the pulmocutaneous artery (Morris et al., 1986a; Morris, Gibbins and Osborne, 1989; Figure 6.5). This corresponds to the position in the pulmonary artery of the frog where there is evidence for an anatomical sphincter (de Saint-Aubain and Wingstrand, 1979). These vagal neurons, which are tonically active in urodeles (Luckhardt and Carlson, 1921), provide a strong constrictor innervation of the pulmonary artery (Campbell, 1971). Indeed, stimulation of vagal neurons supplying the amphibian lung, either electrically, or reflexly during hypercapnia, can lead to an almost complete occlusion of the pulmonary artery (Couvreur, 1889; Smith, 1978b: Johansen, 1982). This vagally mediated vasoconstriction seems to be mostly cholinergic, although a slow neurogenic constriction can be revealed after the destruction of sensory neurons with capsaicin and cholinergic blockade (Davies and Campbell, 1988). This slow constriction is mimicked by somatostatin (Davies and Campbell, 1988), which could be the endogenous mediator of the response. In addition to this major excitatory innervation of the pulmonary artery of the toad, there is a much sparser innervation by VIP-IR neurons whose cell bodies lie in the distal vagal nerve trunk. These neurons are probably responsible for the rapid, non-adrenergic, non-cholinergic dilatation seen after transmural stimulation of the pulmonary artery (Davies, 1989). The cerebral arteries of some amphibians are supplied with a plexus of nerve fibres showing acetylcholinesterase activity (Tagawa et al., 1979b, 1989). By comparison with mammals (see below), the acetylcholinesterase-positive neurons innervating the extraparenchymal blood vessels may well be processes of cranial autonomic neurons that synthesize acetylcholine, while the nerve fibres innervating small blood vessels in the brain parenchyma are thought to originate from central cholinergic neurons (Tagawa et al., 1979b). However, in addition to occurring in cholinergic neurons, acetylcholinesterase activity has been demonstrated in catecholamine-synthesizing neurons and in sensory neurons (Eranko et al., 1970; Furness, 1973; Barajas and Wang, 1975; Papka, 1978; Papka et al., 1981). FIGURE 6.5 Diagram showing the autonomic innervation of the vasculature in an amphibian (Bufo marinus). The density of stippling represents the density of nerve fibres revealed by catecholamine fluorescence procedures or by immunohistochemical demonstration of neuropeptides. Note that the innervation by cranial autonomic neurons (upper left inset) is much more restricted than the spinal autonomic innervation. Abbreviations: ba, brachial artery; cuta, cutaneous artery; cma, coeliacomesenteric artery; da, dorsal aorta; eca, external carotid artery; ia, ileal artery; ica, internal carotid artery; lia, lingual artery; ma, mesenteric artery; pa, pulmonary artery; pea, pulmo-cutaneous artery; ra, renal artery.

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Therefore without a direct demonstration of the origin of the acetylcholinesterasepositive nerve fibres, we cannot be certain that the cerebral blood vessels of amphibians are innervated by cranial autonomic neurons. Innervation by spinal autonomic nerves Most arteries and veins in anuran amphibians are innervated by spinal autonomic neurons that synthesize catecholamines (Banister and Mann, 1966: McLean and Burnstock, 1966; Kirby and Burnstock, 1969a; Leont’eva, 1978; Nilsson, 1978; Morris and Gibbins, 1983; Figure 6.5). The arterial innervation is generally more dense than that of veins, but appears to be less dense than the vascular innervation in reptiles and mammals. In B. marinus and R. catesbeiana, NPY coexists with adrenaline in vascular neurons (Morris et al., 1986a; Horn, Stofer and Fatherazi, 1987), except those innervating the coronary vasculature (Morris et al., 1986a). Furthermore, in toads, many vascular neurons with NPY-IR also contain galanin-IR (Morris, Gibbins and Osborne, 1989). In most vessels, stimulation of the spinal autonomic neurons produces constriction. The vasoconstriction is abolished by drugs that block adrenergic neurons, such as bretylium, and is reduced or abolished by a-adrenoceptor antagonists (Kirby and Burnstock, 1969a; Smith, 1976; Nilsson, 1978; Morris, 1983; Cousins and Campbell, 1988; Stofer, Fatherazi and Horn, 1990; Osborne, 1992). The adrenoceptors that mediate the neurogenic constriction of the renal vasculature and aorta of the toad do not appear to be similar to the adrenoceptors that mediate constrictions by exogenous adrenaline, and do not easily fit into the classification of a r or a 2-adrenoceptors (Morris, 1983). Furthermore, there is evidence that a slower, non-adrenergic constriction occurs in addition to the a-adrenoceptor-mediated constriction in the vasculature of the skeletal muscle in the bullfrog (Stofer, Fatherazi and Horn, 1990), and in the mesenteric artery of the toad (Osborne, 1992). The slow constriction of the vasculature of the skeletal muscle in the bullfrog is mimicked by exogenous NPY (Stofer, Fatherazi and Horn, 1990), and thus may be produced by NPY released from the perivascular nerve terminals. NPY and galanin are both potent pressor agents in B. marinus (Courtice, 1991), but as yet, experiments have not been conducted to demonstrate roles for these peptides in transmission from spinal autonomic neurons to the vasculature. In some parts of the vasculature, adrenaline released from neurons of spinal origin may produce vasodilatation. In the pulmonary vascular bed of the toad, adrenergic dilatation is the predominant response to stimulation of the spinal autonomic nerves (Campbell, 1971). However, as strips of the main pulmonary artery are contracted by adrenaline, the neurogenic dilatation of the pulmonary vasculature may be produced preferentially in smaller arterial branches (Holmgren and Campbell, 1978). There is also evidence that neuronally released adrenaline has access to /3-adrenoceptors that mediate dilatation in the renal vasculature of the toad, even though the predominant adrenergic response in this vascular bed is constriction (Morris, 1983).

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REPTILIAN VASCULATURE Innervation by cranial autonomic nerves As in amphibians, innervation of the reptilian vasculature by cranial autonomic neurons is most prominent in the lingual and the pulmonary vascular beds. In lizards, the neuronal cell bodies that lie along intralingual branches of the facial nerve contain VIP-IR (see Gibbins, chapter 1). These neurons are the likely source of the dense plexus of VIP-IR nerve fibres associated with the lingual arteries (Baecker, Yanaihara and Forssmann, 1983; I. Gibbins, unpublished). These neurons presumably correspond to the VIP-IR glossopharyngeal neurons in the amphibian tongue, although functional experiments have not been conducted in reptiles to determine whether they produce vasodilatation. Postganglionic neurons in vagal pathways to the lung of two species of snake contain VIP-IR (Donald and Lillywhite, 1989a; Donald, O’Shea and Lillywhite, 1990b). These neurons innervate both the extrinsic and the intrinsic segments of the pulmonary vasculature. Physiological studies have demonstrated strong vagal constriction of the pulmonary vasculature in turtles, lizards and snakes, which is located mainly in the extrinsic pulmonary artery (Berger, 1972, 1973; Burggren, 1977; Milsom, Languille and Jones, 1977; Berger and Burnstock, 1979) or in the pulmonary outflow tract of the truncus arteriosus (Smith and MacIntyre, 1979). However, in the ratsnake Elaphe obsoleta, activation of the vagus can also mediate constriction of the intrapulmonary vasculature (Donald, O’Shea and Lillywhite, 1990b). All of these pulmonary vasconstrictions seem to be cholinergic (Berger, 1972, 1973; Burggren, 1977; Milsom, Languille and Jones, 1977; Berger and Burnstock, 1979; Smith and MacIntyre, 1979; Donald, O’Shea and Lillywhite, 1990b). The coronary vasculature of the tortoise also receives a vagal innervation (Drury and Smith, 1924). Stimulation of the vagus produces a cholinergic dilatation of the coronary vessels (Sumbal, 1924). Cerebral vessels of snakes and turtles are innervated by nerve fibres with acetylcholinesterase activity (Iijima, 1977; Iijima et al.9 1977). However, as in amphibians, it is not known whether these fibres are processes of cranial autonomic neurons, or other classes of neurons. Innervation by spinal autonomic nerves The innervation of the vasculature by spinal autonomic neurons that synthesize catecholamines has been well described for several reptilian species. Histochemical studies have demonstrated an innervation of most arteries and many veins by catecholamine-synthesizing neurons (McLean and Burnstock, 1966; Furness and Moore, 1970; Iijima, 1977; Iijima et al., 1977; Donald and Lillywhite, 1988, 1989b; J. Morris, S. Holmgren and I. Gibbins, unpublished; Figures 6.6-6.9). The predominant catecholamine synthesized by these neurons is noradrenaline. The neurons may also contain one or more neuropeptides, particularly NPY or galanin (Davies, Donald and Campbell, 1990; I. Gibbins, J. Morris, S. Holmgren, unpublished). In addition to this innervation by catecholamine-synthesizing autonomic neurons, small, brightly fluorescent cells (extra-adrenal chromaffin cells) are

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FIGURE 6.6- 6.9 Whole-mounts of lizard blood vessels showing innervation by spinal autonomic neurons that synthesize catecholoamines. Glyoxylic acid-induced fluorescence. 6: Systemic arch from the bearded dragon (Pogona vitticeps). 7: Mesenteric artery of the bearded dragon. 8: Small artery supplying the body wall of a goanna (Varanus gouldii). Note the large number of chromaffin cells scattered along the artery. 9: Inferior vena cava of the bearded dragon. Note the clumps of chromaffin cells. Scale bar = 100 ^m for all figures. Figures courtesy of J. Morris, S. Holmgren and I. Gibbins.

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abundant in the adventitia of large blood vessels in some reptiles (Furness and Moore, 1970; Berger et al., 1982; Donald, O’Shea and Lillywhite, 1990b; J. Morris, S. Holmgren and I. Gibbins, unpublished; Figures 6.8 and 6.9). The vessels associated with the gastrointestinal tract in all of the lizards and snakes examined receive a dense innervation (Figure 6.7). However, the density of the innervation of other vascular beds by catecholamine-synthesizing neurons can vary greatly. For example, the common carotid artery and the pulmonary artery in the semiarboreal ratsnake Elaphe obsoleta are only sparsely innervated (Donald and Lillywhite, 1988), compared with the moderate to dense innervation of the same arteries in several species of lizard (Trachydosaurus rugosus: Furness and Moore, 1970; Varanus gouldii; Pogona vitticeps). Furthermore, the posterior arteries and veins in the ratsnake receive a particularly dense innervation by catecholaminesynthesizing neurons (Donald and Lillywhite, 1988). The predominant effect on the systemic vasculature of stimulating spinal autonomic neurons that synthesize catecholamines is constriction mediated primarily by a-adrenoceptors. Thus, the marked craniocaudal gradient in the density of innervation of the systemic vasculature by catecholamine-synthesizing neurons in the semi-arboreal ratsnake can be explained by the need to produce strong constriction of the posterior arteries and veins, in order to maintain blood pressure and to regulate the flow of blood during a vertical posture (Seymour and Lillywhite, 1976; Lillywhite and Seymour, 1978; Lillywhite and Gallagher, 1985; Lillywhite, 1987; Donald and Lillywhite, 1988). The predominant effect of stimulating the spinal autonomic neurons on the pulmonary vasculature of lizards and snakes is dilatation mediated by /3-adrenoceptors (Berger, 1972, 1973; Smith and MacIntyre, 1979; Lillywhite and Donald, 1989; Donald, O’Shea and Lillywhite, 1990b). Pulmonary vasoconstriction mediated by a-adrenoceptors may also occur (Berger, 1973). However, the pulmonary vasculature of turtles seems to lack an innervation from spinal autonomic neurons (Burggren, 1977; Milsom, Languille and Jones, 1977). AVIAN VASCULATURE Innervation by cranial autonomic nerves There have been very few studies of the innervation of the avian vasculature by cranial autonomic neurons. However, scattered observations of the tongue, brain, lung and gastrointestinal tract indicate that blood vessels in these regions may be innervated by cranial autonomic neurons. Nerve fibres with VIP-IR innervate blood vessels in the tongue of the quail (Baecker, Yanaihara and Forssmann, 1983). These fibres are likely to originate from VIP-IR neuronal cell bodies in ganglia located in the posterior region of the tongue, which presumably lie in facial nerve pathways (see Gibbins, chapter 1). As in other vertebrates, nerve fibres with acetylcholinesterase activity supply cerebral blood vessels in birds (Tagawa, Ando and Wasano, 1979). These nerves have a different distribution from the catecholamine-containing neurons, and may represent a

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cranial autonomic innervation. There are reports from functional studies that the vagus also innervates the vasculature of the crop and oesophagus (Couvreur, 1892, cited by Bennett, 1974). Innervation by spinal autonomic nerves The vasculature of birds may be innervated by two main classes of neurons whose cell bodies lie in spinal autonomic ganglia. The most prominent innervation is by catecholamine-synthesizing neurons. These neurons innervate most arteries in the fowl (Bennett and Malmfors, 1970; Bennett, 1971), with the cerebral arteries and cutaneous arteriovenous anastomoses receiving a particularly dense innervation (Tagawa, Ando and Wasano, 1979; Midtg&rd, 1988). Catecholamine-synthesizing neurons also provide a dense innervation of the veins, especially the proximal pulmonary vein, the inferior vena cava and the renal portal vein (Bennett and Malmfors, 1970; Bennett, 1971; Bennett, Cobb and Malmfors, 1974). Many catecholamine-synthesizing neurons supplying the vasculature of the fowl contain NPY-IR (Gibbins et al.9 1988). In most avian blood vessels, stimulation of the catecholamine-synthesizing neurons produces vasoconstriction mediated by a-adrenoceptors (Folkow, Fuxe and Sonnenschem, 1966; Bennett and Malmfors, 1975; McGregor, 1979; Midtgard and Bech, 1981; Hillman, Scott and von Tienhoven, 1982). /3-Adrenoceptors may be present in some vessels, although not dominant (Somlyo and Woo, 1967; Bolton and Bowman, 1969). An exception is the longitudinal muscle layer of the anterior mesenteric artery, in which neurally released catecholamines mediate vasodilatation via /3-adrenoceptors (Bolton, 1968; Bell, 1969). One of the most striking examples of vasoconstriction mediated by catecholamine-synthesizing neurons is the constriction of the extramuscular arteries supplying duck hindlimbs during diving (Folkow, Fuxe and Sonnenschem, 1966). The femoral artery in ducks is much more heavily innervated than the corresponding artery in a non-diving bird, the turkey, or in the cat (Folkow, Fuxe and Sonnenschem, 1966). The extramuscular location of this site of vasoconstriction may be responsible for the ability of birds to maintain a reduced flow of blood to the skeletal muscles during prolonged periods of diving, despite concomitant muscular activity (Folkow, Fuxe and Sonnenschem, 1966). There are several reports of neurogenic vasodilatation in the limbs of birds, which seems to be at least partly mediated by acetylcholine (Feigl and Folkow, 1963; Johansen and Millard, 1974). These responses are likely to be mediated by neurons whose cell bodies lie in spinal autonomic ganglia. Furthermore, it is possible that neurons with VIP-IR innervating arteriovenous anastomoses in the skin of fowl (Midtg&rd, 1988) or the systemic blood vessels of fowl (Gibbins et al., 1988), or acetylcholinesterase-positive nerve fibres in the feet of ducks (Molyneux and Harmon, 1982), might be in spinal autonomic pathways. It is not clear whether the noncholinergic vasodilatation in the hindlimbs of birds (Johansen and Millard, 1974; McGregor, 1979) is due to the activation of spinal autonomic neurons or to antidromic activation of sensory nerve fibres. The longitudinal muscle layer of the anterior mesenteric artery of fowl (Bolton,

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1968; Bell, 1969), and the inferior vena cava of fowl (Bennett and Malmfors, 1975), are innervated by cholinergic constrictor neurons. Again, although the origin of these nerve fibres has not been established, it is possible that they lie in spinal autonomic pathways.

MAMMALIAN VASCULATURE Innervation by cranial autonomic nerves Cranial nerves in facial, glossopharyngeal and vagal nerve pathways innervate many blood vessels in the head, thorax and the upper gastrointestinal tract of mammals. The postganglionic neurons innervating the cephalic vasculature have their cell bodies in a range of ganglia in the head: sphenopalatine ganglion; otic ganglion; submandibular ganglion; sublingual ganglia; and intracranial ganglia associated with the facial or glossopharyngeal nerves. Neurons innervating the thoracic and abdominal vessels have their cell bodies in local ganglia. It is assumed that all of these postganglionic neurons synthesize acetylcholine, and most of them also contain one or more neuropeptides. VIP is the most widespread peptide in vascular neurons in cranial autonomic pathways (Lundberg, 1981; Gibbins, Brayden and Bevan, 1984; Gibbins, 1990), and usually coexists with its gene-related product, peptide histidine isoleucine (Lundberg et al., 1984). Other peptides, such as NPY, may be present as well as VIP and peptide histidine isoleucine (Leblanc, Trimmer and Landis, 1987; Gibbins and Morris, 1988; Sheikh et al., 1988). Almost invariably, these cranial autonomic neurons produce vasodilatation. One exception seems to be the coronary vasculature of some mammalian species, where stimulation of cranial autonomic neurons produces vasoconstriction (Kalsner, 1985; Young, Knight and Vatner, 1987, 1988). Vasodilatation in many parts of the head and airways is associated with glandular secretion, and usually involves noncholinergic as well as cholinergic mechanisms (Heidenhain, 1872; Bevan et al., 1982). A fast, non-cholinergic vasodilatation is apparent in some cerebral arteries. Recent evidence suggests that nitric oxide is involved in these responses, possibly as a neurotransmitter (Toda, Ayajiki and Okamura, 1990). More long-lasting noncholinergic vasodilatations are seen in many parts of the cephalic vasculature, and in the gastric circulation, in response to high-frequency stimulation of cranial autonomic neurons. These non-cholinergic responses often occur together with a cholinergic vasodilatation. There is good evidence that VIP is the transmitter mediating these slow vasodilatations (Lundberg, Anggard and Fahrenkrug, 1981; Bevan et al., 1984; Goadsby and MacDonald, 1985; Brayden and Bevan, 1986; Ito, Ohga and Ohta, 1988). Cephalic vasodilatation produced by stimulation of cranial autonomic neurons is likely to play an important role in thermoregulation, particularly in animals such as the cat, which pants to enhance evaporative water loss (Gibbins, Brayden and Bevan, 1984).

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Innervation by spinal autonomic nerves There are three main classes of spinal autonomic neurons that may innervate the mammalian vasculature: (1) catecholamine-synthesizing neurons whose cell bodies lie in the prevertebral or paravertebral sympathetic ganglia; (2) non-catecholaminesynthesizing neurons with cell bodies in the sympathetic ganglia; (3) neurons, most of which do not synthesize catecholamines, whose cell bodies are located in the pelvic ganglia. Spinal autonomic neurons that synthesize catecholamines innervate most vascular segments in mammals. The density of innervation can vary considerably between species, between different vascular beds and between different segments of the same vascular bed (Gibbins et al,, 1988). Contrary to popular belief, veins may be very densely innervated, some being even more densely innervated than the corresponding arteries. In general, the most densely innervated veins are the large systemic veins close to the heart (Gibbins et al., 1988 cf. Nilsson et al., 1988), the veins of the limbs and branches of the facial vein (Pegram, Bevan and Bevan, 1976). Vascular neurons that synthesize catecholamines in mammals generally contain NPY in addition to noradrenaline (Lundberg et al., 1983; Lundberg and Hokfelt, 1986). The exceptions to this generalization include most of the neurons innervating the cephalic and thoracic vasculature in some Australian marsupials (Morris, Gibbins and Murphy, 1986; Morris, Gibbins and Holmgren, 1992), neurons innervating venous sinusoids in the nasal mucosa of the cat (Lundberg and Hokfelt, 1986), and those neurons supplying the microvasculature in non-hairy skin of guinea-pigs (Gibbins and Morris, 1990). Instead of NPY, these neurons may contain galanin (Morris, Gibbins and Holmgren, 1992) or opioid peptides (Gibbins and Morris, 1990). Furthermore, galanin or opioid peptides coexist with noradrenaline and NPY in some vascular neurons (Fried et al., 1986; Kummer, 1987; Morris, Gibbins and Holmgren, 1992). In most vascular beds, the predominant response to stimulation of catecholaminesynthesizing neurons is vasoconstriction. Nerve-mediated contraction of vascular smooth muscle occurs in elastic arteries, small and large muscular arteries, precapillary arterioles, and medium to large veins. In many cases these constrictions appear to be mediated by noradrenaline acting on vascular a-adrenoceptors. However, constriction of some blood vessels in response to single nerve impulses, or to short trains of impulses, is often resistant to blockade of a-adrenoceptors. There is evidence that in some blood vessels these responses are mediated by neuronally released ATP (Burnstock and Kennedy, 1986), and in other vessels seem to be mediated by a population of specialized adrenoceptors located on the membrane of the smooth muscle, close to the sites of transmitter release (Hirst and Neild, 1981; see Morris and Gibbins, 1992). On the other hand, prolonged vasoconstrictions produced by long trains of nerve impulses, or by impulses delivered at high frequencies, may also be resistant to adrenoceptor antagonists. There is good evidence that NPY is the transmitter responsible for mediating at least some of these long-lasting vasoconstrictions (Lundberg and Hokfelt, 1986; Morris and Murphy, 1988; Ohlen et al., 1990; Morris, 1991).

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In a few specialized vascular beds, such as the coronary arterial bed of some species, the facial veins, and the precapillary arterioles in skeletal muscle beds, the predominant effect of stimulating catecholamine-synthesizing neurons, is vasodilatation mediated by /3-adrenoceptors (Pegram, Bevan and Bevan, 1976; Lundvall, Hillman and Gustafsson, 1982; Jackson, Pope and Lucchesi, 1987; Elkhawad, Al-Zaid and Bou-Resli, 1990). /3-Adrenoceptors are also present in many other vessels, but noradrenaline can only produce vasodilatation after blockade of the a-adrenoceptor-mediated vasoconstrictions. Most of the non-catecholamine-synthesizing neurons with cell bodies in the sympathetic ganglia seem to synthesize acetylcholine. In guinea-pigs, these neurons also contain the peptides VIP, NPY and dynorphin (Gibbins, 1992). It has been known for many years that these “cholinergic sympathetic” neurons innervate resistance vessels in the skeletal muscle of the limbs (Bulbring and Burn, 1935). However, in the hindlimbs of guinea-pigs, this innervation is restricted to the small arteries lying just proximal to the muscle, and does not extend to the intramuscular arterial segments (Gibbins, 1992). Stimulation of the non-catecholamine neurons results in vasodilatation of the vasculature of the skeletal muscle (Bulbring and Burn, 1935; Mellander and Johansson, 1968), which appears to be important in the “alerting response” at the start of exercise (Bell, 1975). Many parts of the vasculature in the pelvic region of mammals receive a dense innervation by autonomic neurons projecting from the pelvic ganglia. Usually this is in addition to the innervation by vasoconstrictor neurons with cell bodies in the sympathetic ganglia. Most of the pelvic vascular neurons do not synthesize catecholamines, although they may contain immunoreactivity to one of the catecholamine-synthesizing enzymes (Morris and Gibbins, 1987). Many of the pelvic vascular neurons synthesize acetylcholine, and most also contain one or more neuropeptides. The most widespread neuropeptides are VIP, NPY and dynorphin (Dail, Moll and Weber, 1983; Morris et al., 1985). Stimulation of the pelvic neurons produces strong vasodilatation, particularly of the blood vessels associated with the genital and reproductive organs (Bell, 1972). There is considerable variability in the proportion of the vasodilatation that is cholinergic between regions of the vasculature, between species and between reproductive states (Bell, 1968; Sjostrand and Klinge, 1979; cf. Creed, Carati and Keogh, 1988). There is evidence for both fast and slow non-cholinergic vasodilatation in the pelvic vasculature (Bowman and Gillespie, 1983; Morris and Murphy, 1988; Morris and Gibbins, 1992). There is increasing evidence that the fast non-cholinergic vasodilatation involves the generation of nitric oxide, which may act as a neurotransmitter (Bowman and Gillespie, 1983; cf. Martin et al., 1988; Ignarro et al., 1990; Holmquist, Hedlund and Andersson, 1991). Furthermore, there is strong evidence that VIP mediates the maintained increase in blood flow in the corpus cavernosum following stimulation of the pelvic nerve (Andersson, Bloom and Mellander, 1984; Andersson et al., 1987; Juenemann, etal., 1987). The relative roles of these different neurotransmitters have not yet been fully defined, however it appears that both acetylcholine and VIP are necessary for the full erectile response in the penis of the dog (Carati, Creed and Keogh, 1988).

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CARDIOVASCULAR REFLEXES The need to preserve a constant inner environment (homeostasis), i.e. a constant milieu for the cells, has induced the evolution of a number of nervous mechanisms that regulate the physical and chemical properties of the blood and other body fluids. For instance, to maintain an adequate perfusion of the organs by the blood, the blood pressure is regulated within certain limits that are adjusted to conform to the physiological situation of the animal, a situation that may vary considerably. Consider, for example, water-breathing fish at zero gravity in a relatively constant environment such as the sea, in comparison with running or flying birds and mammals. In addition, internal changes due to “external” stimuli (exercise, diving, hypoxia, to name a few examples) may occur within the individual organism. Histochemical and pharmacological experiments on isolated tissues and organs have allowed a certain degree of clarity regarding the nature of autonomic fibres, and their possible effects on their targets within the cardiovascular system. However, the actual effects, in vivo, are in most cases obscure in the non-mammalian vertebrates. For example, there is a clear gap in our knowledge regarding which autonomic pathways are responsible for a reflex-induced hypotension. Bradycardia has been the most widely documented effect, but in some cases (teleosts and mammals), there is clear additional evidence for inhibition of the activity of the vasomotor nerves during reflex hypotension. Studies of autonomic components in cardiovascular reflex pathways have been conducted on vertebrates representing all classes. The studies have largely been restricted to the “classical” cholinergic and adrenergic cardioregulatory and vasomotor mechanisms, where simple pharmacological experiments will yield information on patterns of autonomic innervation (Figure 6.10). Thus, information about non-adrenergic, non-cholinergic components in the reflex pathways is scarce in the non-mammalian vertebrates. BARORECEPTOR REFLEXES In mammals, the arterial blood pressure affects the diameter of the larger arteries (aorta and carotid arteries), which in turn causes stretching of sensory nerve endings (baroreceptors), within the arterial wall. The baroreceptors increase their firing rate in response to the elevated blood pressure, and the sensory information enters the central nervous system via the vagus (aortic baroreceptors) or glossopharyngeal nerves (carotid artery baroreceptors). In addition to the arterial baroreceptors, cardiac baroreceptors have also been demonstrated in a number of vertebrate groups (Jones and Milsom, 1982). The location of the arterial baroreceptors varies between the vertebrate groups, but the basic pattern of the cardioinhibitory reflex pathway is the same. The presence of autonomic pathways that regulate the arterial blood pressure via control of the vascular resistance is uncertain in many of the ectothermic groups, but such pathways are well established in mammals (e.g. Folkow and Neil, 1971).

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Baroreceptor input from Gills (elasmobranchs and teleosts) Pulmocutaneous artery (amphibians) Truncus/pulmonary artery (reptiles) Aorta (birds) Aorta/carotid arteries (mammals)

Chemoreceptor input from Gills (elasmobranchs and teleosts) Pulmocutaneous artery and carotid labyrinth (amphibians) Pulmonary artery (?) (reptiles) Carotids (reptiles, birds and mammals) FIGURE 6.10 Diagram summarizing two of the major autonomic pathways involved in circulatory control: the vagal, cholinergic cardioinhibitory pathway and the spinal autonomic vasomotor pathway. The diagram also outlines, for the different classes of vertebrates, the baro- and chemoreceptor input affecting these pathways.

In elasmobranchs, barosensitivity has been demonstrated in areas innervated by branchial nerves, but the role of the pathway in vivo is unclear (Ristori and Dessaux, 1970 a, b; Barrett and Taylor, 1984). Furthermore, a well-documented bradycardia induced by hypoxia and mediated by the vagus in dogfish (Satchell, 1961; Piiper, Baumgarten and Meyer, 1970) was compensated for by an increase in stroke volume due to a Frank-Starling relationship, rendering a constant cardiac output and thus having little effect on the blood pressure (Short, Butler and Taylor, 1977; Taylor, Short and Butler, 1977). In teleosts, several studies demonstrate that an increase in blood pressure induced by an injection of adrenaline, or by elevating the blood pressure in individual branchial arteries, produces bradycardia due to activation of the vagal inhibitory pathways (Randall and Stevens, 1967; Ristori, 1970; Ristori and Dessaux, 1970a, b; Stevens eta l.9 1972; Helgason and Nilsson, 1973; Pettersson and Nilsson, 1980; Wood and Shelton, 1980; Jones and Milsom, 1982; Bagshaw, 1985). Experiments with carp (Cyprinus carpio) suggest that the first two pairs of gill arches are particularly active as baroreceptor areas (Ristori and Dessaux, 1970a, b). In amphibians, baroreceptors that trigger a cardioinhibitory reflex reside in the carotid labyrinth and in the pulmocutaneous artery from which sensory fibres run in the laryngeal nerve (Ishii and Oosaki, 1969; Ishii and Ishii, 1978; Smith, Berger and Evans, 1981; Smith et al.9 1981; Jones and Milsom, 1982; West and Van Vliet, 1983). The receptors in the carotid labyrinth react only to pressures exceeding normal systolic pressure, and they appear to lack short term effects on the circulation (Ishii, Honda and Ishii, 1966; Smith, Berger and Evans, 1981; Smith et al., 1981). In reptiles and birds, baroreceptors occur in the aortic truncus near the heart, or within the heart itself. There is no solid evidence for a major baroreceptor function

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of any equivalent of the mammalian carotid sinus. The autonomic pathways in the reflex are mainly spinal autonomic, with both cardiac and vasomotor components (Seymour and Lillywhite, 1976; Lillywhite and Seymour, 1978; Berger, Evans and Smith, 1980; Berger et al., 1982; Lillywhite and Smith, 1981; Jones and Milsom, 1982).

CHEMORECEPTOR REFLEXES While baroreceptor reflexes directly affect the generation of blood pressure in the vertebrates, the chemoreceptor reflexes are, to a major extent, involved in ventilatory control mechanisms. Both central and peripheral receptor sites have been demonstrated in a number of vertebrate groups, but our knowledge of their relationship with the efferent (autonomic) pathways is rudimentary in the non-mammalian groups. The location of receptors sensitive to changes in the internal chemical milieu, notably C 02/pH and oxygen, varies between vertebrate groups, although the basic arrangement of glomus cells and sustentacular cells in the receptor unit remains surprisingly similar (Jones and Milsom, 1982). Activation of the chemoreceptors produces rapid ventilatory responses, especially in the water-breathing vertebrates. There are generally also effects on both cardiac performance and vascular resistance. Water-breathing species (fish) are far more likely to encounter an hypoxic environment than the air-breathing vertebrates. Thus, external chemoreceptors sensitive to the environmental levels of oxygen occur in the gills of e.g. teleosts (Burleson and Smatresk, 1990a, b; Kinkead et al., 1991; for references, see also Jones and Milsom, 1982; Fritsche and Nilsson, 1992). In the elasmobranch, Scyliorhinus canicula, a diffuse system of chemoreceptors resides in branchial areas innervated by cranial nerves V, VII, IX and X. Bilateral sectioning of all of these cranial nerves completely abolishes the hypoxic bradycardia and releases all of the vagal cardioinhibitory tonus (Butler, Taylor and Short, 1977). In teleosts, rapidly induced hypoxia similarly produces a bradycardia due to activation of cholinergic vagal pathways. Adrenergic, excitatory pathways to the heart are also activated, but their effect is masked by the cholinergic cardioinhibition (Fritsche, 1990; Fritsche and Nilsson, 1990). The location of the oxygen receptors is not entirely clear: both external (exposed to the water) and internal receptors (exposed to the blood) have been postulated (Kinkead et al., 1991). It does, however, appear that the first pair of gill arches, innervated by branches from the glossopharyngeal and vagus nerves, possess most of the receptors responsible for the hypoxic bradycardia reflex (Bamford, 1974; Smith and Jones, 1978; Fritsche and Nilsson, 1989; Burleson and Smatresk, 1990a, b). In addition to the reflexogenic bradycardia, hypoxia induces an elevation of both pre- and postbranchial arterial blood pressure, which in some species can be substantial (Holeton and Randall, 1967; Saunders and Sutterlin, 1971; Fritsche and Nilsson, 1989, 1990, 1992; Fritsche, 1990). In the cod, Gadus morhua, this increase in arterial blood pressure depends largely on an increase in the systemic vascular

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resistance due to an increased adrenergic nervous vasomotor tonus mediated by a-adrenoceptors (Fritsche and Nilsson, 1990). In the tetrapods, the arrangement of possible chemoreceptor sites is diffuse. In amphibians, chemoreceptors have been unequivocally demonstrated within the complex vasculature of the carotid labyrinth (Smith, Berger and Evans, 1981). In lizards, glomus and sustentacular cells are found at the base of the internal carotids, while in birds large congregations of chemoreceptive cells, the carotid bodies, are found near the thyroid (Abdel-Magied and King, 1978). Forced diving (submergence) of ducks produces a number of reflexes originating from stimulation of the nares, pharynx and glottis, but also from chemoreceptors. The treatment normally produces major bradycardia and an increase in the total peripheral vascular resistance, but this effect is largely prevented (bradycardia) or reduced (changes in the vascular resistance) after sectioning of the nerves to the carotid bodies (Jones and Purves, 1970; Jones and Milsom, 1982). EXERCISE Cardiovascular changes in response to exercise are, generally speaking, quite profound in all vertebrates. The general responses of the cardiovascular system to exercise have been relatively well elucidated in mammals. However, many studies of the physiological effects of exercise, especially in the non-mammalian vertebrates, have failed to differentiate between “exercise” and general “stress” due to various aspects of the experimental situation. Thus, our knowledge of the effects of true “exercise” is sometimes obscured by unfortunate experimental designs, which do not allow a reasonable appraisal of “stress levels” that may affect the interpretation. Simple pharmacological experiments have been used to elucidate the nature of the autonomic nervous pathways responsible for cardiac and vasomotor effects associated with exercise. In addition to “true” autonomic nervous effects, a number of other factors have been implicated in the vascular or cardiac responses to exercise and to physiological responses related to exercise. This began with GaskelPs (1880) suggestion of “chemical changes” as the cause for hyperaemia of active muscle, and later studies in mammals show the involvement of several specific factors (e.g. endothelial-derived relaxing factor) that may be triggered by exercise-related factors such as local hypoxia (Burnstock, 1990; Mione, Ralevic and Burnstock, 1990). The role of such factors in non-mammalian vertebrates is, however, not clear. The autonomic nervous effects during exercise in fish have received some attention. In teleosts such as the cod, Gadus morhua, and sea raven, Hemitripterus americanus, exercise reduces the cholinergic vagal tonus on the heart (Axelsson and Nilsson, 1986; Axelsson, 1988; Axelsson et aL> 1989b; Axelsson and Fritsche, 1991), which is similar to the situation in mammals (Folkow and Neil, 1971; Guyton, Jones and Coleman, 1973). In addition, exercise induces an increase in the adrenergic tonus on the heart of the cod (Axelsson, 1988). The adrenergic tonus is chiefly or solely of nervous origin, as demonstrated by the use of bretylium, which blocks adrenergic neurons (Axelsson and Nilsson, 1986; Axelsson, 1988). At high swimming speeds, the levels of catecholamines in the plasma start to increase, and when

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swimming speeds approach Ucrit (critical swimming speed, the maximum (calculated) speed), the increase in the circulating levels of catecholamines has been shown to marginally enhance the swimming performance (Butler et al., 1989). Vasomotor effects by adrenergic fibres become important during exercise. A decrease in vascular resistance occurs in teleosts during exercise, but in the cod this effect is offset by an increase in the adrenergic vasomotor tonus (Kiceniuk and Jones, 1977; Axelsson and Nilsson, 1986). A situation similar to that in exercising teleosts was demonstrated in the toad, Bufo marinus, by Wahlqvist and Campbell (1988). Exercise induces a rapid increase in the heart rate and blood pressure due to vagal withdrawal and an increased adrenergic nervous tonus. Later during the period of exercise, an increase in the levels of circulating catecholamines becomes important for the maintenance of the adrenergic tonus (Wahlqvist and Campbell, 1988). Also, in the urodele amphibian, Necturus maculosus, there is a tachycardia due to withdrawal of the vagal inhibitory tonus to the heart during exercise. Part of the tachycardia is due to an increase in the adrenergic tonus to the heart. However, the heart of Necturus does not seem to receive an adrenergic innervation (Axelsson and Nilsson, 1985), and the effect may be due to an increase in the concentration of noradrenaline in the plasma (Axelsson, Wahlqvist and Ehrenstrom, 1989). Similarly, the adrenergic vasomotor innervation in Necturus is scarce, and exercise-induced change in the vascular resistance is probably due to circulating catecholamines (Axelsson, Wahlqvist and Ehrenstrom, 1989).

CONCLUDING REMARKS The information presented here demonstrates that, with the notable exception of the cyclostomes, there are many similarities between the patterns of autonomic innervation of the heart and blood vessels across the vertebrates (see Figure 6.11). However, perhaps one example of an evolutionary trend in the autonomic control of the cardiovascular system concerns the control of the heart by catecholamines. While an inhibitory vagal innervation is present in all vertebrates (except cyclostomes), an excitatory innervation by spinal autonomic neurons that synthesize catecholamines is only apparent in the teleosts and the tetrapods. In the other fish, excitation of the heart is achieved by the release of catecholamines from chromaffin tissue located within the heart or in the systemic veins. There is now good evidence that an innervation of the vasculature by spinal autonomic neurons that synthesize catecholamines can be found in all vertebrates FIGURE 6.11 Diagram attempting to provide a generalized summary of the autonomic innervation of the cardiovascular system across the classes of vertebrates. Postganglionic neurons are shown as broken lines, preganglionic neurons are shown as solid lines. A vagal cardiac innervation in cyclostomes occurs in lampetroids only. Note that the splanchnic nerve in fish is composed of postganglionic neurons. Abbreviations: cc, chromaffin cells; grc, grey rami communicantes; la, lingual artery; pa, pulmonary artery; sc, sympathetic chain; sg, spinal autonomic (“sympathetic”) ganglion; spl, splanchnic nerve; wrc, white rami communicantes; VII, facial nerve; IX, glossopharyngeal nerve; X, vagus nerve.

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Cyclostome

Elasmobranch

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(cf. Burnstock, 1969). However, it may still be valid to conclude that control of the vasculature by circulating catecholamines is more prominent in fish than in tetrapods. Although the density of innervation of the vasculature by catecholaminesynthesizing neurons is, in general, lower in amphibians than in other tetrapods, there is no systematic difference between the density of vascular innervation in reptiles compared with mammals. In fact, there seem to be greater differences in the patterns of vascular innervation between members of the same vertebrate class that have different life styles, than between representatives of different classes. For example, both tree snakes and giraffes have developed patterns of vascular innervation that differ somewhat from those of other members of their classes, and that can be related to their need to deal with gravitational effects on the circulation (snakes: Lillywhite, 1987; Donald and Lillywhite, 1988; giraffes: Hargens et al., 1987; Nilsson et al., 1988; Kimani, Mbuva and Kinyamu, 1991). All tetrapods have a dense innervation of the buccal vasculature, particularly in the tongue, by cranial vasodilator neurons. Although there are some differences between the cranial nerve pathways in which these neurons lie in different vertebrates, the neurons supplying the tongue in most vertebrates examined contain the potent vasodilator, VIP. It is likely that these cranial neurons play important physiological roles during feeding in all tetrapods, and are also likely to participate in thermoregulation during panting in animals such as carnivores (Gibbins, Brayden and Bevan, 1984), ungulates, most birds and some lizards. An emerging theme in recent studies of autonomic neuroeffector mechanisms in the vertebrates is that transmission from autonomic neurons commonly involves more than one transmitter. In addition to the “classical neurotransmitters”, (nor)adrenaline and acetylcholine, there is good evidence that purines, peptides and nitrodilators play important roles in autonomic neurotransmission. To date, there is very little information on the functions of putative cotransmitters, particularly neuropeptides, in non-mammalian vertebrates. However, the available evidence indicates that there are not likely to be strict evolutionary trends in the choices of cotransmitters used by autonomic neurons. For example, some species of amphibians and mammals have somatostatin in their intracardiac vagal neurons, while intracardiac neurons in other amphibians and mammals lack somatostatin. Furthermore, there is no strict phylogenetic pattern to the distribution of the peptides NPY and galanin in vascular autonomic neurons of spinal origin: both NPY and galanin occur in catecholamine-synthesizing neurons in the sympathetic ganglia of some amphibians, reptiles and carnivores; sympathetic neurons in some Australian marsupials and lizards contain galanin, but lack NPY; whereas most of the catecholamine-synthesizing vascular sympathetic neurons in rodents contain NPY but lack galanin. Future studies providing physiological information on the roles of neuropeptides and other putative transmitters in cardiovascular autonomic neurons in a range of vertebrates will hopefully provide some explanation for the use of different combinations of cotransmitters by different classes of autonomic neurons.

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ACKNOWLEDGEMENTS Work from our own laboratories has been supported primarily by the National Health and Medical Research Council of Australia and the Swedish Natural Science Research Council, respectively.

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10 Chromatophores David J. Grove N uffield Fish L aboratory, School o f Ocean Sciences, University College o f N orth Wales, U.K. The adaptive responses of motile chromatophores of the lower vertebrates are controlled by “chromatic” centres (both excitatory and inhibitory) located in the optic tectum and brain stem. Differences between incident and background-reflected light are detected by the eyes and lead (a) to release of circulating hormones (e.g. melanophore stimulating hormone (MSH), melanine-concentrating hormone (MCH)) and (b) in many teleosts and some reptiles, to activation of nerve tracts which pass along the spinal cord. The central, antagonistic centres control the level of excitation in pre-ganglionic, cholinergic fibres which enter the sympathetic chain, increase neuronal activity and typically promote aggregation of chromatophore pigment through activation of sympathetic ganglion cells. The postganglionic nerve fibres are typically adrenergic and pass directly to the skin to innervate discrete “dermatomes”; the chromatophores lie in a “ground plexus” of terminals and each effector receives multiple innervation. Paling of the skin depends on activation of a-adrenoceptors (resembling the a 2-subtype) in most fish, but anomalous cases involving a cholinergic innervation are also known. Skin darkening may involve /3-adrenoceptor mediation but adenosine compounds, released as co-transmitters, disperse chromatophore pigment by both direct and indirect actions. Central control of complex colour changes in agonistic/courting behaviour and in adaptations to variegated backgrounds is not yet understood. KEY WORDS fish; chromatophores; autonomic; nerves

INTRODUCTION Fish, amphibians and reptiles are able to change shade or colour by redistributing pigment within effector cells (chromatophores). Such “physiological” changes are often contrasted with “morphological” changes caused by variations in the amount, or type, of pigment stored in the skin. However, only the teleost fish (and a few reptiles, see Bagnara and Hadley, 1973) use their autonomic nervous system to coordinate their physiological shade and color changes. The role of endocrine factors - such as the pituitary peptide hormones MSH, melanophore dispersing hormone, intermedin; (Lerner, Shizume and Bunding, 1954) and MCH; (Kawauchi et al., 1983) lie beyond the scope of this report. However, evidence has been put forward that a synergy between neuronal and hormonal factors is needed to obtain a maximal paling of the skin (Green and Baker, 1989). 331

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THE EFFECTORS Chromatophores are often separated into categories depending on the colour and chemistry of the pigment they contain (black or brown melanophores; red or orange lipophores: - erythro- and xanthophores; white guano-, irido- and leuco-phores) but the review that follows will be primarily concerned with the role of melanophores in co-ordinated changes of shade, since it is usually this category of effectors which can be controlled by the autonomic nervous system. Kuntz (1917) made histological studies of the skin of flounder (Paralichthys) and concluded that the cell itself retains a constant form during changes in skin shade, which depend on movements of the intracellular pigment towards or away from the centre of the chromatophore. Changes in the shade (black-grey-white) of a lower vertebrate to match the reflectance of the background are therefore brought about by re-distributing large numbers of 0.5 pun pigment granules (melanosomes) within the relatively static, star-shaped melanophore, thereby masking or exposing the underlying tissues to darken or pale the skin respectively (Figure 10.1 a, b). It has become conventional to refer to aggregation of all the melanosomes to the very centre of the cell as melanophore ‘‘aggregation”, often also called “contraction” or “paling”. The converse, where granules distribute into the cell branches, is called melanophore “dispersion”, as well as “expansion” or “darkening”. Details of the intracellular mechanisms which elicit melanosome movements are not of direct importance to this review but recent results and further references on this topic can be found in publications by Oshima, Inagaki and Manabe (1990a), Oshima, Hayakawa and Sugimoto (1990b), Fujii, Wakatabi and Oshima, (1991) and Rodionov, Gyoeva and Gelfand, (1991). Microscopically-observed changes in chromatophore “shape” in isolated pieces of skin have been followed by allocating an index (the semi-quantitative Hogben Melanophore Index; Hogben and Slome, 1931; Figure 10.1c), or photoelectrically (Fujii, 1959 and co-workers; Grundstrom et al., 1988). Responses in intact animals have often been recorded by visual comparison with standard grey shades (e.g. Healey, 1948 and co-workers; Grove, 1968; Figure 10.2).

COORDINATION OF M ELANOPHORE RESPONSES EYES AND CENTRAL NERVOUS SYSTEM In his review, Waring (1963) distinguished between several types of mechanisms which could affect melanophores. Non-visual responses were recognised (such as a primary or direct local reaction of melanophores to illumination, or even generalised nerve/hormone-linked responses caused by light striking the brain). Coordinated visual responses however require activation of the eyes by light, either to cause darkening (when the fish is on an illuminated dark background and only the ventral retina is stimulated : the secondary ocular response) or to cause paling (when the fish is on an illuminated pale background and the dorsal region of the

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FIGURE 10.1 Diagrams of melanophore activity. A: Melanophore (M) in skin of a teleost adapted to a white background. Aggregation of the melanosomes unmasks the white reflective guanophores causing pallor of the skin. B: After adaptation to a black background, the melanosomes disperse to overlie the guanophores and darken the skin. C: The degrees of pigment dispersion corresponding to the points 1-5 on the Melanophore Index (M.I.) of Hogben and Slome (1931). The unfilled branches of the melanophores are not seen in conventional light-microscopy.

retina was also stimulated: the tertiary ocular response). Such visual responses involve the central nervous system; nerves and/or hormones then change the state of melanophores in the skin to match the prevailing background. The present review discusses the evidence for the participation of the autonomic nervous system in the rapid adjustment of melanophore state, which is often achieved in a few minutes, leading to changes in body shade to match the background. It will become clear that, despite widespread interest in the topic over more than a century, the progress of various authors was greatly limited by the currentlyavailable techniques. Many necessarily resorted to indirect (and often novel) experiments to deduce the underlying mechanisms involved in adaptive shade and colour changes. Most of the detailed studies of the systems which coordinate melanophore responses have been made in fish. Pouchet (1871, 1872, 1876) showed that turbot (Scophthalmus) lose their ability to darken or pale, in response to exposure to black

334 COMPARATIVE PHYSIOLOGY AND EVOLUTION

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Time (minutes) FIGURE 10.2 Changes in skin shade of Blennius pholis, recorded at 14°C, by comparison with the Derived Ostwald Index (DOI) standards of grey (Grove 1968). Low DOI values indicate pallor and high values are dark. At the start, five white-adapted (filled circles) and five black-adapted (open circles) fish were transferred to the opposite background, and returned after 1 hour. Note that a) a new adaptation to a white background requires more than an hour to complete and b) that return to the original background (in either direction) is faster than a new adaptation to that background. Fish with anterior spinal cord section require more than 24 hours to achieve similar changes. This suggests that hormonal influences (MSH, MCH) may also affect the rate of change in vivo.

(B) or white (W) backgrounds, if the eyes were destroyed. The fish retained an intermediate shade of grey when illuminated, according to most authors. However, Biiytendijk (1911) extended this work to indicate the possible involvement of central nervous modulation of the response to blinding. He reported that removal of one eye from Scophthalmus led to preliminary darkening followed by re-adaptation to the existing background. Removal of the second eye, however, caused the animal to remain pale or dark, depending on the background to which the fish was fully adapted before complete blinding, von Frisch (1911) noted that, after removal of one eye, cyprinids darkened but re-adapted to their background while, in salmonids, removal of one eye leads to a permanent darkening of the opposite side of the body!

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Studies using partly/completely-blinded fish, and/or directional light beams, had been undertaken by Semper (1892) on Macropoda, Schondorff (1903) on Salmo, Secerov (1909) on Noemacheilus and by Mayerhofer (1909) on Coitus, Perea and Esox. The general conclusion was that B-adaptation (darkening) and W-adaptation (paling) of the fish depended on differential stimulation of areas of the retina of at least one eye by light. Sumner (1933) used black celloidin to make false corneas thereby obtaining partially-blind Fundulus parvipennis. He concluded that adaptive colour changes were dependent on the relative intensities of illumination of the upper and lower portions of the retina. Illumination of the lower retina “acted positively” to cause darkening. Butcher & Adelman (1937) studied the effect of rotating the eyes of Fundulus surgically (a method also used by Vilter (1937) to study axolotl responses) on colour changes; they too agreed that illumination of the (normally) ventral retina promoted darkening. Later, Butcher (1938) exposed Fundulus to varying ratios of direct to reflected light; he again concluded that adaptive responses of the fishes melanophores depended on this ratio but also suggested that illumination of one region of the eye (e.g. upper retina) can inhibit melanophoric responses (darkening) caused by illumination of other retinal regions. Danielson (1939, 1941) similarly stimulated eyes of Nocomis and Semotilus and concluded that as the ratio of overhead to reflected light increased, so the fish darkened (and vice versa). Hogben and Landgrebe (1940) used directional light stimuli and concluded that the retina of Gasterosteus was differentiated into a ventral region (“B”), which induced the fish to darken, and a dorsal region (“W”), which controlled paling. The information gathered by the eye is transferred to the brain by the optic nerves. Polimanti (1912) recorded that Megarini in 1884 was the earliest worker to envisage specific chromatophore-controlling centres residing in the brains of Phoxinus and Salmo. The detailed studies by von Frisch (1911) on these species showed that blinded individuals developed fully dark skins only when their diencephalon was illuminated; otherwise they were an intermediate shade of grey. He therefore proposed that a melanophore-dispersing centre lay in this part of the fish brain. Electrical stimulation of the medulla oblongata, on the other hand, revealed the presence of a melanophore-aggregating (paling) centre. Schaefer (1921) reached a similar conclusion for Pleuronectes. More recently, Iwata & Fukuda (1973) stimulated the brain of crucian carp, especially the optic tectum, whilst recording melanophore responses in the skin photoelectrically. The median part of the tectum (which receives input from the ventral retina of the contralateral eye) darkens the skin when stimulated, whereas paling is associated with the latero-ventral part of the tectum (which receives projections from the dorsal retina). Neural tracts were detected running to the medulla oblongata and spinal cord in: 1. 2. 3. 4.

tractus tractus tractus tractus

tecto-bulbaris dorsalis tecto-bulbaris ventralis incruciatus (ipsilaterally) tecto-bulbaris ventralis cruciatus(contralaterally) tecto-spinalis.

Electrical stimulation of tract 2 (and less strongly 3 and 4) usually caused paling

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FIGURE 10.3 Diagrams of the early descriptions of central and autonomic nerve tracts which control melanophores in fish. The spinal and peripheral pathways of the “chromatic tracts” in Phoxinus, as described by von Frisch (1911), are shown as continuous lines. The effects on the skin shade of whiteadapted fish after surgical lesion are indicated by the corresponding diagrams below. 1: Anterior spinal cord section. 2: posterior spinal cord section. 3: Bilateral section of the anterior (double) sympathetic chain. 4: Section of the posterior (single) sympathetic chain. 5: Section of 2 spinal nerves on the left side. The dotted lines indicate the postulated course of peripheral, darkening fibres suggested by von Gelei (1942).

whilst slight darkening was associated with tract 1. In the medulla, complex responses were obtained but large parts of the reticular formation evoked darkening, or initial quiescence followed by paling. Such responses could be traced along the dorsal and lateral columns of the spinal cord. (In addition darkening (B) and paling (W) areas were also found in the posterior dorso-lateral and the anterior dorsal regions of the telencephalon respectively, which also appeared to link to the medulla). They concluded that central control of melanophore responses included a dual, antagonistic system of central nerves which are respectively excitatory and inhibitory in their final connections with the medulla oblongata and spinal cord. Using careful surgery to place lesions in the central and peripheral nervous system in Phoxinus (Figure 10.3), von Frisch (1911) had found that nerves from the medullary paling centre pass along the spinal cord to the level of ca. vertebra 15

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before leaving the spinal cord (although Burton in 1964 found that in British stocks of minnow the exit was at vertebra 12). In his study, von Frisch had applied the techniques of nerve lesion and electrical stimulation employed earlier by Pouchet (1871, 1872, 1876) in his elegant study of chromatic tracts in the turbot, Scophthalmus. Similar studies on flatfish were made by Scott (1965) and Fernando and Grove (1974a). Burton (1964, 1966, 1968) used more precisely-located lesions to localise the tracts to the region around the base of the dorsal horns of the spinal cord of Phoxinus. The autonomic chromatic functions are associated with the dorsal part of the corpus commune posterius and substantia gelatinosa rolandi. Partial damage to these regions produces a prominent dark mottled pattern in the skin of white-adapted minnows. Individual melanophores of these fish showed varied degrees of asymmetrical dispersion, suggesting that each melanophore ultimately receives a multiple innervation. Furthermore, fibres of any specific region of the spinal tract seem to be associated with melanophore-controlling fibres with a wide cutaneous distribution, rather than for a given body segment. In crucian carp, surface medial stimulation of the anterior spinal cord induced darkening of the skin, but the active region decreased in size posteriorly. Internal stimulation of the cord caused paling. This led Iwata and Fukuda (1973) to conclude that there are two types of neurones in the spinal, chromatic tract of fish: excitatory (promoting paling) and inhibitory (suppressing activity in the paling motoneurones). It seems that most inhibitory chromatic neurones are found in the anterior spinal cord, since if the darkening were caused by “pigment dispersing nerve fibres” the response should not (as it did) diminish along the cord. This dineuronic model of antagonistic, central neuronic tracts in the anterior spinal cord of the carp supports the observations of Healey (1951, 1954) who concluded that active darkening tracts, if present, must accompany the paling fibres along the anterior part of the spinal cord. THE SYMPATHETIC CHAIN AND PERIPHERAL TRACTS Unlike their findings within the central nervous system, Iwata and Fukuda (1973) could only elicit paling responses when the sympathetic chain of Carassius was stimulated. The chain conducted spontaneous impulses (up to 20 Hz) as long as the spinal pigmento-motor tract was intact, and especially during paling. These discharges decreased in frequency when brain stimulation induced darkening. No bursts of discharges in the chain were detected during darkening of the skin. Apparently skin colour in the carp depends on the rate of activity in a single (paling) class of nerves whose fibres run in the sympathetic chain. Their activity is promoted by excitatory spinal motoneurones, which are themselves subject to control by central excitatory and inhibitory neurones. Presumably the excitatory neurones which leave the spinal cord travel in ventral roots of spinal nerves to join the sympathetic chain by way of grey rami communicantes. This “peripheral mononeuronic theory”, based on electrophysiological techniques, contrasts with the “dineuronic” interpretations described below.

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In the figures presented by von Frisch (1911) and later von Gelei (1942), based on sympathetic chain lesions in Phoxinus, the pre-ganglionic (paling) fibres emerging from the spinal cord apparently run forwards or backwards along the chain. Once in the sympathetic chain, the preganglionic fibres (presumably cholinergic see below) synapse with postganglionic neurones which pass to the skin by way of white rami comunicantes which rejoin the spinal nerves. However, Nicol (1952) found anatomical evidence that single nerve fibres run throughout the length of the sympathetic chain, giving off collateral branches within each segmental ganglion to innervate final motor neurones. In the head, the tracts pass to the skin by cranial (e.g. trigeminal) nerves, van Rynberk (1907) had shown in Solea and Scophthalmus that the pigmento-motor fibres originating in segmental ganglia of the “Nervus sympathiesf* are distributed entirely within that area of skin (dermatome) supplied by sensory nerves of the corresponding spinal ganglion, van Herk (1929) made similar nerve lesions in the flounder Platichthys. He added, however, that each segmental area may overlap its neighbours by somewhat more than half. In his early histological studies, Kuntz (1917) had already suggested that the small, sheathless nerves he saw around flatfish melanophores originated from the sympathetic nervous system. According to all these early reports, and that of Bauer (1910), in vivo nerve sections at any level in the sympathetic chain of a W-adapted fish rapidly induce intense pigment dispersion (darkening) in the downstream, “paralysed” melanophores. This happens despite the presence of circulating hormones (such as MCH). Such denervated melanophores later develop slow, adaptive responses which appear to be under endocrine (e.g. pituitary) control. Electrical stimulation of the same tracts in unoperated, black-adapted fish causes rapid melanophóre aggregation. However, it has become clear that the physiological links within the sympathetic chain are not as simple as von Frisch (1911) envisaged, and early workers found surprising responses to lesion, stimulation and drug tests. Schaefer (1921) stimulated the skin of Pleuronectes electrically and found that the whole animal would pale, even if the stimulated area was completely bounded by cuts in the skin. It was essential for the stimulated area to be in contact with the nervous system for, if the spinal cord and sympathetic chain were destroyed, the pallor would not spread beyond the isolated square of skin. This does not conform with von Frisch’s account of the anatomy of the peripheral chromatic tracts. von Wernóe (1928) adopted a novel approach. He examined the complete body pallor which develops in the skin of freshly-killed fish (plaice, cod and eel). This pallor for any given skin area depended on unbroken connections between the brain (where the paling centre lies), the anterior spinal cord and the sympathetic chain. Spinal paling motoneurones may discharge spontaneously as central functions break down. Apparently the post mortem paling response depends on the integrity of von Frisch’s paling fibre tract. von Wernóe also examined the effects of application of nicotine (to block cholinoceptors on ganglion cells) to stated sympathetic ganglia in freshly-killed fish. Nicotine initially paled the appropriate region of skin as the sympathetic nerve cells of that body segment were excited. The skin subsequently darkened as nicotinic

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blockade set in. If the sympathetic chain was then stimulated electrically, the paralysed dermatomes did not respond, but the other regions paled. He concluded, as Langley had found for tetrapods (Langley and Dickinson, 1889), that where sympathetic paling fibres running in the sympathetic chain synapse in the segmental ganglion before innervating the skin, they were susceptible to nicotinic blockade. Other (pre-ganglionic) fibres passing through that segment of the chain were unaffected by the nicotine. He went on to inject nicotine into the whole fish, rather than “paint” localised ganglia with the drug. This treatment prevented any electricallystimulated pallor in any segment of the body, except that segment where the electrodes were placed, presumably because postganglionic tracts to the spinal nerve were activated directly. This led him to the conclusion that all paling fibres leaving the spinal cord (in von Frisch’s chromatic tract) must synapse at least once before reaching the melanophores. Of greater interest was his finding that fish (that had their brain and spinal cord destroyed previously) paled completely even on weak electrical stimulation of the sympathetic chain in the caudal region of the body. That this was not due to direct stimulation of the melanophores by an unspecific leak of the stimulus was shown by sectioning some spinal nerves, whereupon the appropriate segments of skin failed to respond. If, instead, the sympathetic chain was transected near the middle of the body, the pallor extended only up to the level of the cut. He concluded that caudal stimulation of the sympathetic chain eventually recruits all postganglionic tracts unless blocked by nicotine. von Wernoe was faced with several alternative explanations for his observations, and no adequate study of the connections along the sympathetic chain has yet been made. It was difficult for him to conceive that weak electrical stimulation of the sympathetic chain in the caudal region could pale the anterior part of the fish by leakage of the stimuli to other, anterior, pre-ganglionic chromatic tracts stemming from the central nervous system, or by spread through the skin. He was left with the conclusion that chromatic tracts to different parts of the body are linked functionally. His experiments, in which both of the (double) anterior sympathetic chains were cut, clearly showed that the postganglionic chromatic fibres (e.g. in the tail region) did not enter a peripheral (e.g. dermal) nerve net which could conduct impulses to the head. He discounted the idea that preganglionic tracts or neurones form a specialised nerve net capable of conducting impulses in a variety of directions, as it is difficult with such an anatomical arrangement to understand how (for example) flatfish might adapt to patterned backgrounds (e.g. Mast, 1916). He therefore believed that his work would be best interpreted on the basis that chromatic nerve outflow from the central nervous system led to excitation of one large, “chromatic” neurone in the sympathetic chain whose branches and collaterals innervate every postganglionic, chromatic nerve fibre. It is difficult to accept this conclusion, particularly as it is not easy to see how such a single sympathetic neurone could control often-reported, adaptive changes in body patterns to variegated backgrounds. It is however clear from Wernoe’s studies that all chromatic pathways from the spinal cord to the skin involve synapses in the sympathetic chain, and that the chain can conduct stimuli in a number of directions. In this

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respect at least his work receives some support from the anatomical studies of Nicol (1952; see above). Nevertheless, studies of this type should be repeated with modern techniques so that the problems raised can be answered. MONO- OR DI-NEURONIC CONTROL OF EFFECTORS? Because of the prevailing view, based on mammalian studies, that tissues innervated by “sympathetic” nerves would also have anatomically-distinct, antagonistic “parasympathetic” nerves, von Frisch (1911) envisaged the possible existence of a set of opposing, darkening (= melanophore dispersing) fibres running alongside the aggregating fibres; he had no direct evidence for this, save that fish (like reptiles) could change shade quickly from white to black (and vice versa) and that both these fast responses to a new background shade were abolished by a single lesion in the spinal cord, the sympathetic chain or individual spinal nerves. Later Healey (1940 et seq.) and Hogben and Landgrebe (1940) carefully recorded the time course of such shade changes, a contribution which came to play a significant part in interpreting experimental results. In 1916, Spaeth demonstrated that melanophores aggregate when treated with adrenaline. Giersberg (1930) injected Phoxinus with an a-adrenoceptor antagonist (ergotamine), which also possesses weak intrinsic agonist activity of its own, thereby causing a mild paling of the fish skin. He then injected acetylcholine to “enhance’’ the activity of a postulated cranial para-sympathetic system (in keeping with the prevailing view of physiology at that time), which he believed should control darkening of the minnow skin. When the spinal cord was stimulated electrically, he found that the fish indeed darkened. He took this as evidence in support of the proposed melanophore-dispersing nerves postulated by von Frisch. In 1942, von Gelei combined this work with the addition of crude lesions made in the sympathetic chain, an operation performed by introducing a sharp knife through the body wall of the fish, cutting other tissues and blood vessels. Melanophore dispersion, elicited by electrical stimulation of the hind brain or anterior end of the spinal cord, was always limited posteriorly by the site of a lesion in the sympathetic chain. This claim differed dramatically from the pattern described by von Frisch for paling fibres: in the anterior of the body (in front of vertebra 15) a sympathetic chain lesion forms the anterior limit of the paling fibres’ influence; anterior portions of the fish darken, von Gelei believed that he had demonstrated that a special tract of melanophore-dispersing nerve fibres emerged from the minnow spinal cord through the rami communicantes of the 1st and 2nd spinal segments. This set of nerves would pass along the sympathetic chain to achieve “double” innervation of the skin melanophores by way of the spinal nerves (Figure 10.3). von Gelei’s conclusions have been disputed by a number of workers. Healey (1951, 1954) conducted an exhaustive series of experiments involving severance of the spinal cord of minnows anterior to the 1st vertebra (which would disconnect both von Gelei’s “dispersing” and von Frisch’s aggregating tracts to the melanophores). When such fish were transferred between different backgrounds, he could detect no difference in the rate and degree of shade change between such spinal-

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sectioned fish and others which had received a spinal lesion at a site varying between the 1st and 15th vertebrae (thereby presumably disconnecting only the W-fibre tract). The latter group would have their paling, but not darkening, nerve tracts to the skin severed, on von Gelei’s hypothesis; the timing of shade changes in the second group should have been much faster from W to B than the reverse. Healey and Ross (1966) attempted to repeat von Gelei’s experiments. They again injected ergotamine to block a-adrenoceptors and then stimulated the anterior spinal cord, whereupon the whole animal became dark - interestingly without the need for the injections of acetylcholine used by von Gelei. They concluded that ablockade had “reversed” the action of adrenergic transmitter, presumably by unmasking an effect on /3-adrenoceptors which mediate dispersion. Such qualitative “reversal” of sympathetic nerves had been known since the work of Dale (1913) and had later been suggested by Gray (1956) as an explanation for both Giersberg and von Gelei’s findings. In a later in vitro study, Pye (1964a, b) attempted to reverse the paling action of adrenaline on isolated Phoxinus skin, by using ergotamine. The paling action of the catecholamine was not “reversed” to cause melanophore dispersion, casting doubt on the idea that an adrenergic darkening (melanophore dispersing) nerve tract had been exposed by in vivo ex-adrenergic blockade, when combined with sympathetic nerve activation or stimulation used by other authors. Unfortunately, at this time the development of suitable /3-adrenoceptor blocking agents was in its infancy and the experiments could not be completed. Pye (1964a) also challenged von Gelei’s conclusion on the precision of gross electrical stimulation methods to determine the likely anatomical pathways of “dispersing” fibres. He found that melanophore responses could be evoked using an electrode placed at any level of the spinal cord of the minnow (whether anterior or posterior to the point of chromatic tract outflow to the sympathetic chain). He even found responses when a small, posterior (non-“chromatic”) section of the spinal cord, isolated by two consecutive transverse sections but in connection with the peripheral nervous system (by spinal nerves and rami communicantes), was stimulated electrically! He assumed that the known chromatic tracts (mapped by von Frisch using lesions) had been indirectly stimulated in his own experiments by pathways as yet undescribed. Pye concluded, in agreement with Healey, that if darkening fibres do innervate the skin melanophores, they follow paths similar to those taken by the paling fibres. Such reports draw into question the use of crude electrical stimulation of chromatic nerve tracts to deduce anatomical pathways, whereas similar criticism may not apply to interpretation of the effects of carefully introduced nerve lesions in vivo, nor to the use of microelectrodes. Because of these uncertainties, other workers have attempted novel and at times imaginative techniques to study the nerve tracts as they approach the periphery to innervate melanophores. In 1924, Wyman showed that small lesions in the fins of teleosts such as Fundulus caused dispersion of “downstream” melanophores which was not due to indirect effects such as damage to the local vascular bed. Parker and his school claimed that their studies on such temporary, dark caudal bands in Ameiurus and Fundulus (reviewed in 1948) demonstrated the presence of a system

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of melanophore dispersing nerves which oppose the effects of the well-established paling fibres described by Pouchet, von Frisch and others. Two of the experiments cited in support of his claim seem to be important. First, a new cut within a recently “denervated” and faded band produces melanophore re-dispersion both ortho- and anti-dromically. Secondly, application of a cold tube distal to a new cut limits the spread of the region of dispersed melanophores, presumably by interrupting neural conduction. Parker argued that the initial dispersion was an active process produced by repetitive discharges in segments of darkening fibres, and originating from a permanently depolarised region at the site of the nerve lesion. Such activity might be analagous to the condition set up in pain fibres after trauma. Parker and Rosenblueth (1941) claimed to have caused the skin of Ameiurus to darken by stimulating B-fibres in the chromatic tract selectively, using rectangular, 300-500 msec pulses at a rate of 1-2 pulses per second. Pye (1964a) pointed out that such stimulation used inappropriate electrodes and also approximated the application of a direct current, thereby simply interfering with on-going activity in the W-fibres. Gray (1956) repeated Parker’s fin-cutting experiments using Phoxinus instead of Fundulus. Cuts made in pale, either intact or hypophysectomised, fish effectively disposed of the concept that severance merely released melanophores from aggregating nervous control, thereby permitting circulating MSH to disperse the melanophore pigment. After an initial cut, melanophore dispersion was complete in 15 minutes, and disappeared in 12 hours if the animal was kept on a white background. As the denervated area faded, the pallor encroached from the lateral edges of the cut band towards the centre. On transferring the fish to black, the skin darkened uniformly in both innervated and denervated areas (without encroachment of pallor from the sides). Gray drew the conclusion, based on these and other experiments, that: 1. 2. 3.

immediately on denervation, an inherent dispersing mechanism in melanophores comes into play. subsequently they lose refractoriness to diffusing neurohumours, and may even become hypersensitive. Asymmetrical and partial responses of melanophores after nerve section indicate that each cell is controlled by more than one fibre (observations which confirm earlier reports by Mills 1932 a, b, c and Abramowitz 1936a, b).

These responses he believed are most easily explained by a dual-innervation model based on both B and W fibres. His first point may be supported by the discovery of adenosine receptors (Miyashita, Kumazawa and Fujii, 1984) and the second point is confirmed by the sensitivity changes following denervation described by Grove (1969a), both of which are discussed below. However, Vilter (1939) studied the effect of lesions in various cyprinid, gobiid and other fish. He confirmed that adjacent nerves running to the skin dermatomes overlapped (to an extent varying with species). A new, distal cut can affect nerve branches previously unaffected by the first cut. Such innervation patterns call

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Parker’s interpretation, that damage activates or re-activates the living terminations of cholinergic B-fibres, into question. Healey (1967) also placed lesions in the chromatic tract of Phoxinus, during its passage along the anterior part of the spinal cord. Fish were left for up to 11 months (which allowed at least partial nerve regeneration) and their remaining slow or rapid responses to background changes carefully analysed. He concluded: 1. 2. 3.

That a number of chromatic neurones were present in the anterior spinal cord. That each of these neurones contributes to the state of excitation of the postganglionic, autonomic nervous system which affects all the melanophores of the skin. That, during nerve regeneration, asymmetrical responses of some melanophores indicated that both B- and W-nerve fibres pass to the skin.

With the procedures used to date, the problem of whether two antagonistic types of autonomic nerves effect coordinated melanophore responses can not be resolved. PERIPHERAL INNERVATION OF THE SKIN Early histological studies (Ballowitz, 1893; Eberth and Bunge, 1895; Whitear, 1952) revealed the presence of nerve nets in the dermis; each melanophore is surrounded by many fine branches. Whitear described fibres which branch at some distance from the melanophores, travel in all directions and which reach several effector cells. Fujii and Novales (1969) studied the response of individual melanophores to electrical stimulation using “split fin” preparations. Individual melanophores can be easily observed, drugs and hormones penetrate the exposed dermis rapidly and the main chromatic nerves run parallel to the fin rays. They found that even a single electric shock to the nerves induced detectable melanophore pigment aggregation; the response was not “all or none” but increased with stimulus intensity. They concluded that several fibres control each melanophore cell, in a pattern similar to that found for the autonomic ground plexus in higher vertebrates (Burnstock, 1970). It is generally accepted that most fish control rapid pigment-aggregating responses of the melanophores by way of autonomic postganglionic fibres (Iwata and Fukuda, 1973). The terminal regions of postganglionic neurones reaching the melanophores and forming a ground plexus in the dermis have been shown by fluorescence histochemistry to be adrenergic, with recognisable varicosities, where catecholamines (mainly noradrenaline) are thought to be released (Jacobowitz and Laties, 1968; Falck, Muntzing and Rosengren, 1969; Fernando, 1989). The adrenergic nature of the terminal nerve fibres has also been demonstrated by microscopic autoradiography of split tail fins labeled with 3H-noradrenaline (Miyata and Yamada, 1987). In the fins, the peripheral fibres may form parallel bundles (Fujii and Taguchi, 1970; Miyata and Yamada, 1987). Dense-cored vesicles believed to contain catecholamines have been detected in the abundant pre-synaptic endings on the melanophores of Lebistes (Fujii and Taguchi, 1970), and electrical stimulation or application of KC1 induces a marked reduction in the number of these vesicles (Miyata, 1987).

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A number of authors have used simple salt solutions on isolated skin or scales to cause the melanophores to fully disperse or aggregate their pigment prior to experimental tests with agonists and/or antagonist drugs. Treatment with K+, especially in the presence of Ca2+, induces skin pallor, caused by release of catecholamine, whilst Na+ causes skin darkening (e.g. Fernando and Grove, 1974b; Ali and Ovais, 1983; Miyata, 1985; Patil and Jain, 1989; Burton 1990; Burton and Everard, 1990; Miyashita and Moriya 1990). Purines may be simultaneously released by K+ (Kumazawa and Fujii, 1984). K+ also releases acetylcholine to induce skin pallor in the cholinergically-innervated catfish, Kryptopterus bicirrhi (Kasukawa and Fujii, 1984). Electrical stimulation of the fish skin also shows that the dermal net or plexus conducts both ortho- and antidromically, thus aggregating melanophores, suggesting that the paling neurotransmitter is released from a number of sites along the nerve terminal (Fujii and Novales 1969). Obika (1988) has shown that the peripheral nerves in Oryzias may control melanophore aggregation and leucophore dispersion, leading to a double mechanism controlling skin paling; a similar pattern was also found in Odontobutis (Iga, Takabatake and Watanabe, 1987). MELANOPHORE PHARMACOLOGY AND RECEPTORS The idea of a dual peripheral innervation of melanophores by antagonistic autonomic nerves, on the evidence reviewed so far, remains unproven. Further information has been obtained by examining the types of neurotransmitter associated with the nerve-melanophore “junction’’ and the receptors which melanophores carry. Many authors have found that directly- and indirectly-acting adrenergic agonists induce melanophore aggregation in the majority of teleosts by way of «-adrenoceptors; this action is suppressed by «-adrenergic blockade (reviewed by Grove 1969a, Fernando and Grove 1974a, b; Kasukawa et al., 1985). Agents, such as reserpine, which deplete the neurotransmitter stores (Jacobowitz and Laties 1968) suppress both rapid melanophore aggregation and dispersion in vivo (Grove 1969a). Results from a series of pharmacological studies using several fish species strongly suggest that the «-adrenoceptors are of the «2 subtype, and mediate aggregation of the melanosomes via inhibition of adenylate cyclase activity (Andersson, Karlsson and Grundstrom, 1984; Karlsson et al., 1987; Svensson, Karlsson and Grundstrom, 1989). However, there are clear signs of phylogenetic divergence in receptor characteristics amongst these species (Karlsson et al., 1987). The «-adrenoceptors of Symbranchus marmoratus melanophores cannot be classified as or «2, but might on the other hand not be innervated (Abrao et al., 1991). Pigment aggregation in erythrophores and xanthophores, and iridophore pigment movements which alter reflectance to longer wavelengths, are also mediated by «-adrenoceptors (Fujii and Oshima, 1986; Iga, Takabatake and Watanabe, 1987; Karlsson et al., 1988a; Nagaishi and Oshima, 1989b; Oshima, Kasukawa and Fujii, 1989). The control of iridophore movements in the freshwater goby, Odontobutis obscura, is mediated by «-adrenoceptors which are activated by the same sympathetic nerve fibres that control adjacent melanophores (Iga, Takabatake and

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Watanabe, 1987). The a-adrenoceptors on the erythrophores of the squirrelfish, Myripristis occidentalism have been classified as a 2-adrenoceptors (Karlsson et al., 1988a). Interruption of the paling nerve tract, by lesions or by drug treatment, causes notable super-sensitivity of the melanophores (Grove 1969a), in the increasing order: control < spinal section < cocaine < spinal nerve section. Such super sensitivity follows Cannon’s “Law of denervation” as extended by Trendelenburg (1966); the in vivo ED50 of fully denervated melanophores of Phoxinus is 0.2ng/g compared with 7 ng/g in unoperated fish. The melanophores of cuckoo wrasse, Labrus ossifagus, were studied in vitro by Karlsson et al. (1988b). They confirmed that cocaine caused supersensitivity to noradrenaline (8-fold) during the 3 days that the nerve terminals remained functional. Prolonged culture (8 days or more) led to a much greater supersensitivity (300-fold) which apparently depended on changed melanophore physiology, such as an increase in the adrenoceptor population. Such studies suggest that there is no nerve/nerve synapse outside the sympathetic chain before the postganglionic neurones reach the melanophores. A number of authors have also pointed out that different (micro and macro) melanophores in the skin of untreated fish differ noticeably in their sensitivity to catecholamines. The macromelanophores of Phoxinus are the last to pale and the first to darken during coordinated shade changes. The pattern made by these effectors causes “bars” to appear on the fish, presumably aiding as a disruptive camouflage for the individual during adjustment to new backgrounds. It should be emphasised that the melanophore-aggregating effect of the pituitary hormone, MCH, does not exert its effects by way of the same receptors that are activated by the nerves. The hormone effect persists in those fish which have cholinergic receptors (Nagai, Oshima and Fujii, 1986) and is not abolished by receptor blockade (Oshima et al., 1985; D.J. Grove, unpublished observations). Spaeth and Barbour (1917) showed that the aggregating action of adrenaline was transformed to a darkening action after treatment with ergotoxin. Several more recent studies have been carried out on “adrenaline reversal”, to test whether melanophore dispersion can be caused by the action of catecholamines on /3adrenoceptors. Although Pye (1964b) and Grove (1969a) were unable to detect /3-responses in Phoxinus, Fujii and Miyashita (1976b) were able to show that /3agonists do disperse Lebistes melanophores after a-adrenoceptor blockade. This dispersion effect has been observed in other species for melanophores (Iga, 1983; Kasukawa et al.9 1985; Morishita, Katayama and Yamada, 1985) and other chromatophores (Iga, Yamada and Iwakiri, 1977; Matsumoto etal., 1978; Yamada, 1980; Morishita Katayama and Yamada, 1985). Enami (1940) and Fujii and Miyashita (1976a) found that Parasilurus melanophores were especially unusual in that «-adrenoceptors are absent; adrenaline dispersed them, by an action on /3adrenoceptors. The precursor dopamine had a similar effect, but acted on separate receptors. Parker (1948) and his co-workers had proposed that teleost fish possess “parasympathetic” darkening fibres and believed that these were cholinergic. Responses of melanophores to treatment with cholinesters are inconsistent and contradictory

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(Grove, 1969b). Furthermore, the muscarinic blocker atropine, which should theoretically enhance paling on Parker’s model, has consistently been found to disperse melanophore pigment. Mention should again be made however of the catfish (Parasilurus), as well as the glass catfish (Kryptopterus) and other silurids (Fujii and Miyashita, 1976a; Kasukawa, Oshima and Fujii, 1986). In all these species, extrinsic catecholamines disperse the melanophores by action on jS-adrenoceptors whilst the sympathetic tracts which aggregate the melanophores are, unexpectedly, cholinergic and atropine-sensitive. In the more aberrant “upside-down” catfish (Synodontis), the dorsal melanophores receive adrenergic nerves, which induce paling by way of a-adrenoceptors. In addition, /3-adrenergic responses, mediating dispersion (see below), can be demonstrated (Nagaishi and Oshima, 1989a). The small ventral melanophores are less sensitive to a-agonists. Instead, the j8adrenoceptor-mediated dispersion effect is dominant, thereby keeping the ventral (upper) skin darker, presumably to effect countershading camouflage. Several authors have re-examined melanophores to test whether rapid, nervemediated pigment dispersion (darkening) can be demonstrated to play a part in chromatic responses. Fujii (1959) had developed his “split fin” preparation to study such chromatophore responses in fish. He found that denervated melanophores dispersed more slowly than non-denervated ones, when recovering in Ringer after catecholamine treatment. He suggested that sodium ions might activate dispersing fibres. However, denervated melanophores are markedly super sensitive to catecholamines (see above) and this change can explain the delayed recovery. Watanabe, Izumi and Iwata, (1962) made the interesting observation that dispersion of melanophores in Ringer solution after adrenaline-induced aggregation was more rapid at higher amine concentrations, and that a second component of catecholamine action must exist. Novales and Fujii (1970) and Fujii and Miyashita (1976b) had also shown that the jS-adrenergic dispersion of Lebistes melanophores was mimicked by cyclic-AMP. This agent, and especially its dibutyryl derivative, has a clear dispersing effect on melanophores of Pleuronectes (D. J. Grove and G.M. Campbell, unpublished observations), and may act as an internal, second messenger (e.g. for the hormone MSH). However, other adenine derivatives are also effective in dispersing fish melanophores, and are now known to act by way of adenosinespecific receptors (Miyashita, Kumazawa and Fujii, 1984). Adenosine itself inhibits release of noradrenaline from aggregating nerves (Oshima, 1989). ATP is released as a co-transmitter from the adrenergic nerves (Kumuzawa et al. 1984, Kumuzawa and Fujii, 1984). It may function (after dephosphorylation) to promote rapid pigment dispersion when activity ceases in the paling fibres and the catecholamine transmitter has been rapidly inactivated and/or removed from the nervemelanophore junction. This last finding is of great interest. A major stumbling block to proponents of the mononeuronic model of melanophore control has been the rapid dispersion of freshly-denervated effector cells (in the face of circulating pituitary hormones) when a fish adapts to a dark background, or especially when a chromatic tract is sectioned. If a longer-acting, antagonistic co-factor is released by the same paling nerve terminals, this weakens the case for presence of separate B-fibres. In support of this

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FIGURE 10.4 Diagram to show the likely central and peripheral autonomic tracts which control melanophore responses in teleost fish (but not Siluridae [see text]). Incident light rays (ir) stimulate the ventral retina (v) and activate “darkening” neurones (filled circles with dashed lines) passing to the medial part of the optic tectum (ot). Incident rays may also cause the pineal gland (pin) to release the hormone melatonin (mel) which stimulates melanophore aggregation. Light rays reflected from a white background (rr) stimulate the dorsal retina (d) and activate “paling” neurones (open circles, continuous lines) which pass to the latero-ventral part of the tectum. Tracts pass to the medulla oblongata (mo) where both darkening and paling centres are found. Centres in the brain also activate pathways leading to the release of darkening hormone (MSH) or paling hormone (MCH) from the pituitary gland (pit). The medullary paling centre sends excitatory tracts to motoneurones in the spinal cord (sc), which pass as pre-ganglionic fibres into the sympathetic chain (ac). The medullary darkening centre sends inhibitory tracts to the same spinal moto-neurones. In the chain, preganglionic fibres run in both cephalad and caudal directions. They have multiple collateral branches which form cholinergic synapses in the sympathetic ganglia (sg). Similar fibres innervate the chromaffin tissue of the head kidney (hk). The autonomic ganglion cells (gc) are adrenergic and send long postganglionic axons (post) in the segmental spinal nerves to reach the skin. The terminal parts of the axons branch into a ground plexus around melanophores (m). These terminals carry varicosities (v) which release catecholamines (cat) to act on a-adrenoceptors (a) and promote pigment aggregation. /3-Adrenoceptors (/3) may mediate melanophore dispersion, at least in some fish. Nucleotides and purines, released as co-factors induce pigment dispersion by way of adenosine receptors (an). Hormones (MSH, MCH, mel) as well as adrenaline and diffusing neurotransmitters, also arrive in blood vessels (bv).

finding, Grove (1969a, b) found that treatment with adrenergic neurone blockers (guanethidine, bretylium or reserpine) abolished both rapid paling and darkening in intact Phoxinus. Postoperative darkening, after in vivo lesions in the chromatic tract, is not abolished but it is greatly curtailed (from 1 day to 3 hours). Chromatic nerve terminals remain alive-for hours in isolated skin and scales, and can be activated electrically, or by drug and chemical treatment. If ATP or a similar cofactor is spontaneously released in isolated skin, this would

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explain the dispersion of melanophores which is regularly seen in such isolated preparations. Furthermore, little attempt has yet been made to search for nerves and secretory cells with other messengers (including peptides) which may affect melanophores. A schematic plan of the likely autonomic contribution to melanophore control during physiological changes in shade is presented in Figure 10.4. However, it still seems unlikely that a simple mono-neuronic model of this kind can control the elegant pattern responses which have been described for some fish (e.g. Mast, 1916). Patches of skin possessing melanophores with different sensitivity to transmitters or hormones, combined with different rates of nerve discharges, may contribute to the response of flatfish to patterned backgrounds, but this aspect of the life of lower vertebrates remains a challenging area of study.

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352 COMPARATIVE PHYSIOLOGY AND EVOLUTION Rodionov, V.I., Gyoeva, F.K. and Gelfand, V.I. (1991). Kinesin is responsible for centrifugal movement of pigment granules in melanophores. Proc. Natl. Acad. Sei. USA, 88, 4956-4960. van Rynberk, G. (1907). On the segmental skin innervation by sympathetic nervous system in vertebrates, based on experimental researches about the innervation of the pigment cells in flat fishes, and of the pilo-motor muscles in cats. Proc. Kon. Acad, van Wetensch. (Amsterdam), X, 332-342. Schaefer, J.G. (1921). Beiträge zur Physiologie des Farbenwechsels der Fische. I. Untersuchunger an Pleuronectiden. II. Weitere Untersuchungen. Pflüg. Arch. ges. Physiol., 188, 25-48. Schöndorff, A. (1903). Über den Farbenwechsel bei Forellen. Arch. f. Naturgeschichte., 69, 399-426. Scott, G.T. (1965). Physiology and pharmacology of colour change in the sand flounder, Scophthalmus aquosus. Limnol. Oceanog., 10, Supplement: A.C. Redfield 75th Anniversary Volume R230-R246. Secerov, S. (1909). Farbenwechselversuche an den Bartgrundel (Noemacheilus barbatula L). Archiv, fü r Entw. Mech. der Organismen., 28, 629-660. Semper, S. (1892). Farbwechsel an Macropoden. Arb. aus d. zoolog. Institut, (Würzburg), 10, 31-58. Spaeth, R.A. (1916). Evidence proving the melanophore to be a disguised type of muscle cell. J. Exp. Zool., 20, 193-215. Spaeth, R.A. and Barbour, H.G. (1917). The action of epinephrine and ergotoxin on single, physiologically isolated cells. J. Pharmacol. Exp. Ther., 9, 431-440. Sumner, F.B. (1933). The differing effects of different parts of the visual field upon the chromatophore responses of fishes. Biol. Bull. (Woods Hole), 65, 266-282. Svensson, S.P., Karlsson, J.O.G. and Grundström, N. (1989). Melanophores in isolated scales of Labrus berggylta (Ascanius): innervation and alpha-2-adrenoceptor-mediated pigment aggregation. Comp. Biochem. Physiol., 93C, 247-252. Trendelenburg, U. (1966). Mechanisms of supersensitivity and subsensitivity to sympathomimetic amines. Pharmac. Rev., 15, 225-276. Vilter, V. (1937). Les rapports entre les localisations rétiniennes et la polarisations dorsoventrale de la livrée mélanique chez l’axolotl. C.r. Ass. Anat., 32, 448-461. Vilter, V. (1939). Configuration des dermatomes pigmento-moteurs chez les téléosteéens et modalités de leur recouvrement réciproque. C.r. Soc. Biol. (Paris), 130, 388-390. Waring, H. (1963). Color change mechanisms o f cold-blooded vertebrates. New York: Academic Press. Watanabe, M., Izu mi, I. and Iwata, K.S. (1962). The action of adrenaline on the melanophore of Oryzias with special reference to its pigment dispersing action. Biol. J. Okayama Univ., 8, 95-102. von Wernöe, Th. B. (1928). Über den verlauf und die Verteilung präganglionärer sympatischer Bahnen bei Fischen. In Physiological papers dedicated to August Krogh. Heinemann; Copenhagen. Whitear, M. (1952). The innervation of the skin of teleost fishes. Quart. J. Microsc. Sei., 93, 289-305. Wyman, L.C. (1924). The reactions of the melanophores of embryonis and larval Fundulus to certain chemical substances. J. Exp. Biol., 40, 161-180. Yamada, K. (1980). Action of sympathetomimetic amines on leucophores in isolated scales of a teleost fish with special reference to beta-adrenoceptors mediating pigment dispersion. J. Sei. Hiroshima Univ., Ser. B. Div. 1. 28, 95-114.

11 The Iris Stefan Nilsson Department o f Zoophysiology, University o f Göteborg, Göteborg, Sweden The anatomical arrangement of the autonomic pathways that innervate the vertebrate iris is relatively uniform among the groups studied. Pathways of cranial autonomic (‘parasympathetic’) origin reach the ciliary ganglion of all groups, except possibly amphibians, along the oculomotor nerve and short ciliary root, and postganglionic neurons reach the iris in the ciliary nerves. Nerve fibres from the anterior sympathetic chain ganglia enter the ciliary nerves. The nature of the autonomic fibres controlling the iris is amazingly variable, and there are no traces of any ‘phylogenetic trends’ in the evolution of the innervation patterns. A survey of the different vertebrate groups yields examples of cholinergic, adrenergic and non-adrenergic, non-cholinergic (NANC) innervations of the sphincter and dilator muscle, that combines striated and smooth types of muscle, a- and jS-adrenoceptors and muscarinic and nicotinic cholinoceptors. In addition, the iris sphincter in many fish, amphibians and reptiles, and in the embryonic chicken, contains a light-sensitive pigment and constricts in direct response to light. KEY WORDS iris; autonomic nervous system; ciliary ganglion

INTRODUCTION The vertebrate iris works much like the aperture of a camera, adjusting the pupillary diameter to regulate the amount of light that reaches the retina. This process depends on two sets of antagonistically acting muscles: a ring of circular muscles at the pupillary margin (the sphincter pupillae), and a set of radial muscle bundles (the dilator pupillae). Contraction of the sphincter, associated with relaxation of the radial muscle causes narrowing of the pupil (miosis), while the opposite (contraction of the radial muscles and relaxation of the sphincter) opens wide the pupil (imydriasis). In most vertebrates, the iris is composed of smooth muscle but in reptiles and birds the iris muscle is (at least mainly) striated. In some species of fish, amphibians and reptiles the dilator muscle may be absent. In addition to the autonomic innervation of the iris (see later), it was observed more than 100 years ago that the sphincter pupillae of many fish, amphibians and reptiles possess a supplementary regulatory mechanism. The sphincter muscle 353

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contains a light-sensitive pigment, which causes a direct (i.e. unrelated to nerves) pupilloconstriction in response to light (Steinach, 1890, 1892; Guth, 1901). The presence of a light-sensitive pigment in the iris muscle is not surprising, in view of the origin of this tissue fro.m the embryonic retina (Romer, 1962; Hoperskaya, 1972). Mydriasis occurs in humans (and other mammals) if the excitatory innervation to the sphincter pupillae is blocked. Since this innervation is largely cholinergic, blockade can be established by atropine or related muscarinic cholinoceptor antagonists. In times passed, fashion dictated that the eyes of women should be dark and mysterious (?) and to achieve this, atropine solution could be dripped onto the cornea producing a wide-open pupil. One of the plants that has been used for extraction of atropine, Atropa belladonna, has been given a name that implies its use in the beautification of women (bella donna = beautiful woman). The iris of the albino rat was used in the first successful attempt to actually see neuronal noradrenaline, and thus histochemically demonstrate adrenergic nerve fibres. The experiments were carried out in August 1961 in the laboratory of Nils-Ake Hillarp and Bengt Falck at Lund (Owman and Bjorklund, 1978), and the technique developed employed the use of formaldehyde condensation of the monoamines. This method was known to produce fluorescent compounds in adrenomedullary chromaffin tissue (Eranko, 1952, 1955). The key to the success was the use of formaldehyde vapour rather than aqueous solution, and that the very thin, delicate iris preparation dried in air so quickly that the neuronally stored noradrenaline was prevented from diffusing and blurring the image.

Mammal

Teleost

FIGURE 11.1 Simplified and generalized summary of the anatomical arrangement of nerves in the ciliary ganglion region of mammals and teleosts. Legend: cil brev, ciliaris brevis, short ciliary nerve; cil g, ciliary ganglion; cil long, ciliaris longa, long ciliary nerve; Gass g, Gasserian ganglion; prof g, profundus ganglion; r brevis, radix brevis, short ciliary root; r long, radix longa, long ciliary root; r symp, radix sympathica, sympathetic root; symp, nerve fibres from the sympathetic chains. Ill and V refer to the oculomotor and trigeminal nerves, respectively. Redrawn with permission from Nilsson (1983).

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The iris and ciliary muscle of the eye generally receive an autonomic innervation from both spinal autonomic sources (the sympathetic chains) and cranial autonomic sources (the oculomotor nerve, III, which arises in the Edinger-Westphal nucleus of the midbrain). The anatomical arrangement of the various components of the nerve supply to the eye is relatively similar in all vertebrates. For instance, the ciliary ganglion generally receives its input from three sources: the long and short ciliary roots and a sympathetic root (Figure 11.1). The nature of the innervation of the iris sphincter and dilator (if present) does, however, show an amazing degree of heterogeneity between the different groups and species studied.

ELASMOBRANCHS The innervation of the elasmobranch iris was first studied in detail by Young (1933a), who demonstrated a direct effect of light on the iris sphincter in studies of Scyllium catulus, Scyllium (Scyliorhinus) canicula, Mustelus laevis and Trygon violaceus. The iris sphincter was insensitive to both acetylcholine and adrenaline, and apparently devoid of an autonomic innervation. Oculomotor nerve stimulation produced contraction of the radially arranged dilator muscles, probably by a cholinergic mechanism since atropine caused narrowing of the pupil. There is no evidence in elasmobranchs for a contribution to the iris innervation by ‘sympathetic’ fibres from the paravertebral ganglia (Young, 1933a). Thus, in elasmobranchs the pupillary diameter is controlled by the antagonistic actions of the sphincter, which constricts in direct response to light, and the radial dilator muscle, controlled by a cholinergic excitatory autonomic innervation running in the oculomotor nerve (Figure 11.2).

TELEOSTS A direct effect of light on the sphincter is evident in a number of teleost species (Steinach, 1890, 1892; Guth, 1901; Brown-Sequard, 1847; Nilsson, 1980), but may be absent in some (e.g. Uranoscopus scaber and Lophius piscatorius; Young, 1931, 1933b), although more recent studies suggest that the iris sphincter of Lophius is light sensitive (Rubin and Nolte, 1981). The pattern of autonomic innervation is quite varied between species: Young (1933b) demonstrated an innervation of the dilator muscles by cholinergic fibres running in the oculomotor nerve in Uranoscopus, while no such control could be seen in Lophius. A cholinergic innervation of the sphincter was demonstrated in both species, and this innervation originates in the cephalic part of the sympathetic chains (Figure 11.2; Young, 1933b). The iris of Lophius and Opsanus tau is also constricted by adrenergic agonists, but the response in these species is thought to be nonspecific (Rubin and Nolte, 1981).

356 COMPARATIVE PHYSIOLOGY AND EVOLUTION

Elasm obranch

Uranoscopus

FIGURE 11.2 Generalized summary of the patterns of autonomic innervation of the iris in some vertebrate species and groups. Legend: cil g, ciliary ganglion; a and (3, alpha and beta adrenoceptors, respectively; m, muscarinic cholinoceptors; n, nicotinic cholinoceptors; sc, sympathetic chains; sg, sympathetic ganglia. Solid lines, cholinergic neurons; dashed lines, adrenergic neurons; dotted lines, neurons of unknown nature. + and — refer to excitatory and inhibitory effects, respectively.

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FIGURE 11.3 Cod iris sphincter preparation in vitro. Effect of electrical stimulation of the ipsilateral cephalic sympathetic chain posterior to the facialis-trigeminal outflow with pulses of 10 Hz, 1 ms pulse duration and 3 V for 20 sec every 4 min, compared to the effect of illumination of the preparation for 2 min with bright light. Reproduced with permission from Nilsson (1980).

In the Atlantic cod, Gadus morhua, the effects of nerve stimulation and light are smaller than in Uranoscopus or Lophius, although easily recordable (Figure 11.3; Nilsson, 1980). Dilator muscles appear to be absent, but there is a distinct sphincter which contracts in response to both acetylcholine and adrenaline, and also in response to tyramine, an amine that produces its effects mainly by releasing stored catecholamines from adrenergic nerve endings. Destruction of adrenergic nerve terminals by injection of 6-hydroxydopamine (see Holmgren and Nilsson, 1976, 1982) abolished the response to tyramine, and produced a presynaptic super sensitivity to adrenaline (Nilsson, 1980). Electrical stimulation of either the ciliary nerve or the cephalic part of the ipsilateral sympathetic chain produced rapid constriction of the pupil, which could be blocked by the a-adrenoceptor antagonist phentolamine, but not by similar concentrations of atropine. The excitatory innervation of the sphincter in the cod is thus adrenergic, acting via an a-adrenoceptor mechanism. Pharmacological analysis of drug effects on the sphincter reveals the additional presence of ^-adrenoceptors. The adrenergic fibres to the cod iris sphincter have their cell bodies in the cephalic sympathetic chain ganglia, and histochemical evidence suggests that adrenergic cell bodies occur as far anterior as within the ciliary ganglion itself (Figure 11.2; Nilsson, 1980 and unpublished results). The control of the teleost iris thus shows a number of different arrangements. A direct constriction of the sphincter in response to light is present in many, but not all, teleosts. The innervation of the sphincter originates in the sympathetic chains, runs in the long or the short ciliary nerve to the eye, and may be either cholinergic (excitatory) or adrenergic (excitatory). The dilator muscle, if present, is innervated by excitatory, cholinergic fibres which run in the oculomotor nerve.

AM PHIBIANS The direct pupilloconstrictor effect of light on the amphibian iris sphincter is well documented (Steinach, 1890, 1892; Guth, 1901; Brown-Sequard, 1947; Barr and

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Alpern, 1963; Glaus-Most, 1969; Morris, 1976). Dilator muscles appear to be absent in both anurans (Armstrong and Bell, 1968; Morris, 1976), and the urodele, Cynops pyrrhogaster (Okamoto, 1988). The light-induced constriction of the sphincter is antagonised by a dense, mainly 0-adrenoceptor mediated, inhibitory adrenergic innervation of the sphincter. In the anurans, the pathways leave the spinal cord in the 2nd, 3rd and 4th spinal roots (Langley, 1911; Armstrong and Bell, 1968; Ber and Singer, 1969; Morris, 1976; Srivastava and Jaju, 1982; Okamoto, 1988).

REPTILES A constriction of the sphincter in response to oculomotor nerve stimulation is the general pattern in reptiles, birds and mammals, but is contrary to the situation in fish where oculomotor fibres run to the dilator muscle (if present). In reptiles, the iris sphincter consists mainly of striated muscle, and stimulation of the oculomotor nerve causes miosis. The pupil of the alligator (Alligator mississippiensis) constricts in response to the cholinoceptor agonists nicotine and pilocarpine, and the effect of oculomotor nerve stimulation is antagonised by atropine but not by curare (Iske, 1929). In the turtle (Emys blandingi) there is a pupillodilation after atropine, which further supports the idea of a muscarinic-type cholinoceptor mechanism in the sphincter (Iske, 1929). Whether this is due to a special type of atropine-sensitive cholinoceptors in the striated muscle, or if the innervation mainly affects smooth muscle elements in the sphincter, is not clear. Dilator muscles are absent in the alligator iris, but old studies in turtles describe a slight dilation of the pupil in response to sympathetic chain stimulation (Gaskell and Gadow, 1884; Mills, 1885). The nature of this response remains unknown: it could either be an excitatory innervation of dilator muscle, or an inhibitory effect on the sphincter (Figure 11.2).

BIRDS The developing chick iris is sensitive to light, but this light response disappears in the adult (Pilar et al.y 1987). Similar to the situation in reptiles, the avian iris muscle is chiefly of the striated variety, and oculomotor fibres innervate the iris sphincter. The cell bodies of the postganglionic neurons of these pathways lie within the ciliary ganglion, and the neurotransmission includes both ‘traditional’ chemical synapses, and a type of electrical junction (Marwitt, Pilar and Weakly, 1971; Gabella, 1976). At least two distinct types of postganglionic neurons are present in the avian ciliary ganglion. One type innervates smooth muscles of the choroid coat and the vasculature of the choroid layer, and the neurons contain somatostatin as a cotransmitter of acetylcholine. The second type of neuron is cholinergic, with no known cotransmitter, and innervates the iris and ciliary body (Meriney et al.9 1987; Gray, Pilar and Ford, 1989; Coulombe and Nishi, 1991). During development of the embryo, the somatostatin-immunoreactivity of the neurons is stimulated by the

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presence of a substance released from choroid cells (Coulombe and Nishi, 1991). Contrary to the observations in the alligator, most of the effects on the bird iris caused by electrical stimulation of the oculomotor nerve can be abolished by curare, but not by atropine. The nerve endings possess motor end plates of the type seen in skeletal striated muscle, rather than the more widespreading varicose terminals of autonomic neurons (Iske, 1929; Campbell and Smith, 1962; Pilar and Vaughan, 1969; Bennett, 1974). It is not clear to what extent contributions from the sympathetic chain reach the avian iris, but in old studies an innervation of the striated dilator muscle by fibres from the trigeminal nerve has been postulated (Zeglinski, 1885; Jegorow, 1887, cf. Koppanyi and Sun, 1926). A functional equivalent of the iris dilator muscle is found in the anterior epithelial layer of the avian iris, which includes both circular smooth muscle within the sphincter, and radially arranged smooth muscle innervated by both cholinergic and adrenergic fibres (Ehinger, 1967; Nishida and Sears, 1970; Dietrich, Fischer and Hiller, 1988). In the chick embryo, the myoepithelial smooth muscle develops first, and is functionally innervated by cholinergic fibres exerting muscarinic effects. Later the striated muscle of the sphincter occurs, and receives an innervation which acts via nicotinic cholinoceptors (Pilar et al., 1987). Both types of cholinoceptors occur also in the dilator muscle, which is composed of both striated and smooth muscle elements (Pilar et al., 1987).

MAMMALS The mammalian iris sphincter is controlled mainly by excitatory cholinergic fibres (see earlier), and by adrenergic fibres from the superior cervical ganglion that produce dilation of the sphincter via both a- and ^-adrenoceptor mechanisms (van Alphen, Robinette and Marci, 1964; Ehinger and Falck, 1965, 1966; Tranzer and Thoenen, 1967; Ehinger, Falck and Persson, 1968; Ochi et al., 1968; Persson and Sonmark, 1971). The iris dilator of the rat contracts in response to catecholamines, which can be antagonized by the a-adrenoceptor antagonist phentolamine. Low concentrations of acetylcholine produced relaxation, while higher concentrations contract the preparations; both responses are antagonized by atropine. Electrical stimulation produces mixtures of contractions and relaxation: the contraction is adrenergic, and there is an additional cholinergic inhibitory component (Narita and Watanabe, 1982). Of the neuropeptides that have been demonstrated in autonomic neurons of mammals, the distribution and possible functions of substance P has received some interest. Substance P-immunoreactive nerve terminals occur both in the iris sphincter and near the dilator muscle of the rabbit, and the peptide produces strong contractions of the isolated sphincter in vitro (Leander et al., 1981; Ueda et al., 1981; Tornqvist et al., 1982). Capsaicin, a drug known to activate substance P-containing neurons, mimics the response to substance P, and it was therefore suggested that

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substance P serves as an additional excitatory neurotransmitter in the rabbit iris sphincter (Ueda et al., 1981).

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Leander, S., Häkansson, R., Rosell, S., Folkers, K., Sundler, F. and Törnqvist, K. (1981). A specific substance P antagonist blocks smooth muscle contractions induced by non-cholinergic, nonadrenergic nerve stimulation. Nature., 294, 467-469. Marwitt, R., Pilar, G. and Weakly, J.N. (1971). Characterization of two ganglion cell populations in avian ciliary ganglia. Brain Res., 25, 317-334. Meriney, S.D., Pilar, G., Ogawa, M. and Nunez, R. (1987). Differential neuronal survival in the avian ciliary ganglion after chronic acetylcholine receptor blockade. J. Neurosci., 7, 3840-3849. Mills, T.W. (1885). The innervation of the heart of the slider terrapin (Pseudemys rugosa). J. Physiol. (Lond.), 6, 248-286. Morris, J.L. (1976). Motor innervation of the toad iris bufo-marinus. Am. J. Physiol., 231, 1272-1278. Narita, S. and Watanabe, M. (1982). Response of isolated rat iris dilator to adrenergic and cholinergic agents and electrical stimulation. Life Sei., 30, 1211-1218. Nilsson, S. (1980). Sympathetic nervous control of the iris sphincter of the Atlantic cod Gadus morhua. J. Comp. Physiol., 138, 149-156. Nilsson, S. (1983). Autonomic Nerve Function in the Vertebrates, Berlin, Heidelberg, New York: Springer-Verlag. Nishida, S. and Sears, M. (1970). Fine structure of the anterior epithelial cell layer of the iris of the hen. Exp. Eye Res., 9, 241-245. Ochi, J., Konishi, M., Yoshikawa, H. and Sano, Y. (1968). Fluorescence and electron microscopic evidence for the dual innervation of the iris sphincter muscle of the rabbit. Z. Zellforsch., 91,90-95. Okamoto, M. (1988). Fine structure of the iris muscle in the Japanese common newt Cynops pyrrhogaster with special reference to innervation. Zool. Sei. (Tokyo), 5, 337-346. Owman, C. and Björklund, A. (1978). Current research on the histochemistry and function of biogenic amines. A tribute to Bengt Falck. Acta Physiol. Scand., 102 (Suppl. 452), 5-8. Persson, H. and Sonmark, B. (1971). Adrenoceptors and cholinoceptors in the rabbit iris. Eur. J. Pharmacol., 15, 240-244. Pilar, G. and Vaughan, P.C. (1969). Electrophysiological investigations of the pigeon iris neuromuscular junctions. Comp. Biochem. Physiol., 29, 51-72. Pilar, G., Nunez, R., McLennan, I.S. and Meriney, S.D. (1987). Muscarinic and nicotinic synaptic activation of the developing chicken iris. J. Neurosci., 1, 3813-3826. Römer, A.S. (1962). The vertebrate body, Philadelphia, London: Saunders Company. Rubin, L.R. and Nolte, J. (1981). Autonomic innervation and photo sensitivity of the sphincter pupillae muscle of 2 teleosts Lophius piscatorius and Opsanus tau. Curr. Eye. Res., 1, 543-552. Srivastava, Y.P. and Jaju, B.P. (1982). Characterization of adrenoceptors of frog Rana tigrina iris. Indian J. Exp. Biol., 20, 612-614. Steinach, E. (1890). Untersuchungen zur vergleichenden Physiologie der Iris. Erste Mitteilung. Pflügers Arch. Physiol., 47, 289-340. Steinach, E. (1892). Untersuchungen zur vergleichenden Physiologi der Iris. Zweite Mitteilung. Pflügers Arch. Physiol., 52, 495-525. Törnqvist, K., Mandahl, A., Leander, S., Lor6n, I., Häkanson, R. and Sundler, F. (1982). Substance P immunoreactive nerve fibers in the anterior segment of the rabbit eye: Distribution and possible physiological significance. Cell Tissue Res., 222, 467-478. Tranzer, J.P. and Thoenen, H. (1967). Electronmicroscopic localization of 5-hydroxytyramine (3, 4, 5-trihydroxyphenyl-ethylamine) a new “false” sympathetic transmitter. Experientia, 23, 743-745. Ueda, N., Muramatsu, I., Sakakibara, Y. and Fujiwara, M. (1981). Noncholinergic, nonadrenergic contraction and substance P in rabbit iris sphincter muscle. Jap. J. Pharmacol., 31, 1071-1080. Young, J.Z. (1931). The pupillary mechanism of the teleostean fish Uranoscopus scaber. Proc. R. Soc. Lond. [Biol.], 107, 464-485. Young, J.Z. (1933a). Comparative studies on the physiology of the iris.-I. Selachians. Proc. R. Soc. Lond. [Biol.], 112, 228-241. Young, J.Z. (1933b). Comparative studies on the physiology of the iris.-II. Uranoscopus and Lophius. Proc. R. Soc. Lond. [Biol.], 112, 242-249. Zeglinski, N. (1885). Experimentelle Untersuchungen über die Irisbewegung. Arch. Anat. Physiol., 1-37.

Index accomodation, 11 acetylcholine (ACh), 70 effects on colour change, 344 effects on glands, 172, 174, 178-179, 184-185, 187 effects on the iris, 357-359 effects on the gut, 121, 123, 125, 131, 135, 142, 144 effects on the heart, 197-198, 204, 206-207 effects on the urino-genital system, 317, 321, 323-324 effects on the vasculature, 220-221, 223 ACh (see acetylcholine) acetylcholinesterase, 212, 214, 217, 219, 293 AChE (see acetylcholinesterase) acid secretion (see secretion) ACTH ( = adenocorticotropic hormone), 179 adenosine, 73, 127, 199 adenosine 5 '-triphosphate (ATP), 73, 119, 127, 199, 203, 222, 264, 276, 346 a , 0-methylene ATP, 203 adenyl compounds, 73 adipose tissue, 32, 52, 101, 103 adrenal gland ( = adrenal; see also chromaffin tissue), 6, 31, 47-48 adrenaline, 71 effects on colour change, 344-346 effects on glands, 176, 184 effects on the iris, 355-359 effects on the gut, 121 effects on the heart, 197-206 effects on the lung, 274, 284 effects on the swimbladder, 295, 297 effects on the urino-genital system, 312, 315 effects on the vasculature, 203, 279 in chromaffin tissue, 38

adrenergic neurons, 71, 180, 183-184, 198, 263, 270, 276, 313 adrenoceptors, a-(alpha-) in colour change, 341, 345 in glands, 184 in the gut, 122-123 in the iris, 356 in the lung, 274, 279 in the melanophores, 341 in the spleen, 248, 251, 253 in the swimbladder, 295, 297 in the urino-genital system, 323 in the vasculature, 219, 220, 222, 279 adrenoceptors, 0-(beta-) in colour change, 341, 345 in glands, 184 in the iris, 356 in the gut, 122-123 in the heart, 197, 198, 199, 203-205, 208 in the lung, 279 in the swimbladder, 295, 297 in the urino-genital system, 318, 319, 325 in vasculature, 216, 219, 220, 223, 279 afferent gill arteries, 212 air breathing, 194 airways, 18, 32, 262 alytesin, 76 amphibians, 101 anatomy o f the autonomic nervous system, 8-9, 11, 13, 21, 23, 25, 27-30, 43-47, 52-54 innervation o f the vasculature, 213-216 innervation o f glands, 171-172, 174, 176-178 innervation o f the gut, 125-126, 128-129, 134-135, 137, 139, 141-142, 144, 152-154 innervation o f the heart, 200-203 363

364

COMPARATIVE PHYSIOLOGY AND EVOLUTION

innervation o f the iris, 357 innervation o f the lung, 271, 273-281 innervation o f the spleen, 249 innervation o f the urino-genital organs, 314, 318, 323 amylase, 180, 182-184 antidiuresis, 314, 316 anuran amphibians (see amphibians) aortic baroreceptors, 224 apnea, 272 apodan amphibians (see amphibians) APPs ( = avian pancreatic polypeptides), 79 APY ( = anglerfish peptide YG), 81 arginine vasotocin, 314, 316 aromatic amino acid decarboxylase (see DOPA decarboxylase) ATP (see adenosine 5 '-triphosphate) atropine, 174, 178, 197, 317-318, 322-323 Auerbach’s plexus (see myenteric plexus) Australian lungfish, 200 axillary body, 36, 38 baroreceptor, 224-225 bat, 207 Bidder’s ganglion, 27 binucleate neurons, 5 bird anatomy o f the autonomic nervous system, 11, 25-29, 47, 50 innervation o f the lung, 269 innervation o f the iris,. 358 innervation o f glands, 185 innervation o f the gut, 139, 141-143, 153 innervation o f the vasculature, 219-221 innervation o f glands, 171-174, 180, 183-185, 187 innervation o f the heart, 205 innervation o f the iris, 358-359 innervation o f the lung, 269-271 innervation o f the urino-genital organs, 324 bladder (see swimbladder and urinary bladder, respectively) bombesin, 75-76, 124, 127, 170-180, 185, 187, 197, 210, 278 brachial plexus, 31, 35, 37, 40-41, 43, 47-49, 51, 54 brachiopterygian fish (see also holosteans), 126, 128, 141, 146, 152 branchial arches, 9, 13, 29 ganglia, 20

hearts, 195 nerves, 30, 209, 212 rami o f the vagus, 30, 39 vasculature, 20, 209-210, 213 bretylium, 203, 216, 227, 313 Brockmann bodies, 179-180, 182-184 bulbus cordis, 195 buoyancy regulation, 290 BW 284 C51, 254 Caecilia (see amphibians) caerulein, 77-78, 174, 177, 182 calcitonin gene-related peptide ( = CGRP), 88, 145, 156, 171, 180, 187, 206-207, 209, 268, 280 capsaicin, 263, 268 carassin, 85-86 carbachol, 174, 177-178, 199 cardiac nerves, 20, 27, 29-33, 39, 44, 49, 205 cardiac ganglion, 18, 49, 205, 207 cardiac gland, 179 cardiovascular reflexes, 224 carotid artery baroreceptors, 224-225, 227 carotid labyrinth, 225-227 carotid nerves, 32 catecholamines (see also adrenaline and noradrenaline) in SIF cells, 108 in chromaffin tissue (Ch. 3), 97 ff. in colour change, 341 in glands, 171-172, 178, 184, 187 in the iris, 355-359 in the gut, 121 in the heart, 197, 203, 205 in the lung, 274, 284 in the swimbladder, 295, 297 in the urino-genital system, 314, 319, 324 in the vasculature, 210, 219, 222, 227 in endothelial cells, 200 caudal heart, 195 cavernous plexus, 17-18 CCK (see cholecystokinin) cephalic vasculature, 221 cephalic phase, 173 cerebral vasculature, 17-19, 29, 214, 220 cervical ganglia, 49-51, 54 cervical sympathetic trunk, 31, 48, 50 CGRP (see calcitonin gene-related peptide) ChAT (see choline acetyltransferase) chelonians, 8, 11, 13, 23, 48 chemical codes, 2-3 chemoreceptor reflexes, 107, 225-226

INDEX

chief (peptic) cells, 176 cholecystokinin, ( = CCK), 76, 155, 171-180 choline acetyltransferase ( = ChAT), 70, 184, 254, 295 cholinergic neurons, 11, 16, 19-20, 27, 32, 50, 70, 119, 121, 125, 145-148, 180, 210, 212, 221, 226, 263, 276 supersensitivity, 254 sympathetic neurons, 42, 52, 223 cardiac tonus, 199 cholinesterase (see acetylcholinesterase) chondrostean, 19, 32, 42-43 chorda tympani, 13-15, 17-18, 24-25, 29 choroid, 12, 358 choroidal vasculature, 11 chromaffin systems (Chromaffin Systems Ch. 3), 97 ff. chromaffin cells (chromaffin tissue), 5-6, 18-19, 27, 31-36, 38, 42-44, 47-48, 52-54, 72, 98, 104, 196, 218 chromatic tracts, 336, 337 chromatophores (Chromatophores Ch. 10), 42, 48, 331 ff. ciliary ganglion, 9-12, 14, 19, 22, 24, 39, 354 ciliary muscles, 11-12 ciliary nerves, 28, 38-39, 44, 49, 354, 357 cocaine, in presynaptic super sensitivity, 251 cod (see Gadus morhua) coeliac artery, 213 coeliac ganglion, 32, 39, 41-42, 44-46 coeliac plexus, 48, 50-51 coeliaco-mesenteric plexus, 34, 50 colour change (see Chromatophores, Ch. 9), 331 ff. conduction velocities, 6, 44-45 conus arteriosus, 195 coronary vessels, 197 coronary vasculature, 210, 217, 221, 223 corpus cavernosum, 223 corticotropin-releasing factor ( = CRF), 178 cranial autonomic system, 2, 19-10, 14-15, 17, 24, 38-40, 43-44, 48, 210, 214 craniocervical sympathetic trunk, 49 crocodiles, 23, 25, 28, 49-50 cyclic AMP, 346 cyclostomes anatomy o f the autonomic nervous system, 9, 13, 19, 34, 43, 52-55 innervation o f glands, 179-180 innervation o f the gut, 125-126, 128-129

365

innervation o f the vasculature, 209 innervation o f the heart, 196 innervation o f the urino-genital organs, 312 darkening area in colour change, 336 darkening neurones, 347 DBH (see dopamine-p-hydroxylase) dendrites, 3-8, 12 desmethylimipramine ( = DMI), 251 developmental studies, 52 dihydroergotamine, 197 dilator pupillae, 353 dipnoans ( = lungfish), 199 anatomy o f the autonomic nervous system, 19, 21, 43, 53 innervation o f the gut, 126, 128-129, 135, 139, 141-142, 144, 146 innervation o f the lung, 258, 288 diving, 205, 220, 227 dogfish Xsee elasmobranchs) DOPA decarboxylase ( = aromatic amino acid decarboxylase), 71 dopamine, 203 dopamine-j3-hydroxylase ( = DBH), 12, 16, 71, 170, 183, 184, 187 ducts o f Cuvier, 20 duodenum, 18, 27 dynorphin, 82, 206, 208, 223 eel intestinal peptide ( = EIPP), 88 elasmobranchs anatomy o f the autonomic nervous system, 9, 11-13, 19-21, 28, 35-37, 42-43, 52-55 innervation o f the vasculature, 209 innervation o f glands, 172, 174, 176-177, 179-180, 182-187 innervation o f the gut, 125-126, 128-129, 134-135, 137-138, 144-146, 151-153 innervation o f the heart, 197 innervation o f the iris, 355 innervation o f the spleen, 248 innervation o f the urino-genital organs, 312, 316, 321 electrical transmission, 11 eledoisin, 85, 174 endorphin, 82, 142 endothelial-derived relaxing factor (see also nitric oxide), 227 enkephalin, 16, 82, 141, 170-171, 182, 187, 206, 296 enteric neurons, 2, 7, 30, 33, 120, 209

366

COMPARATIVE PHYSIOLOGY AND EVOLUTION

enteroglucagon, 171-172 epicardium, 202 epididymal duct, 324 epinephrine (see adrenaline) ergot alkaloids, 203 erythrophore, 332 ethmoid ganglion, 22-25, 28 exercise, 212, 223, 227-228 exocrine glands, 21, 32, 180 exocytosis, 107 extra-adrenal chromaffin cells, 98, 109, 217, 319 facial nerve, 9, 10, 12-13, 15-16, 18-20, 22-24, 29, 39, 47-48, 219 facial veins, 217, 221-223 facial-trigeminal, 40 femoral artery, 220 filamental sphincters, 212 fish (see cyclostomes, dipnoans, elasmobranchs and teleosts, respectively) FMRF, 180 foramen Panizzae, 196 Frank-Starling relationship, 225 frog (see amphibians) GABA ( = 7 -aminobutyric acid), 156 galanin, 23, 25, 27, 35, 44, 88, 137, 155-156, 172, 179, 182, 187, 200-202, 204, 208, 213-217, 222, 319 gallbladder, 137, 155, 179, 212 ganglia, 27, 37, 39 ganoid fish, 13,43 gas bladders (see swimbladders) gas gland, 290-294 gastric ganglia, 36-38 gastric inhibitory peptide ( = GIP), 171-173, 178 gastrin/CCK ( = gastrin/cholecystokinin), 76, 134, 170-175, 177-182, 185, 187 gastrin releasing peptide ( = GRP), 75-76, 129, 170, 176, 178, 180, 185 gastrointestinal canal (The Gastrointestinal Canal C h.4), 119 ff. anatomy o f the autonomic nervous system, 19-20 GFR (see glomerular filtration rate) GIP (see gastric inhibitory peptide) gill, 19, 212 giraffes, 230 gizzard, 8 glands (Glands Ch. 5), 169 ff.

arrangement o f autonomic innervation, 23-24 glomerular filtration rate (GFR), 312-315 glomus cells, 226 glossopharyngeal nerve, 9-10, 13, 16-20, 17, 24-25, 28-29, 38-41, 47-48, 50, 213, 217, 221 glucagon, 171-172, 179-180, 182, 184 gonads (see Urinogenital Organs Ch. 9), 311 ff. greater superficial petrosal nerve, 14, 16, 28 guanophore, 332 gut (see The Gastrointestinal Canal, C h.4), 119 ff. hagfish, 35, 312 hair-follicles, 16 hard palate, 28 Harderian glands, 16, 21, 24 head kidney, 42 heart (see The Circulatory System Ch. 6), 193 ff. arrangement o f the autonomic innervation, 18-19, 27, 32, 38, 46, 48, 54 innervation in amphibians, 200-203 birds, 205 dipnoans, 199-200 cyclostomes, 196-197 elasmobranchs, 197-198 mammals, 205-208 reptiles, 204-205 teleosts, 198-199 helodermin, 87 helospectins, 87 hibernating mammal, 207 histamine, 170, 172, 174-175, 177, 179, 324 Hogben melanophore index, 332 holocephalan elasmobranchs, 11, 126, 128, 135, 152, 180, 182-183 holosteans, 19, 38, 43, 54, 142, 153, 172 5-HT, see 5-hydroxytryptamine 5hydroxytryptamine ( = 5-HT, serotonin), 72, 73 in glands, 170-171, 175, 177, 179, 187 in the gut, 9, 119, 125, 147, 158 in the urinogenital organs, 34, 318, 323 in the heart, 203 in the lung, 277 in the vasculature, 210, 212 6hydroxydopamine, 251, 254

INDEX

367

hylambatin, 85 hyomandibular ramus o f the facial nerve, 10, 13, 20, 25, 39 hyperaemia, 227 hypogastric plexus, 32-33 hypoglossal nerve, 25 hypoxic bradycardia, 212, 225-226

innervation in amphibians, 273-281, 285-288 birds, 269-271 fish, 288-290 mammals, 261-269 reptiles, 281-288 lungfish (see dipnoans)

iliac plexus, 44 inferior mesenteric ganglion, 32-34 inferior pharyngeal ramus, 30 inferior vena cava, 220-221 infra-orbital ganglion and nerve, 13, 22, 28 infundibulum, 324 insulin, 174, 179-180, 182-184 interatrial septum, 27 intermaxillary gland, 21 intermedin, 331 intermesenteric plexus, 32 interneurons, 5-8, 27, 203 intestine (see The Gastrointestinal Canal C h.4), 119 ff. intracardiac neurons, 200-202, 205 intralingual neurons, 18, 25 intrapulmonary vasculature, 217 iridiophore, 332 iris (Iris Ch. 11), 353 ff. arrangement o f the autonomic innervation, 11-12

major pelvic ganglion, 33 mammals adrenal gland, 103 anatomy o f the autonomic nervous system, 7, 11-12, 18, 25, 28, 30, 54-55 innervation o f the lung, 261-269 innervation o f glands, 172, 176 innervation o f the heart, 205-208 innervation o f the iris, 359 innervation o f the spleen, 250 innervation o f the vasculature, 221-223 mandibular (third) division o f the tri-geminal nerve, 16, 17, 24 marsupials, 31, 208, 222 maxillary division o f the trigeminal nerve, 13, 16, 22, 24 MCH (see melanophore concentrating hormone) MDH (see melanophore dispersing hormone) mediastinal ganglia, 32, 208 medulla oblongata in colour change, 335 Meissner’s plexus (see submucous plexus) melanocyte stimulating hormone ( = MSH), 331, 347 melanophore (see Chromatophores Ch. 10), 331 ff. melanophore concentrating hormone ( = MCH), 338, 345, 347 melanophore dispersing hormone ( = MDH), 331 melanosome, 332 melatonin, 347 mesenteric artery, 210, 213, 216 mesenteric nerves, 42 met-enkephalin (see enkephalin) microganglia, 14, 16-18, 20, 23, 29, 32, 209 miosis, 353 monotreme, 7, 31 motilin, 171, 179 motor neurons, 8 MSH (see melanocyte stimulating hormone) mucus and poison glands in the skin, 45

Jacobson’s anastomosis, 13 juxtaglomerular cells, 314-315 kassinin, 85-86 kidney (see Urinogenital Organs Ch. 9), 311 ff. arrangement o f the autonomic innervation, 18, 34, 37 lacrimal gland, 16 lampetroids (see Cyclostomes) lepidosaurs, 28 lesser petrosal nerve, 15-17 leucophore, 332 leu-enkephalin (see enkephalin) light-sensitive pigment, 354 lingual nerve, 16-17, 25, 213, 217 litorin, 76 lizards (see reptiles) lung (see Lungs and Swimbladders Ch. 8), 257 ff. arrangement o f the autonomic innervation, 21, 26-27, 29-30, 43, 46, 54

368

COMPARATIVE PHYSIOLOGY AND EVOLUTION

multinucleate neurons, 6 multipolar neurons, 9 muscularis mucosae o f the swimbladder, 295 mydriasis, 353 myenteric plexus ( = Auerbach’s plexus), 4, 7, 8, 18, 120 myxinoids (see cyclostomes) NANC ( = Non-Adrenergic, Non-Cholinergic), 214, 263, 270, 318, 320, 323-324 nasal mucosa, 16, 24, 222 nerve of the pterygoid canal (see Vidian nerve) neural crest cells, 7, 52, 55 neurogenic vasodilatation, 220 neurokinins, 85, 206, 268 neuromedins, 75-76, 85 neuropeptide gamma ( = N Py), 85 neuropeptide Y ( = NPY), 5, 16, 23, 44, 49, 79, 137, 183, 198, 203-208, 216-217, 220-223, 251, 268, 276, 279 neuropeptide K, 85, 268 neuropeptides (see Comparative Aspects o f the Biochemical Identity of Neuro-Transmitters o f Autonomic Neurons Ch. 2) in adrenomedullary cells, 103 neurotensin, 81, 133, 138, 171-172, 178, 180, 296 nitric oxide ( = NO), 221, 223, 264, 278, 298 nitric oxide synthase ( = NO synthase), 265, 275 NKA (see neurokinins) non-adrenergic, non-cholinergic (see NANC) non-adrenergic vasoconstriction, 216 non-cholinergic vasodilatation, 223 non-cholinergic mechanisms, 202, 220 non-noradrenergic sympathetic neurons, 32 noradrenaline (see also adrenaline), 71 effects on glands, 177-178, 184 effects on the gut, 121-122 effects on the lung, 293 effects on the urino-genital system, 315 effects on the vasculature, 217, 222 noradrenergic, 49, 121 norepinephrine (see noradrenaline) NPY (see neuropeptide Y)

oculomotor nerve, 9, 11-12, 22, 24, 39, 354-355 oesophagus (see The Gastrointestinal Canal C h.4), 119ff. arrangement o f the autonomic innervation, 18, 26, 27, 30 ophthalmic division o f the trigeminal nerve, 10-11, 13, 22, 24 opioid peptides, 82, 141, 178, 182, 208,

222

organ o f Zuckerkandl, 109 otic ganglion, 14, 16-17, 29 oval o f the swimbladder, 292 ovary (see Urino-Gential organs Ch. 9), 311 ff. oviduct, 324 oviposition, 324 oxygen receptors, 226 oxynticopeptic cells, 176, 187 oxytocin, 88, 183 palate, 13, 21 palatine ganglion, 22-24, 28 palatine ramus o f the facial nerve, 10, 12-13, 19, 21-24, 39-40 palatine mucosa, 21, 25 paling area in colour change, 336 paling neurones, 347 pancreas, 18, 21, 28, 177, 179-180, 182-184 pancreatic polypeptides, 79, 137, 172, 178-179, 184 parabronchi, 269 paracervical ganglion, 33 paraganglia, 6, 104 parasympathetic division o f the autonomic nervous system (see Comparative Anatomy and Evolution o f the Autonomic Nervous System Ch. 1), 1 paravertebral sympathetic chains, 43, 47 paravertebral ganglia, 31, 47 parietal (oxyntic) cells, 172 parotid salivary gland, 17, 29 pelvic ganglion and plexus, 2, 32-34, 37, 42, 44, 46-48, 50, 54-55, 223 pelvic vascular neurons, 223 pelvic ganglia - Remak’s nerve, 50 penis, 33 penna-motor muscles, 50 pepsin, 172, 174, 176-177, 187 pepsinogen, 176-177 peptide YY ( = PYY), 79, 137 peptide histidine isoleucin ( = PHI), 87, 183, 221, 264

INDEX

peptide histidine methionine ( = PHM, human PHI), 264 pharynx, 17, 19 phenylethanolamine-N-methyl transferase ( = PNMT), 71, 102-103, 183 phyllocaerulein, 77-78 phyllomedusin, 85 physalaemin, 85-86, 174 physoclist, 258, 290 physostome, 258, 290 pigment aggregation, 347 pigment cells, 40, 42 pilo-erector nerves, 32 pneumatic duct, 291 PNMT (see phenylethanolamine-N-methyl transferase) polynergic neuron, 254 portal heart, 195 post-branchial plexus, 37 postganglionic neurons, 3, 6, 34, 200,

210

post-trematic branchial ramus, 20, 25 PP (see pancreatic polypeptide) pre- and postbranchial arterial blood pressure, 226 pre- and post-trematic branchial rami, 20 pre-aortic plexus, 32-33 precapillary arterioles in skeletal muscle beds, 223 preganglionic neurons, 6, 11, 17-18, 24, 33-34, 37-38, 40, 45-48, 53, 107 pregastric ganglia, 37 preglomerular sphincters, 313-314 pre-orbital ganglion, 15, 22-24 preprocaerulein, 78 preprocholecystokinin, 78 preproenkephalin A and B, 83 preprogastrin, 78 preprosomatostatin, 84 preproVIP, 87 presacral plexus, 32 presynaptic supersensitivity, 251 pre-trematic rami, 20 prevertebral sympathetic ganglia, 31-32, 37-38, 41, 46, 48-50, 53-57 primary and secondary sympathetic chains, 53 profundus nerve, 11 pro-opiocortin, 83 prostaglandins, 324 proximal colon, 18 pseudobranch, 12 pterygopalatine ganglion, 16, 24 pulmocutaneous artery, 214, 225, 279

369

pulmonary (see also lung) vasculature, 213-214, 216-217, 220, 262, 270 nerves, 27, 30, 48 pupil, 11, 353 purinergic nerves, 73, 265 purines, 127 purinoceptors, 197 ramus ophthalmicus profundus, 11 ranatensin, 76 rays (see elasmobranchs) rectal gland, 185 rectin, 185 rectum (see The Gastrointestinal Canal C h.4), 119 reflexes, cardiovascular, 224 Remak’s ganglion, 27 Remak’s nerve, 50, 55 renal nerves (see also Urino-Genital Organs C h.9), 311 ff. renal ganglion, 32, 45-46 renal vasculature, 216 renin, 315 reptiles adrenal gland, 102 anatomy o f the autonomic nervous system, 11, 13, 15, 22-23, 25, 27-29, 47, 49-50, 54 innervation o f the vasculature, 217-219 innervation o f glands, 172, 174, 177, 183-184 innervation o f the gut, 128-129, 134-135, 137-139, 141-142, 144, 153-154 innervation o f the heart, 204 innervation o f the iris, 358 innervation o f the lung, 281-285 innervation o f the spleen, 00 innervation o f the urino-genital organs, 319, 324 reptiles, mammal-like, 29 retro-orbital plexus, 28 retroperitoneal chromaffin tissue, 109 sacral parasympathetic outflow, 7, 47 salivary glands o f lizards, 21, 23, 25, 32, 179 salt secreting glands, 184-185, 187 satellite ganglion, 42 SCG (see superior cervical ganglion) scyliorhinin, 85 sebaceous glands, 16 secretin, 171-172, 176, 179-180, 185 secretion (see Glands Ch. 5), 169 ff.

370

COMPARATIVE PHYSIOLOGY AND EVOLUTION

o f alkali, 178, 187 basal, 172-173 o f bicarbonate, 177 o f gas in swimbladder, 291 o f gastric acid, 18, 173-176, 187 o f mucus, 177, 179, 270 secretomotor neurons, 6, 13, 16-17, 24-25, 32, 46 selachians (see elasmobranchs) sensorimotor innervation, 280, 285 sensory neurons, 7-9, 200, 209, 214 sequences, peptide, 74 bombesin-like, 76 CCK/gastrin, 77 N PY /PY Y /PP-like, 80 neurotensin-like, 82 somatostatin-like, 83 tachykinins, 85 VIP-like, 87 serotonergic (see 5-hydroxytryptamine) serotonin (see 5-hydroxytryptamine) sharks (see elasmobranchs) SIF cells ( = small intensely fluorescent cells), 100, 106, 319-320, 324 sinoatrial nerve plexus, 198 sinus venosus, 20, 27, 195 skates (see elasmobranchs) sublingual gland ganglion, 14 small intensely fluorescent cells (see SIF cells) small intrinsic neurons, 203 submandibular ganglion, 14, 24 snakes (see reptiles) somatostatin, 74, 83 in glands, 170-173, 175-177, 179-180, 182, 184-185, 187 in the iris, 358 in the gut, 144, 158 in the heart, 200-204, 206, 207 in the lung, 273, 276 in the urino-genital system, 318-319 in the vasculature, 210, 214 SP (see substance P) specialized adrenoceptors, 203, 222 sphenopalatine ganglion, 5, 12, 14, 16, 24, 25, 28 sphincter pupillae, 353 spinal autonomic outflow, 2, 31, 55 spinal sympathetic ganglia, 40, 49-50 splanchnic nerves, 21, 32, 36-38, 42, 44, 46, 50, 104, 107, 121, 125, 127, 248, 251 spleen (see The Spleen Ch. 7), 247 ff. stellate ganglion, 31, 33, 49, 54, 201, 204, 208

stomach (see The Gastrointestinal Canal Ch. 4), 119 ff. stress, effects on the circulatory system, 227 structure o f autonomic neurons, 3 subclavian plexus, 43-44 subcutaneous nervous plexus, 35 sublingual gland, 17, 29 submandibular ganglion, 16, 25, 29, 17 subraucous plexus ( = Meissner’s plexus), 7, 120 substance P ( = SP), 73, 85 in glands, 170-171, 176, 179, 183, 187 in the iris, 359 in the gut, 124, 126, 135, 142-143, 145, 158 in the heart, 206-207 in the lung, 268, 276, 280 in the spleen, 251 in the swimbladder, 296 in the urino-genital system, 318-319 in the vasculature, 209, 213 sulphated cholecystokinin 8 (see gastrin/CCK) superior mesenteric ganglion, 32 superior cervical ganglia ( = SCG), 14, 31, 45 supersensitivity, 254, 345 supra-orbital salt gland, 24 supra-renal bodies, 37, 38 supreme cervical ganglion, 50-51 sustentacular cells, 226 sweat glands, 32, 52 swimbladder ganglion, 21, 293 swimbladders (see Lungs and Swimbladders Ch. 9), 257-258, 290 sympathetic chain, 31, 37, 40-41, 44-45, 48-49, 51 sympathetic chain in colour change, 337 sympathetic division o f the autonomic nervous system (see Comparative Anatomy and Evolution o f the Autonomic Nervous System Ch. 1), 1 tachykinins, 84, 145, 172, 174, 177, 187, 209, 268 teleosts anatomy o f the autonomic nervous system, 11, 13, 19, 21, 30, 38-39, 42, 52-55 innervation o f the vasculature, 210-212 innervation o f glands, 171-173, 174, 177, 179-180, 182-183, 187

INDEX

innervation o f the gut, 125-126, 128-129, 134-135, 137-139, 141-142, 144-146, 152-154 innervation o f the heart, 198 innervation o f the iris, 355 innervation o f the spleen, 248 innervation o f the urino-genital organs, 313, 316, 321 terminal arterioles, 313, 314 testis (see Urinogenital Organs Ch. 9), 311 thermoregulation, 221, 230 thyroid gland, 18 thyrotropin-releasing hormone ( = TRH), 178 toad (see amphibians) tongue, 17, 219 trachea (see Lungs and Swimbladders Ch. 8), 257 tractus tecto-bulbaris, 335 transverse commissure, 40 trigeminal nerve, 13, 22, 24, 38-39, 47-48, 354 truncus arteriosus, 217 turtles (se reptiles) tympanic ramus o f the glossopharyngeal nerve, 14-17 tyramine, 197, 253 tyrosine hydroxylase ( = TH), 12, 71, 183,

201

uperolein, 85 ureter (see Urinogenital Organs Ch. 9), 311 ff. urinary bladder, 33, 42, 46, 212, 316-320 urodele amphibians (see amphibia) urogastrone, 179 urotensin, 83 uterus (see Urinogenital Organs Ch. 9), 311 ff. uvea, 16 vagal sensory ganglion complex, 38 vagally mediated vasoconstriction, 214 vagus nerve

371

anatomical arrangement, 4, 7, 9-10, 13, 18-24, 26-27, 29, 33, 37, 40, 41-43, 46-48, 50 innervation o f the vasculature, 214, 217 innervation o f glands, 173 innervation o f the heart, 196-200, 204-205, 207 innervation o f the lung, 263-265, 269-284, 288 vagosympathetic nerve trunk, 46, 199-200, 204, 212, 292 vas deferens, 324 vasculature (see The Circulatory System Ch. 7), 193 ff. vasoactive intestinal polypeptide ( = VIP), 87 anatomical considerations on distribution, 4-5, 16, 18, 23, 25, 27 in glands, 170, 172, 174, 176, 178-179, 184-185, 187 in the iris, 00 in the gut, 124, 151, 158 in the heart, 199-200, 206-207 in the lung, 264-265, 273, 276, 282-283 in the spleen, 251 in the swimbladder, 296-297 in the urino-genital system, 318, 323 in the vasculature, 210, 212-214, 217, 219-221, 223 vasoconstriction, 209, 213, 216, 222 vasodilatation, 12-13, 16-18, 24-25, 32, 46, 209, 213, 216, 221, 223 vasomotor neurons, 6, 46 vasopressin, 319 vasotocin, 314 Vidian nerve (nerve o f the pterygoid canal), 14, 16, 24, 28 VIP (see vasoactive intestinal polypeptide) vis a tergo, 195 vis a fronte, 195 visceral ramus o f the vagus, 10, 30, 46 xanthophore, 332 Zuckerkandl, organ of, 109

Biosystematìc Index Acipenser sp., 8, 19, 42, 43 Acrochordus granulatus, 281-282, 284 Agama agama, 123 Aldrichetta fors teri, 126 Aldrichetta sp., 9 Alligator sp., 23, 49 Alligator mississippiensis, 128, 144, 146,

Bufo calamita, 128, 129, 134, 135, 141, 142, 146, 153, 154

Bufo marinus (and Bufo sp.), 4, 8, 21, 25,

27, 44-46, 53-54, 83, 84, 123, 126, 141, 144-146, 148, 150, 152, 200, 202-203, 206, 216, 228, 249, 272-274, 278-279, 286-287, 314-315, 318, 324 Bufo viridis, 273 Bufo vulgaris, 274

153, 358

Alytes obstetricans, 152 Ambystoma sp., 9, 21, 43, 54, 278, 281,

Caiman crocodylus crocodylus, 23, 27, 49,

285, 287

Ambystoma mexicanum, 86, 141, 142,

124, 128, 130, 131, 140-142, 146, 148, 150, 153, 282 Calamoichthys sp., 257-258, 288 Carassius sp., 9, 314 Carassius auratus, 79, 86, 126, 138, 139, 141, 146, 152 cat (= Fe/is artws), 11-12, 15-18, 31-32, 208 Centrina sp., 35 Centrolabrus exoletus, 139, 141, 152 Chamaleo sp., 47-48

146, 153

Ambystoma tigrinum, 128 Ameiurus sp., 341, 342

ctf/vtf, 54, 129, 136, 152, 258

Ammotretis rostrata, 123 Ammotretis sp., 42 Amphibolurus barbatus, 282 Amphioxus (see Branchiostoma) Amphiuma sp., 9, 23 Anarhichas sp. 40, 42 platyrhynchos, 269, 270

Channa argus, 42 Chelodina sp., 282, 284, 285 Chelone sp., 48 Chelydro sp., 23-24, 48, 54 chicken, see Gallus domesticus Chimaera monstrosa, 126, 128, 135, 152,

, 50

Anguilla anguilla, 141, 146, 152 Anguilla australis, 126 Anguilla sp., 9, 142, 290-291, 293, 295 Anolis sp., 23 Anser anser, 269, 270 ^4/Jser

5 /7 . ,

172

Chimaera sp., 11, 19-20, 35 Ciona intestinalis, 11 Ciliata mustela, 128, 135, 138, 146, 152 Columba livia, 11, 50, 205, 271 Conger conger, 40-41 Corydoras schultzei, 141, 152 Cottus scorpius, see Myoxocephalus scorpius Coturnix coturnix sp., 146, 153

50

Atelops oxyrhyncus, 152 Barbus conchonius, 139, 141, 144, 146, 152

Bombina bombina, 146, 152, 273 Bombina variegata, 75, 273 Brachydanio rerio, 152 Branchiostoma lanceolatum (= Amphioxus), 9, 179 Bufo bufo, 146, 152

crocodiles, 23, 25, 28, 49-50 Crocodylus niloticus, 128, 144, 146, 153 373

374

COMPARATIVE PHYSIOLOGY AND EVOLUTION

C rocodylus p o ro su s, 124, 130, 148, 150,

185

C rocodylus jo h n sto n i , 282 C ryptobranchus s p . , 280 Ctenolabrus rupestris , 128, 139, 141, 146,

152, 292-295

C tenophorus s p ., 23, 47 C ygnopis cygn o id , 270 Cygnus olor, 324 C yn ops hongkongensis , 153 C ynops pyrrh o g a ster , 358 C yprinus carpio , 86, 128, 135, 138, 142,

143, 146, 148, 152, 225

D erm ophis s p ., 23, 47 D esm ognathus s p . , 9 dog ( = Canis fam iliaris), 12, 18, 32, 208 E laphe o b so leta , 11, 23, 27, 49, 204, 217,

219, 281-282, 284-285 F/wys 5/7., 48 Fmys blandingi , 358 F/wys orbicularis E ndoshenus ja p o n ic u s , 196 E ptatretus sp ., 196 E retm ochelys sp ., 23, 48 Fso* lucius, 294 Fe/is

5/7.

(see cat)

Fundulus parvipen n is, 334 Fundulus heteroclitu s, 136, 314 Fundulus s p ., 341, 342

G*w/ m5 m orhua, 19-20, 38, 40, 42, 73, 76, 77, 79, 87, 122-124, 126, 128, 130-132, 134-136, 138, 141, 143, 145, 146, 148, 153-156, 171, 172, 173, 199, 213, 226, 227, 248, 291-295, 317, 322, 357 Galeus s p ., 35 G allus dom esticu s (also chicken), 50-54, 126, 139, 140, 141, 146, 148, 152-156, 316 G em m atophora s p ., 47 G eoclem ys s p ., 8 G eotrypes s p ., 23, 47 G illichtys m irabilis, 139, 141, 146, 152 G obius s p ., 293 G ongylus s p ., 48 guinea-pig ( = Cavia cavia ), 5, 8, 11-12, 16-18, 31-32, 52, 206 G yrinochelius aym onieri, 152 H aplochrom is s p ., 128, 139, 152 H atteria s p ., 47

H eloderm a horridum , 87 H eloderm a su spectu m , 87 H elo sto m a tem m inicki, 139, 141, 152 H em igram m us ocellifer, 152 H em itripteru s am ericanus, 227 H erpele s p ., 23, 47 H etero d o n tu s p o rtu sja ck so n i, 35, 38,

198 horse ( = Equus caballus ), 17 human ( = H o m o sapiens ), 11-12, 16-18, 31-32, 52 i / w 5 0 huso, 199 //W 5 0 5 /7 ., 42-43 H ydrolagu s s p ., 11, 19-20, 35 H y d ro m a te s italicus, 128 H y la crepitans, 153 H y la arborea, 153, 273, 278 H yp o g eo p h is s p ., 47 Ich th yoph is s p ., 23 Ictalurus m etas, 155, 156 Ictalurus s p ., 20, 83 K assina senegalensis, 86 K ryto p te ru s bicirrhi, 344, 346 L a b ra x sp. ( = A narhichas sp .), 38 L abru s berggylta, 139, 141, 146, 152 L abru s m ixtus, 139, 141, 152 L abru s ossifagus, 345 L acerta viridis, 153 L acerta viviparus, 137 L acerta s p ., 23, 47-48, 54, 281, 324 L am pan yctu s sp. 11, 19-20, 38, 40 L a m p etra flu via tilis, 124, 126 L a m p etra ja p ó n ica , 128, 129 L a m p etra plan eri, 126 L a m p etra sp ., 9, 19, 34-35, 196, 197, 209,

312

L eb istes sp. 343, 345, 346 L epidosiren p a ra d o x a , 19-20, 126, 129,

135, 139, 141, 142, 144, 147, 148, 152, 199, 200, 288-289 L episosteu s platyrh incus, 128, 139, 152, 153, 199, 212 L episosteu s s p ., 43, 288-289 L ep o m is m acrochirus, 136, 152 Leuciscus idus, 128, 135, 139, 146, 152 L im an da yo k o h a m a e, 316, 322 L im n odyn astes s p ., 44 L o p h iu s s p ., 83 L o p h iu s am ericanus, 79, 127, 183 L o p h iu s piscatoriu s, 318, 322, 355

BIOS YSTEMATIC INDEX M eleagris g a llo p a v o , 316 M e n id ia s p ., 12, 19-20, 38-39 M erlangius m erlangus , 86 M icropterus s p . , 20 M in iopterus schreibersii , 207 mouse ( = M u s m usculus ), 17, 32 mudpuppy (see N ecturus maculosus) M ustelus s p ., 9, 19, 35, 198, 355 M ustelus laevis , 355 M yliobates s p ., 35 M yoxocephalus s p . , 8 M yoxocephalus scorplu s (C o ttu s scorpiu s),

126, 128-131, 135, 137-139, 141, 146, 152 M yripristis occidentalis , 345 M yxine glu tin osa , 35, 124, 126, 151 Myxi/ie 5/7., 120, 196, 197, 209

N atrix natrix , 324 N ecturus m aculosus , 8, 43, 54, 128-131,

133-136, 138-146, 148, 150, 153-156, 200-203, 228, 280, 319 N ecturus sp ., 176, 178 N eoceratodus fo r s te r i , 200, 258, 288

O don tobu tis obscura, 344 O ncorhynchus m ykiss ( = Salm o gairdnen),

101, 128, 130-132, 136, 138, 139, 140, 142, 143, 145, 146, 148, 152, 155, 172, 199, 213, 313-314 O ncorhynchus kisutch, 78, 248 O phiocephalus s p ., 8 O psanus s p ., 54, 292 O psanus tau, 355 O reochrom is m ossam bicus, 126, 139, 141, 152 O reochrom us niloticus, 322 O ryzias sp ., 321, 323 P antodon bu ch h olzi , 152 P aralichthys s p ., 332 Parasilurus s p ., 8, 345, 346 Parasitus s p ., 42 P aroph rys vetulus, 314 Paras cristatus, 75, 141-143, 146, 153 P elm atoch rom is pu lch er, 141 P elom edusa s p ., 287 Perca flu via tilis, 20, 128, 138, 139, 141,

142, 146, 152, 292-293

P etrom yzon s p ., 19, 35 P hoxinus phoxin u s, 336, 338, 340-342,

345, 347

P h yllodactylu s sp ., 23, 47 P hyllom edu sa sp ., 86

375

pigeon (see C olum ba livid) P latich th ys flesu s, 126, 128, 138, 139, 141, 146, 152 P latich th ys s p . , 338 P latycephalu s s p ., 8-9, 38, 40-42 P latypoecilu s variatus, 139 platypus ( = Ornithorhyncus anatinus), 1 P leuronectes pla tessa , 126, 146, 152 P leuronectes sp ., 8, 20, 338, 346 P o d a rcis hispanica, 146, 153 P oecilia reticulata, 128, 138, 139, 141, 146 P og o n a sp ., 23, 47 P o g o n a barbatus, 128, 139, 146, 153 P o g o n a viticeps, 219 P ollachius pollachius, 128, 135, 138, 146, 152 Pollachius s p ., 291, 295 P o ly o d o n spath ula, 9, 19 P o lyp teru s bichir, 257-258, 288 P o lyp teru s senegalensis, 26, 128, 141, 142, 146- 148, 152 P rio n o tu s s p ., 293 P ro to p teru s aethiopicus, 110 P ro to p te ru s annectens, 128, 129, 134, 139, 141, 144, 146 P ro to p te ru s s p ., 20, 43, 199, 200, 288-289 P seu dem ys scripta elegans, 120 P seu dem ys s p ., 8, 282, 284, 286-287 P yth o n sp ., 23 rainbow trout (see O ncorhynchus m yk iss = Salm o gairdneri) R a ja batis, 86 R a ja clavata, 87, 128, 132, 152 R a ja erinacea, 19, 128, 129, 132 R a ja m icrooscellata, 128, 138 R a ja m ontagui, 128, 138 R a ja naevus, 128, 138 R a ja radiata, 87, 138, 148, 152, 185 R a ja rhina, 210 R a j a s p ., 20, 35, 120, 127, 136, 137, 147148, 151, 182, 197, 210 R ana catesbeiana, 44, 46·, 79, 102, 126, 174, 178, 179, 216, 314-315, 319 R an a esculenta, 44-46, 53, 77, 146, 153, 203 R ana lessonae, 273 R an a pip ien s, 44, 46, 77 R ana ridibunda, 79, 102, 144, 146, 153 R an a sp ., 8-9, 11, 21, 44-45, 54, 274, 278-279, 287, 234 R ana tem poraria, 44, 46, 79, 137, 141, 142, 146, 153, 315, 318 R an iceps raninus, 128, 135, 138, 146, 152

376

COMPARATIVE PHYSIOLOGY AND EVOLUTION

rat ( = R attu s sp .), 11-12, 16-18, 32-33, 52, 105 R h om bosolea s p ., 9, 42 R h om bosolea tapirina , 123 Saccobranchus s p . , 8 Salam andra salam andra , 8, 43, 145, 146,

153, 324

Salm o sp. (see also O ncorhynchus m ykiss = Salm o gairdnen ), 8-9, 20, 42,

180, 182, 293, 313

Salm o tru tta , 126, 213 Sander s p . , 20 Scaphyrhynchus , 19 Scophthalm us sp ., 333, 338 Scorpaeichthys m arm oratu s , 130 Scyliorhinus caniculus/canicula, 85, 87,

128, 129, 185, 197, 210, 226, 248, 355

Scyliorhinus stellaris, 126, 128, 135,

144-146, 151, 152

Scyliorhinus s p ., 9, 36-37, 86, 120, 312,

355

ScyIlium catulus, 355 Scyllinum s p ., 9, 35, 198, 321, 355 Seriola quinqueradiata, 248 Serranus s p ., 40 Silurus s p ., 8 Siredon s p ., 43 Siren lacertina, 102 Solea sp ,, 338 Sphenodon pu n cta tu s, 271 Squalus sp ., 182 Squalus acanthias, 9-10, 19, 35, 54, 73,

77, 78, 110, 120, 124, 126, 128-130, 134-137, 144-148, 150-156, 185, 210, 248 Squatina aculeata, 128, 129 Squatina s p ., 35 Sym branchus m arm oratu s, 344

S yn oodn tu s s p ., 346 Tejus s p ., 47 T estudi kleim anni, 86 T estudo s p ., 48, 282, 284-285, 287, 324 T etractenos glaber, 126 Tetractenos s p ., 292-295 Thalassochelys s p ., 48, 54 T ham nophis sp ., 23, 49, 282, 284 Tiliqua rugosa (see also Trachydosaurus rugosus ), 120 Tiliqua s p ., 8, 23, 25, 42, 47 Tinca tinca, 293 T orpedo m atm orata, 79, 83, 84, 86 T orpedo s p ., 35, 74, 80, 86, 321 Torquigener s p ., 9 Trachydosaurus rugosus, 219, 250,

281-284, 319

Trichiurus sp ., 19-20 Trionyx s p ., 23, 43-44, 48 Triturus alpestris, 280 T ropidon otu s s p ., 23 Trygon s p ., 35 Trygon violaceus, 355 T ym panocryptu s s p ., 47

Turkey, 153

U ranoscopus scaber, 19-20, 38, 40, 42,

322, 355

Varanus s p ., 22-23, 47 Varanus gou ldii, 128, 134, 135, 137-139,

144, 146, 153, 219

Vipera aspis, 153 Vipera berus, 128, 137-140, 146, 148, 150,

153, 254

Xenopus laevis, 9, 77, 78, 82, 153, 202, 285, 287, 315, 324