The Enteric Nervous System
John Barton Furness PhD, FAA Department of Anatomy and Cell Biology, University of Melbourne...
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The Enteric Nervous System
John Barton Furness PhD, FAA Department of Anatomy and Cell Biology, University of Melbourne, Victoria, Australia
The Enteric Nervous System
The Enteric Nervous System
John Barton Furness PhD, FAA Department of Anatomy and Cell Biology, University of Melbourne, Victoria, Australia
© 2006 John B. Furness Blackwell Publishing, Inc., 350 Main Street, Malden, Massachusetts 02148-5020, USA Blackwell Publishing Ltd, 9600 Garsington Road, Oxford OX4 2DQ, UK Blackwell Publishing Asia Pty Ltd, 550 Swanston Street, Carlton, Victoria 3053, Australia The right of the Author to be identified as the Author of this Work has been asserted in accordance with the Copyright, Designs and Patents Act 1988. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by the UK Copyright, Designs and Patents Act 1988, without the prior permission of the publisher. First published 2006 Library of Congress Cataloging-in-Publication Data Furness, John Barton. The enteric nervous system / John B. Furness. p. ; cm. Includes bibliographical references and index. ISBN-13: 978-1-4051-3376-0 ISBN-10: 1-4051-3376-7 1. Gastrointestinal system--Diseases. I. Title. [DNLM: 1. Enteric Nervous System--physiology. 2. Neurons --physiology. WL 600 F988e 2006] RC817.F87 2006 616.3--dc22 2005024527 ISBN-13: 978-1-4051-3376-0 ISBN-10: 1-4051-3376-7 A catalogue record for this title is available from the British Library Set in 10/13½ Sabon by Sparks, Oxford – www.sparks.co.uk Printed and bound by Narayana Press, Odder, Denmark Commissioning Editor: Alison Brown Editorial Assistant: Saskia Van der Linden Development Editor: Rob Blundell Production Controller: Kate Charman For further information on Blackwell Publishing, visit our website: http://www.blackwellpublishing.com The publisher’s policy is to use permanent paper from mills that operate a sustainable forestry policy, and which has been manufactured from pulp processed using acid-free and elementary chlorine-free practices. Furthermore, the publisher ensures that the text paper and cover board used have met acceptable environmental accreditation standards.
Contents
Preface, ix Abbreviations, xi
1: Structure of the enteric nervous system, 1 The enteric plexuses, 3 Interconnections between the plexuses, 14 Extent of the ganglionated plexuses, 15 Intramural extensions of extrinsic nerves, 17 Electron microscope studies, 17 Enteric glia, 20 The structural similarities and functional differences between regions may have an evolutionary basis, 21 Development of the enteric nervous system, 23 Maturation of enteric neurons and development of function, 26 Changes in enteric neurons with aging, 27 Summary and conclusions, 28
2: Constituent neurons of the enteric nervous system, 29 Shapes of enteric neurons, 31 Cell physiological classifications of enteric neurons, 43 Functionally defined enteric neurons, 53 Neurons in human intestine with equivalence to those investigated in laboratory animals, 76 Summary and conclusions, 78
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3: Reflex circuitry of the enteric nervous system, 80 Evolution of ideas about enteric circuitry, 80 Motility controlling circuits of the small and large intestine, 81 Intrinsic secretomotor and vasomotor circuits, 88 Assemblies of neurons, 93 Circuits in the esophagus and stomach, 96 Co-ordination of motility, secretomotor, and vasomotor reflexes, 98 Circuits connecting the intestine, biliary system, and pancreas, 98 Sympathetic innervation of the gastrointestinal tract, 99 Summary and conclusions, 101
4: Pharmacology of transmission and sites of drug action in the enteric nervous system, 103 Chemical coding and multiple transmitters, 103 Transmitters of motor neurons that innervate the smooth muscle of the gut, 104 Transmitters at neuro-neuronal synapses, 111 Sites within the reflex circuitry where specific pharmacologies of transmission can be deduced to occur, 120 Transmission from entero-endocrine cells to IPANs, 126 Roles of interstitial cells of Cajal in neuromuscular transmission, 127 Transmitters of secretomotor and vasodilator neurons, 128 Synapses in secretomotor and vasodilator pathways, 130 Transmitters of motor neurons innervating gastrin cells, 130 Summary and conclusions, 130
5: Neural control of motility, 132 Rhythmic activity of gastrointestinal muscle, 132 Structure and properties of interstitial cells of Cajal, 134 Relationship between slow wave activity and neural control, 138 Gastric motility, 140 Patterns of small intestine motility and their intrinsic neural control, 147 Motility of the colon, 157 Neural control of the esophagus, 159 Gall bladder motility, 160 Sphincters, 161 Muscle of the mucosa, 165
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Mechanism of sympathetic inhibition of motility in non-sphincter regions, 166 Sympathetic innervation of the sphincters, 169 Physiological effects of noradrenergic neurons on motility in undisturbed animals, 170 Reflex activities of sympathetic neurons that affect motility, 171 Summary and conclusions, 178
6: Enteric neurons and the physiological control of fluid secretion and vasodilation, 180 Water and electrolyte secretion in the small and large intestines, 180 Reflex control of water and electrolyte secretion, 182 Secretion of gastric acid, 189 Pepsinogen secretion, 194 Gastric secretion of bicarbonate, 195 Secretion into the gall bladder, 195 Pancreatic exocrine secretion, 196 Summary and conclusions, 198
7: Disorders of motility and secretion and therapeutic targets in the enteric nervous system, 200 Therapeutic endpoints for motility disorders, 201 Therapies for secretory diarrheas, 205 Enteric neuropathies involving neuronal loss or phenotypic changes, 206 Mitochondriopathies with intestinal manifestations, 207 Irritable bowel syndrome and plasticity of enteric neurons, 208 Summary and conclusions, 210 Epilogue: the future of enteric neurobiology, 211 References, 214 Index, 267
Preface
The enteric nervous system is of special interest because it is the only substantial grouping of neurons outside the central nervous system that form circuits capable of autonomous reflex activity. In humans it contains around 500 million neurons that fall into about 20 functional classes. Because of its size, complexity, and certain structural similarities, it has been likened to a second brain. Although the enteric nervous system was discovered almost 150 years ago, and several remarkably insightful hypotheses about its functions were made in the 19th century, a long period ensued in which progress was meagre in comparison to the effort made, because methods available were not adequate to determine the intrinsic circuitry of the enteric nervous system and the properties of its constituent neurons. In the last 20–30 years, new techniques, and excellent application of such techniques, have provided a wealth of information on the structural complexity, neuron types, and connectivity of the enteric nervous system and on the transmitters and cell physiology of enteric neurons. Beginning at an earlier time, and proceeding in parallel, have been investigations of the patterns of movement and secretory functions of the digestive tract, and their control. This book aims to integrate the detailed cellular knowledge of the enteric nervous system with the more macroscopic information that is provided by physiological studies of organs, especially in the living animal or human. In doing so, I have tried to deal with the emergence of knowledge in historical perspective, where possible by drawing on early information to acknowledge the contributions made by pioneers of enteric neurobiology, and in places to reproduce original illustrations from early publications. I hope that the reader will enjoy this approach. I have also created many new illustrations, especially of the organization of enteric nerve circuits, which I hope will provide an understanding of the enteric nervous system that the written word cannot easily convey. The first four chapters lay the groundwork, by dealing with the structure of the enteric nervous system, the defining cell physiological, morphological, ix
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and neurochemical properties that allow its neurons to be functionally classified, the enteric neurotransmitters and the intrinsic nerve circuits within the alimentary tract. This is followed by two chapters on gastrointestinal physiology, first on the contractile activity of the muscular walls of the digestive tract and the second on secretory function. In these two chapters I try to develop an understanding of the roles of enteric neurons and how they perform these roles. I have also sought to relate control through enteric circuits to control exerted by the vagus and the sympathetic innervation of the digestive organs, and to a lesser extent through the pelvic nerves. The involvement of altered structure and function of the enteric nervous system in some disease states is well recognized. Nevertheless, how to use the new-found knowledge of the enteric nervous system to understand the relations between changes in the neurons and clinical manifestations of disease is a challenge. Moreover, how the neurons might be manipulated by therapeutic compounds to ameliorate disorders of the digestive system is elusive, in many cases. The problems of understanding and treating digestive diseases that involve the enteric nervous system, or functions controlled by the enteric nervous system, are touched on throughout the book, and are specifically discussed in Chapter 7. In writing this book I have relied on the assistance and advice of many colleagues who have generously read and commented on parts of book, in some cases through several drafts. My special thanks go to Dr Paul Andrews, Dr Joel Bornstein, Dr Axel Brehmer, who also helped me with the interpretation of some of the older literature published in German, Dr Nadine Clerc, Dr Helen Cox, Dr Roberto de Giorgio, Dr Giorgio Gabella, Dr Peter Holzer, Dr Terumasa Komuro, Dr Alan Lomax, Dr Kulmira Nurgali, Dr Michael Schemann, Dr Keith Sharkey, Dr Henrik Sjövall, Dr Werner Stach, who provided previously unpublished micrographs, Dr Jean-Pierre Timmermans, Dr Marcello Tonini and Dr Heather Young. For assistance in the preparation of the illustrations I am very grateful to Melanie Clarke, Anderson Hind, and Trung Nguyen, and for editorial help and assistance with the references, to Emma James. I would also like to thank the many colleagues who gave permission for illustrations to be included in the book. I hope that this book succeeds in linking the extensive knowledge of the structure and cell physiology of the enteric nervous system to an understanding of digestive physiology, and that in so doing it helps provide a rational basis for therapeutic intervention, and even reasons why some interventions may fail. I enjoyed writing the book, although at times it was a hard task. I hope that in reading the book you encounter only the enjoyment. John B Furness Melbourne, May 2005
Abbreviations
AC, adenylyl cyclase ACh, acetylcholine AChE, acetylcholine esterase ADP, after-depolarizing potential AH, designation of neurons having slow after-hyperpolarizing potentials AHP, after-hyperpolarizing potential AMP, adenosine monophosphate AMPA, alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid AP, action potential ATP, adenosine triphosphate BK, large-conductance potassium channel BMP, bone morphogenic protein BN, bombesin (the mammalian form also referred to as GRP, below) cAMP, cyclic adenosine monophosphate CCK, cholecystokinin CFTR, cystic fibrosis transmembrane conductance regulator CGRP, calcitonin gene-related peptide ChAT, choline acetyltransferase CM, circular muscle CNS, central nervous system DAG, diacyl glycerol DMP, deep muscular plexus DMPP, dimethyl phenyl piperazinium DYN, dynorphin ECL, enterochromaffin-like (cell) EJP, excitatory junction potential ENK, enkephalin EPSP, excitatory post-synaptic potential GABA, γ-aminobutyric acid GAL, galanin gCav, voltage-sensitive calcium conductance gK , Ca2+-dependent K+ conductance Ca gNav, voltage-dependent Na+ conductance GRP, gastrin-releasing peptide (also known as mammalian bombesin) xi
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Gs, stimulating G-protein 5-HT, 5-hydroxytryptamine (serotonin) HCN, hyperpolarization activated non-specific cation conductance HVA, high-voltage activated calcium current IAHP , AHP current IBS, irritable bowel syndrome ICav, voltage-sensitive calcium current ICC, interstitial cell(s) of Cajal Ih, hyperpolarization-activated cation current IK, intermediate-conductance potassium channel IKATP, ATP-dependent potassium current IPAN, intrinsic primary afferent neuron IPSP, inhibitory post-synaptic potential LM, longitudinal muscle MAP2, microtubule associated protein 2 MELAS, multisystem mitochondriopathy MMC, migrating myoelectric complex MNGIE, mitochondrial neurogastrointestinal encephalomyopathy MP, membrane potential Muc, mucosa L-NAME, L-nitro-arginine methyl ester nAChRs, nicotinic acetylcholine receptors NANC, non-adrenergic, non-cholinergic NFP, neurofilament protein Nic, nicotinic NK, neurokinin NO, nitric oxide NOS, nitric oxide synthase NPY, neuropeptide tyrosine, usually known as neuropeptide Y P2X, purine receptor 2X P2Y, purine receptor 2Y PACAP, pituitary adenylyl cyclase activating peptide PCR, polymerase chain reaction PDBu, phorbol dibutyrate PHI, peptide histidine isoleucine PHM, peptide histidine methionine PKA, protein kinase A PKC, protein kinase C PLC, phospholipase C PPADS, pyridoxal-phosphate-6-azophenyl-2΄,4΄-disulfonic acid PVG, prevertebral ganglion PYY, peptide tyrosine tyrosine Rin, input resistance RT, room temperature SAC, stretch activated channel SGLT1, Na+/glucose co-transporter 1 SK, small-conductance potassium channel
A BBR EV IAT IONS
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SM, submucosa SOM, somatostatin SSPE, sustained slow post-synaptic potential STC, slow-transit constipation TEA, tetraethylammonium TK, tachykinin TRH, thyrotropin-releasing hormone TTX, tetrodotoxin TTX-R INaV, TTX-resistant sodium current VIP, vasoactive intestinal peptide VPAC, vasoactive intestinal peptide; pituitary adenylyl cyclase activating peptide
1: Structure of the enteric nervous system
A vast amount of neural tissue, which constitutes the enteric nervous system, is embedded in the wall of the gastrointestinal tract. Within the enteric nervous system, nerve cells and supporting (glial) cells are grouped in small clusters, the enteric ganglia, which are interconnected by nerve fiber bundles (Fig. 1.1). The individual ganglia are small, but are so numerous that the system as a whole contains millions of nerve cells. The processes of these nerve cells connect with other neurons and innervate the muscle, secretory epithelium, and blood vessels of the digestive tract, biliary system, and pancreas. Processes of nerve cells from outside the digestive tract also connect with enteric neurons, and intermingle with processes of enteric neurons. A remarkable aspect of the enteric nervous system is that its reflex circuits are capable of directing the functions of the digestive system without relying on commands from the brain or spinal cord. This independence is modulated by the rich interchange of signals between the enteric and central nervous systems. The first clear descriptions of ganglionated plexuses within the wall of the digestive tract were those of Meissner (1857), Billroth (1858), and Auerbach (1862a,b, 1864). Remak (1840, 1852) had earlier noted the presence of microscopic ganglia in the walls of the pharynx and stomach, but his descriptions do not suggest that he recognized a ganglionated plexus. Following their discovery, the enteric ganglia and plexuses attracted considerable attention and numerous descriptions of their organization were published, including those of Henle (1871), Drasch (1881), Dogiel (1895b, 1899), Cajal (1911), Kuntz (1913, 1922), Hill (1927), Schabadasch (1930a,b), Stöhr (1930), and Irwin (1931). These studies, and the contemporary literature they cite, provide detailed information on the sizes, arrangements and interconnections of the ganglia. The descriptions that Meissner, Billroth, and Auerbach provided of the general organization of the ganglionated plexuses, based on quite primitive techniques to reveal the nerve tissue, were not superseded by work in the subsequent 100 years and the descriptions of the arrangements of the enteric plexuses that 1
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are set out in the following pages were essentially established by the time of the reviews of Schabadasch (1930a,b) and Stöhr (1930). An English translation of Auerbach’s 1864 description has been published (Furness & Costa 1987). The enteric nervous system of the tubular digestive tract (the esophagus, stomach, and intestines) is formed of a number of interconnected networks, or plexuses, of neurons, their axons, and enteric glial cells (Fig. 1.1). In the
Fig. 1.1 The enteric plexuses as they are seen (A) in wholemounts and (B) in transverse section.
The drawings depict the small intestine. There are two ganglionated plexuses, the myenteric and the submucosal plexuses, in addition to nerve fibers that innervate the muscle layers, the mucosa and intramural arterioles. Nerve fibers enter the intestine with mesenteric blood vessels in paravascular nerves (B). Adapted from Furness and Costa (1980).
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small intestine and colon, most nerve cells are found in two sets of ganglia, the ganglia of the myenteric (Auerbach’s) plexus and of the submucosal plexus (often referred to as Meissner’s plexus, but see below). The axons of these nerve cells innervate other ganglia and the tissues of the digestive organs, such as the muscle layers and the mucosa. The enteric plexuses Myenteric plexus The myenteric plexus is a network of nerve strands and small ganglia that lie between the outer longitudinal and inner circular muscle layers of the external muscle coat of the intestine (Fig. 1.2). The network is continuous around the circumference and along the gastrointestinal tract (Fig. 1.3). The myenteric ganglia vary in size, shape, and orientation between animal species and from
Fig. 1.2 Drawing of a whole-
mount of the myenteric plexus of the human small intestine, prepared by Auerbach and published in Henle’s Textbook of Histology in 1871. Myenteric ganglia, internodal strands, and small nerve trunks of the secondary component of the myenteric plexus (arrows) can be seen. The secondary nerve strands supply fibers to the circular muscle and deeper layers. Calibration: 1 mm.
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Fig. 1.3 Schabadash’s (1930a) depiction of the myenteric plexus of the pyloric canal in the cat. The large dark areas are the ganglia, which are connected by nerve fiber bundles of various calibers. The continuity of the plexus around and along the gut wall can be clearly seen. Calibration: 2 mm.
one part of the intestine to another (Fig. 1.4), but the shape of the meshwork is usually characteristic and readily identified in any major region from a particular species (Irwin 1931, Gabella 1981a). Although the pattern is easily recognized, considerable variation in the size of ganglia is encountered. In the ileum of the guinea-pig, ganglia range in size from 5 to over 200 nerve cell bodies. Single nerve cell bodies are occasionally encountered outside the main meshwork of the plexus, usually adjacent to a nerve strand. The ganglia are sometimes referred to as the nodes of the plexus because they lie at the junctions of nerve strands, which in turn are called internodal or interganglionic strands, or sometimes interganglionic connectives. The ganglia are deformable and are distorted by the movements of the muscle. Thus measurement of such features as their shape and spatial density must take into account the state of contraction of the gut wall (Gabella & Trigg 1984). Three components of the myenteric plexus are described (Fig. 1.5): a primary plexus, a secondary plexus, and a tertiary plexus (Auerbach 1864, Schabadasch 1930a,b, Stöhr 1930, Li 1940, 1952). Together, the ganglia and internodal strands make up the primary meshwork of the myenteric plexus. Many of the nerve fibers in an internodal strand do not enter the ganglion with which a strand connects, but pass over the ganglion, usually between the ganglion and the longitudinal muscle, and continue in another internodal strand. Finer nerve fiber bundles, constituting the secondary component of the plexus, branch from the primary internodal strands or arise from ganglia, but do not usually link adjacent ganglia. The secondary strands run parallel to the circular muscle bundles and often cross internodal strands. They run on the inner aspect of the primary plexus and ganglia, between the primary plexus and the circular muscle (Schabadasch 1930b, Stöhr 1952). Auerbach (1864) traced nerve processes from the secondary strands to the circular muscle, a connection that has been confirmed (Wilson et al. 1987). The secondary
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Fig. 1.4 Drawings of the myenteric plexus in different regions of the gastrointestinal tract of the guinea-pig: A, esophagus; B, pylorus; C, duodenum; D, ileum; E, colon; F, rectum. Note that patterns of the ganglia differ between regions. They also differ between species. All at the same magnification, except B. The calibration lines are 1 mm apart. Reproduced from Irwin (1931).
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1 1
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Fig. 1.5 The three components of the myenteric plexus found in small animals are shown in a drawing of a wholemount from the guinea-pig small intestine. Common to all species is the primary component of the plexus (1), consisting of the ganglia and internodal strands (interganglionic connectives), and the secondary component (2), consisting of nerve strands lying parallel to the circular muscle (across the page). The tertiary plexus (3) is found only where the longitudinal muscle is thin; in such regions, few nerve fibers are found within the longitudinal layer. Neuron cell bodies are depicted as white ovals in the ganglia. Redrawn from Furness and Costa (1987). Calibration: 100 μm.
strands can be seen in Auerbach’s drawing (Fig. 1.2). The tertiary meshwork (tertiary plexus) is made up of fine nerve bundles that meander in the spaces between the meshwork formed by the primary plexus (Richardson 1958, Llewellyn Smith et al. 1993, Furness et al. 2000) (Fig. 1.5). Nerve bundles of the tertiary plexus can be traced from primary internodal strands, ganglia and secondary strands. The definition of the tertiary plexus given here accords with that of Stöhr (1930), which is different from that given by Schabadasch and Li. The definitions of the latter authors combine the secondary and tertiary plexuses under the name secondary plexus and they call the tertiary plexus those fine fibers that run parallel to and extend into the circular muscle and which join the deep muscular plexus. I refer to these fibers as the circular muscle plexus, or simply as the circular muscle innervation. Submucosal plexus A submucosal ganglionated plexus is found in the small and large intestine (Figs 1.6, 1.7), and was first described in the mid-19th century by Meissner (1857) and Billroth (1858). Although scattered ganglia are found in the submucosal layer in the esophagus and stomach, these do not form a ganglionated plexus comparable to that of the intestines. In general, the interconnecting strands of the submucosal plexus are finer and the ganglia are smaller than those of the myenteric plexus (Henle 1871,
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Fig. 1.6 Distribution of enteric ganglia in the tubular digestive tract. The gastrointestinal tract is represented schematically in longitudinal section to reveal the myenteric ganglia, which form a continuous plexus from the upper esophagus to the internal anal sphincter, and the submucosal plexus, which is prominent in the small and large intestines. Isolated ganglia occur in the gastric and esophageal submucosa and in the mucosa throughout the digestive tract. From Furness et al. (1991).
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Fig. 1.7 Drawing of the submucosal plexus of the small intestine of a 6-day-old child, published by Billroth (1858). It accurately depicts the ganglia, with nerve cells drawn as small circles, and the connecting strands. Note that Billroth depicts ganglia and nerve strands at two levels. Calibration (approx): 250 μm.
Goniaew 1875, Timmermans et al. 2001). The plexus is continuous around the circumference and along the length of the small and large intestines. The arrangements of ganglia in the submucosal plexus, and the functional types of neurons in these ganglia, differ between species (Scheuermann et al. 1987b,c, Hoyle & Burnstock 1989, Timmermans et al. 1990). In large animals, good examples being the pig and human, submucosal ganglia form distinct, but interconnected, plexuses that lie at different levels, as first clearly described by Schabadasch (1930b). Two or sometimes three layers of ganglia have been distinguished (Schabadasch 1930b, Gunn 1968, Hoyle & Burnstock 1989, Timmermans et al. 2001). Ganglia at different depths contain different populations of neurons, these variations being apparent in the shapes and chemical natures of the constituent nerve cells. The inner ganglionated plexus (closer to the gut lumen) has been likened to the plexus described by Meissner (1857) and the outer has been identified with that described by Henle (1871) and Schabadasch (1930b). Because it is not completely clear who should be credited with the discovery of individual components of the submucosal plexus, it seems sensible to refer to the most obvious groupings as the inner and outer submucosal plexuses (Timmermans et al. 2001), the inner being that closer to the intestinal lumen, and the outer that closest to the circular muscle layer. Among the neurons of the outer plexus are some that supply innervation to the circular and even to the longitudinal muscle (Sanders & Smith 1986, Furness et al. 1990a, Timmermans et al. 1994, 1997,
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Porter et al. 1999). The outer submucosal plexus also supplies innervation to the mucosa. The inner submucosal plexus has few neurons that supply the muscle, but many that innervate the mucosa (Porter et al. 1999, Timmermans et al. 2001). In small mammals, typified by the guinea-pig, there is generally a single layer of submucosal ganglia, and these ganglia contain secretomotor neurons, but not motor neurons that supply the external muscle; in fact, in the guinea-pig there are four main types of neurons in the submucosal ganglia of the small intestine (Furness et al. 1984, 2003a). The ganglia of the submucosal plexus in small mammals most closely resemble those of the inner submucosal plexus of larger species. Paucity of ganglia in the submucosa of the esophagus and stomach Extensive submucosal ganglionated plexuses, such as those found in the small and large intestines, do not occur in the esophagus (Harting 1934, Schofield 1960, Rash & Thomas 1962, Christensen & Rick 1985a, Izumi et al. 2002). Small groups of nerve cell bodies are occasionally found adjacent to submucosal mucus-secreting glands that are scattered along the esophagus, although some investigators have reported that there are no nerve cell bodies at all in the submucosa of the esophagus (Christensen & Rick 1985a). Submucosal ganglia are absent or extremely rare in the stomach of small animals (guinea-pig and rat) and are sparse, but clearly present, in larger mammals, such as dog, human, and cat (Schabadasch 1930a, Kyösola et al. 1975, Stach et al. 1975, Radke et al. 1978, Christensen & Rick 1985a, Furness et al. 1991, Schemann et al. 2001, Colpaert et al. 2002) and denervation and tracing experiments show that the intrinsic innervation of the gastric mucosa is derived almost entirely from the myenteric ganglia (Furness et al. 1991, Pfannkuche et al. 1998). Submucosal nerve cells that do occur in the stomach are more common in the antrum. Some of the myenteric ganglia extend into the clefts (septa) between the large blocks of circular muscle in the stomach and can be mistaken for submucosal ganglia. Ganglia in the mucosa Small groups of nerve cell bodies occur in the lamina propria of the mucosa in the small and large intestine, and, rarely, in the stomach (Drasch 1881, Vau 1932, Stöhr 1934, Ohkubo 1936, Isisawa 1939, 1949, Lassmann 1975, Newson et al. 1979, Fang et al. 1993, Balemba et al. 1998). Stöhr (1934) has suggested that these are displaced (ectopic) submucosal ganglia. These nerve cells are almost always close to the muscularis mucosae, that is, they are close to the inner submucosal plexus.
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Subserosal plexus This is a plexus of fine nerve bundles that is found in the connective tissue layer at the surfaces of digestive organs, for example between the serosal lining of the peritoneal cavity and the external muscle of the intestine (Schabadasch 1930b). These nerve bundles connect extrinsic nerves and nerves of the deeper layers of the gut wall, as was recognized and described by Auerbach (1864). Small ganglia sometimes occur in the subserosal plexus, particularly in the esophagus and stomach and near the mesenteric attachment of the intestine and on the surface of the rectum. Some subserosal ganglia lie within or adjacent to the branches of the vagus nerves as they enter the walls of the stomach and esophagus. Longitudinal muscle innervation and the tertiary plexus The longitudinal muscle is innervated by a longitudinal muscle plexus, which consists of fine bundles of nerve fibers that run parallel to and within the muscle, or by the tertiary component of the myenteric plexus, which consists of axons in bundles that lie against the inner surface of the muscle (Richardson 1958). How the muscle is innervated seems to be simply determined by its thickness. In large animals, and in small animals where this muscle layer is thickened, for example in the teniae which occur in the large intestines of some species, a longitudinal muscle plexus is observed. Where the muscle layer is less than about 10 muscle cells thick, it is innervated exclusively by fine nerve bundles of the tertiary component of the myenteric plexus. These bundles are frequently found in small grooves at the inner surface of the muscle (Llewellyn Smith et al. 1993). The tertiary plexus is described in more detail above (see Fig. 1.5). Circular muscle innervation Fine nerve bundles that run parallel to the length of the muscle cells are found throughout the thickness of the circular muscle (Fig. 1.8). These bundles connect with the primary and secondary components of the myenteric plexus and with the deep muscular plexus in the small intestine. The nerve fiber bundles of the circular muscle plexus form a continuous meshwork both around the circumference of the intestine and, through oblique interconnecting nerve strands, along its length. In small mammals, most of the axons within the circular muscle plexus derive from motor neurons with cell bodies in the myenteric ganglia, but there are some fibers that come from nerve cells in the outer ganglia of the submucosal plexus. Fibers that originate from submucosal ganglia are more numerous in larger species (see above and Chapter 2).
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Fig. 1.8 Nerve fibers in the cir-
cular muscle. This micrograph is of a wholemount preparation of the circular muscle of the guinea-pig small intestine, stained with the Champy-Maillet zinc iodide and osmium technique. The level of focus corresponds to the deep muscular plexus. Major nerve fiber bundles run approximately parallel to the long axes of the muscle cells and there are many connections between these bundles. Calibration: 50 μm.
Deep muscular plexus and submuscular plexus An aggregation of nerve fiber bundles is found near the inner part of the circular muscle layer of the small intestine (Li 1937, 1940, Taxi 1965, Gabella 1972b, 1974) (Fig. 1.9) and also of the large intestine (Stach 1972, FaussonePellegrini & Cortesini 1984, Faussone-Pellegrini 1985, Christensen & Rick 1987b). These concentrations of innervation were described by Cajal (1895,
Fig. 1.9 Diagram to illustrate the nerve supply to the mucosa of the small intestine, as seen in histological section. The nerve fibers are in small bundles that form a continuous network in the connective tissue of the mucosa, the lamina propria (lp). The mucosal nerve network can be divided into interconnecting subglandular, periglandular, and villous components. Muscularis mucosae, mm; gland, gl.
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1911) who provided two descriptive names, plexus musculaire profond (deep muscular plexus) and plexus sous-musculeux (submuscular plexus) (Cajal 1911). A concentration of fibers near the inner surface of the circular muscle coat is not observed in the canine stomach (Furness et al. 1990a), although there is some degree of concentration of innervation in the human stomach (Faussone-Pellegrini et al. 1989). It is of interest that Cajal provided two names, because there is a subtle difference that these two names accommodate. In the small intestine, the plexus separates a thin layer of muscle cells from the bulk of the circular muscle, and here it has been generally referred to as the deep muscular plexus (Li 1937, Gabella 1974). In some regions, good examples being the dog and pig colon, there is no layer of muscle internal to the plexus, which lies at the extreme inner surface of the circular muscle, adjacent to the submucosa (Stach 1972, Christensen & Rick 1987b, Furness et al. 1990a). In this position, it can be called the submuscular plexus. The nerve bundles of the deep muscular and submuscular plexuses form continuous meshworks around the circumference and along the intestine. Their predominant orientation is parallel to the direction of the circular muscle, with frequent oblique connections between adjacent bundles (Fig. 1.8). The reason why part of the innervation of the circular muscle is concentrated close to its inner surface is not known, but the axons have the same spectrum of neurochemical types as axons in the rest of the circular muscle. It may simply be that the circular muscle is innervated asymmetrically, just as the innervation of arteries is asymmetric (in arteries, the axons are primarily at the outer border of the muscle coat) and the innervation of the longitudinal muscle through the tertiary plexus is asymmetric. Interstitial cells of Cajal (ICC) lie in close proximity to nerve fibers of the deep muscular plexus and they have a critical role as intermediates in transmission between the axons of motor neurons and the smooth muscle cells (Chapters 4, 5). Innervation of the muscularis mucosae The layers of smooth muscle at the surface of the mucosa, adjacent to the submucosa, are known as the muscularis mucosae. In general, this consists of inner bundles of circularly disposed smooth muscle cells and outer longitudinally oriented smooth muscle, but in some places it is thin and has smooth muscle bundles at various orientations. In the esophagus, the muscle bundles are arranged primarily in a longitudinal direction. Fine nerve fiber bundles that run parallel to the long axes of the muscle cells make up the innervation of the muscularis mucosae. In the small intestine, bundles of smooth muscle
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that are considered part of the muscularis mucosae make finger-like intrusions into the cores of the villi and in the stomach similar slivers of muscle are found between gastric glands. These intra-mucosal muscle bundles are also accompanied by nerve fibers. Mucosal innervation The structure of the mucosa varies more from one part of the gastrointestinal tract to another than do the structures of other layers. In the small intestine, it consists of the muscularis mucosae, the connective tissue (lamina propria), into which simple tubular glands protrude (the intestinal crypts, or glands of Lieberkuhn), and the villi. The lining of the glands and the surface of the villi are a single layer columnar epithelium. A dense network of fine interconnecting nerve bundles that is found throughout the connective tissue (lamina propria) of the mucosa (Fig. 1.9) makes up the mucosal innervation and was described in the 19th century (Billroth 1858, Drasch 1881, Müller 1892, Berkley & Baltimore 1893, Cajal 1895, 1911). The mucosal innervation in the small intestine can be divided into different components: a subglandular plexus, a periglandular plexus, a villous subepithelial plexus, and a plexus of the villous core. There is some specificity in the nerve fibers that contribute to the different components. For example, in guinea-pig small intestine, calretinin immunoreactive secretomotor neurons selectively supply the subglandular and periglandular components (Brookes et al. 1991a, Clerc et al. 1998b). In the stomach and colon, nerve fibers are found throughout the depth of the mucosa, and there is also a dense mucosal innervation in the gall bladder. A sparser plexus of nerve fibers occurs adjacent to the mucosal epithelium in the esophagus, which is a protective stratified epithelium that is devoid of secretory elements. The nerve fibers that innervate the mucosa lie in the connective tissue of the lamina propria, they do not penetrate the epithelium, which is a single layer in the stomach, small intestine, and colon. Some nerve fibers, which are believed to be sensory, penetrate the inner layers of the stratified epithelium that lines the esophagus (Rodrigo et al. 1975, Clerc & Condamin 1987). Nerve fibers in the mucosa inevitably come close to entero-endocrine cells of the gastric and intestinal epithelium. It is difficult to define any special relationship with the entero-endocrine cells, even using electron microscopy. Nevertheless, there are functional interactions between nerve fibers of the mucosal plexus and the endocrine cells (Chapter 2). Nerve fibers in the mucosa also come close to cells of the immune system, e.g. lymphocytes, and to the lymph nodules (Peyer’s patches) that occur in the mucosa (Chapter 2).
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Vascular innervation Two types of nerve fiber bundles are associated with arteries and arterioles of the alimentary tract (and of other organs). The first of these are paravascular nerve fibers which follow the arteries, using the tract taken by the arteries as a conduit. Paravascular fibers carry axons that are destined to supply other structures, as well as axons that innervate blood vessels. In the alimentary tract these other targets include the enteric ganglia, intestinal smooth muscle, and the mucosa. In addition, the arteries and arterioles are surrounded by a continuous meshwork of fine anastomosing nerve fiber bundles, the perivascular plexus, which contains both motor fibers to the blood vessels and vascular primary afferent (sensory) nerve fibers. Veins of the mesentery also have a perivascular plexus, but very few nerves have a perivascular relation with veins within the gut wall. Nerve fibers associated with intestinal lymphatic vessels are quite rare (Furness 1971). Nerve fibers associated with Brunner’s glands and glands in the esophagus The numerous small mucus-secreting glands in the submucosa of the duodenum, the glands of Brunner, are innervated by fine nerve fibers (Drasch 1881, Stach & Hung 1978, Ferri et al. 1984), as are the mucus-secreting glands in the esophagus. Functional studies suggest that the innervation of Brunner’s glands comes from the vagus, not from neurons of the enteric nervous system (Moore et al. 2000). The observation that vagal nerve stimulation promotes secretion of mucus, presumed to derive from Brunner’s glands, is an old one (Wright et al. 1940), but it is only this recent study (Moore et al. 2000) that suggests that the effect is not via the enteric nervous system. In the esophagus I have observed small ganglia in the submucosa adjacent to the glands, confirming earlier reports (Kadanoff & Spassowa 1959). Interconnections between the plexuses Although the plexuses have been described above as if they are separate entities, they are in fact joined by numerous nerve fiber bundles. Auerbach (1864) observed connections between extrinsic (vagal and mesenteric) nerves and the myenteric plexus via the subserosal plexus, and he also observed connections between the myenteric and submucosal plexuses. Drasch (1881) confirmed the connection of the myenteric plexus and the submucosal plexus and recognized that fibers from the submucosal plexus innervate the mucosa. Various authors detected fibers running from myenteric neurons into the circular muscle (Dogiel 1899, Stöhr 1930). Connections within the myenteric plexus, with one neuron sending a process to end on another myenteric neuron of
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the same or an adjacent ganglion were also described by several investigators (Dogiel 1895b, 1899, Kuntz 1922, Waddell 1929). Kuntz also found similar connections in the submucosal plexus. The fiber bundles that connect the myenteric and submucosal plexuses run almost perpendicular to the circular muscle layer. These have been called “penetrating fiber bundles” or “vertical fibers” (Furness et al. 1990a, Brehmer et al. 1998). Extent of the ganglionated plexuses Ganglia are found in the wall of the pharynx near its junction with the esophagus (Remak 1840, 1852, Shimazaki 1998). There are also small ganglia within the tongue (Remak 1852, Sbarbati 2002). I am hesitant to identify oral and pharyngeal ganglia as part of the enteric nervous system, because there is no evidence that they provide the type of local control that is exerted by the enteric nervous system. On the other hand, there is a well-developed myenteric plexus from the most oral part of the esophagus to the stomach in all species (Sabussow 1913, Abe 1959, Mann & Shorter 1964, Christensen & Robison 1982, Wu et al. 2003b). The myenteric plexus is then present continuously along the digestive tract until the internal anal sphincter (Schofield 1968). The ganglionated submucosal plexus is prominent and continuous from the first part of the duodenum to the level of the internal anal sphincter (Fig. 1.6). The numbers of nerve cells contained in the myenteric and submucosal plexuses have been estimated by counts of nerve cell bodies per unit area of gut surface (Furness & Costa 1987, Gabella 1987). Myenteric nerve cells are numerous throughout the digestive tube, varying in density from about 1000 to about 15 000 cell bodies per square centimeter. The lower values are found in the esophagus and proximal stomach, high values in the distal stomach and large intestine, and intermediate values in the small intestine. In total, the enteric nervous system of an individual mammal contains 2–1000 million nerve cells, depending on the size of the mature animal (Furness & Costa 1987, Gabella 1987, Karaosmanoglu et al. 1996). The total number of nerve cells in the myenteric plexus of the sheep small intestine has been estimated to be 31.5 million, in the guinea-pig, 2.75 million, and in the mouse, 403 000 (Gabella 1987). Total numbers in the submucosa of the small intestine were: sheep, 50 million; guinea-pig, 950 000; and mouse, 330 000. It has been estimated that the number of nerve cells in the enteric ganglia is about the same as the number in the spinal cord (Furness & Costa 1980). Although no direct counts have been made, the total number of enteric neurons in the human gastrointestinal tract, estimated by extrapolating data from other mammals, is probably in the range of about 200–600 million.
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Ganglia of the biliary system The biliary tract and the gall bladder develop from a diverticulum of the small intestine, and the ganglia that have been described in the walls of the gall bladder, cystic duct, hepatic duct, and common bile duct (Gerlach 1873, Dogiel 1895a, Harting 1930, Ottaviani 1941, Hermann 1951, Sutherland 1966, 1967, Cai & Gabella 1983, Mawe & Gershon 1989b) are regarded as part of the enteric nervous system. A ganglionated plexus lies external to the muscle coat in the gall bladder. Ganglia are found between the muscle bundles and internal to the muscle in the lamina propria of the mucosa (there is no muscularis mucosae in the gall bladder). In some species, for example the guinea-pig, nerve cells in the lamina propria are very rare (Sutherland 1966, Cai & Gabella 1983). The ganglionated plexus of the cystic duct, hepatic duct, and upper part of the common bile duct is also external to the muscle. In the part of the common bile duct closer to the duodenum, ganglia lie between the longitudinal and circular muscle layers and are also found internal to the musculature. There is continuity between the ganglionated plexuses of the biliary system and those of the duodenum (Cai & Gabella 1983, Mawe & Gershon 1989a, Padbury et al. 1993). Pancreatic ganglia Ganglia in the pancreas have been described by many authors (Cajal 1891, Müller 1892, De Castro 1923, Coupland 1958, Watari 1968). These ganglia are small and are connected to each other by nerve fiber bundles. They are probably best regarded as enteric ganglia because the pancreas develops as an outgrowth of the midgut, and because of their direct connections to duodenal ganglia (Kirchgessner & Gershon 1990). On the other hand, they can be looked upon as parasympathetic ganglia, analogous to the ganglia of the salivary glands, which are also secretory glands of the digestive tract. Ganglia in the trachea and bronchi The lungs develop as outgrowths of the foregut, and, like the digestive system, they have intrinsic ganglia, in this case in the walls of the trachea and bronchi (Fisher 1964). The ganglia form a well-developed plexus and appear to be a principal source of innervation of the airways of the lung. Nevertheless, the innervation of the trachea retains a peculiar relationship with the enteric nervous system into adult life: neurons with the phenotype of inhibitory neurons to gut muscle (immunoreactivity for VIP and NOS) project from the esophagus, where their cell bodies reside, to the trachea (Fischer et al. 1998; Moffatt et al. 1998). These neurons were identified by injecting a retrogradely
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transported dye into the trachea, which was subsequently detected in nerve cells in the esophagus. Intramural extensions of extrinsic nerves Branches of the vagus and pelvic nerves continue to be distinct and recognizable for considerable distances after they enter the walls of the esophagus, stomach, and rectum. The intramural pelvic nerves run along the large intestine, between the longitudinal and circular muscle layers, and can be traced at least as far as the transverse colon (Iljina & Lawrentjew 1932, Fan 1955, Lee 1956a,b, Christensen et al. 1984, Fukai & Fukuda 1984, McRorie et al. 1991). A good description is provided by Stach (1971). Myelinated fibers in the intramural nerves can be shown by degenerative section or tracing to arise from the pelvic nerves (Christensen & Rick 1987a). Branches of the vagus nerve enter the stomach at its junction with the esophagus and fan out to supply prominent intramural vagal branches that ramify at the level of the myenteric plexus (Mitchell 1940, Christensen & Rick 1985b, Wang & Powley 2000). Mesenteric nerve fiber bundles, containing sympathetic efferent axons and afferent nerve fibers, follow the small arteries into the gut wall and branch with them. They soon become very fine and cannot be recognized as distinct entities within the enteric plexuses. Electron microscope studies Ultrastructure analysis shows that enteric ganglia are remarkably compact, consisting of the cell bodies of neurons, supporting cells (enteric glia), and nerve cell processes (Richardson 1958, Hager & Tafuri 1959, Taxi 1959, 1965, Ono 1967, Baumgarten et al. 1970, Gabella 1972a, Cook & Burnstock 1976a, Yamamoto 1977, Wilson et al. 1981, Komuro et al. 1982b). A lowpower view typically shows nerve cell bodies and areas consisting of numerous axons with thin processes of enteric glial cells (Fig. 1.10). The enteric ganglia, unlike other autonomic ganglia, do not contain blood vessels, connective tissue cells or collagen fibrils. The absence of connective tissue and the close packing of neurons and glia give enteric ganglia an appearance similar to the central nervous system. The ganglia are not encapsulated, but lie in the connective tissue framework between the muscle layers or in the submucosa. Within the ganglia, individual nerve cells are only partly surrounded by glial cells. The surface elements of the ganglia have a thin layer of basal lamina and beyond this are fibroblasts and collagen fibrils which, although in parts aligned with the surface, form a discontinuous covering (Fig. 1.10). Thus the enteric ganglia receive their nutrient by diffusion from the blood vessels
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Fig. 1.10 The ultrastructure of enteric ganglia. A: At a low power, enteric nerve cells (N = neuronal nucleus) can be seen to present bare surfaces to the extracellular space (arrows). Glial cells are found in the ganglia, but only partly surround the nerve cells and axon bundles (G = glial cell nucleus). From Komoro et al. (1982). B: An area of neuropil adjacent to a nerve cell (N = neuronal nucleus) and a glial cell (G = glial cell nucleus) in a myenteric ganglion. The neuropil contains many profiles of axons (and perhaps dendrites), some of which contain transmitter vesicles, and some of which contain neurofilamants and neurotubules. Arrow points to a synapse. All axons are unmyelinated. From Gabella (1981).
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through the connective tissue that surrounds them. Tracers, such as peroxidase, injected into the blood stream diffuse to the ganglia and penetrate the clefts between their neuronal and glial elements (Jacobs 1977). Thus there is no effective blood–ganglion barrier, and drugs that do not enter the central nervous system have effects at enteric ganglia. It is common for enteric neurons to have large parts of their surfaces bare at the ganglion surface (Gabella 1972a, Komuro et al. 1982b). Sections through myenteric nerve cell bodies reveal profiles that vary considerably in size and in fine structural detail. Cook and Burnstock (1976a) distinguished eight types of neuron within the myenteric ganglia of the guinea-pig small intestine. However, how these relate to functional types is unknown. The only neurons in which ultrastructure and cell type have been clearly related are Dogiel type II neurons, which contain large numbers of mitochondria (Chapter 2). However, neurons in submucosal ganglia of the guinea-pig small intestine do not vary much in their ultrastructural features and, in conventional electron microscopy, appear to belong to one class (Wilson et al. 1981), although of course this is not true in a functional sense. It seems that all nerve cells in the myenteric and submucosal plexuses receive ultrastructurally recognizable synaptic inputs (Cook & Burnstock 1976a, Wilson et al. 1981, Komuro et al. 1982a, Pompolo & Furness 1988). The synapses typically show pre- and post-synaptic densities and presynaptic accumulations of transmitter storage vesicles (Fig. 1.11), similar to synapses elsewhere. There are also frequent close appositions, without pre- and post-synaptic densities,
Fig. 1.11 Enteric axon varicosities containing transmitter vesicles. A: An axon (Ax) forming a synapse on a myenteric nerve cell (N). There are both pre- and post-synaptic densities at the synapse (arrows). B: An axon (Ax) at the surface of a nerve fiber bundle within the muscle coat. The axon contains transmitter vesicles and is bare at the surface that faces the muscle (m). Axons within the nerve bundle are only partly separated by the processes of glial cells (G). Both micrographs from Gabella (1972).
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between synaptic vesicle-containing axonal profiles and nerve cells. Bare surfaces of axons are encountered where nerve bundles come close to other effectors, such as smooth muscle cells, the interstitial cells of Cajal, or the mucosal epithelium (Fig. 1.11) (Gabella 1972a). In general, presynaptic specialization is not recognized at these close approaches, although rare instances of presynaptic densities have been described (Klemm 1995). Enteric glia The enteric ganglion cells and nerve fiber tracts are supported by numerous glial cells. Nucleated satellite cells (glial cells) around nerve cell bodies of the enteric ganglia were described by Dogiel (1899), and many subsequent light microscopists pointed out these cells, both in the ganglia and nerve strands of the gastrointestinal plexuses, and in general referred to them as Schwann cells (see Stöhr 1952). More recent studies, which have used immunohistochemical markers to locate enteric glia, reveal that they are common in the ganglia and nerve fiber bundles. Enteric glia express glial fibrillary acidic protein (Jessen & Mirsky 1980, 1983) and the S-100 Ca2+ binding protein (Ferri et al. 1982b), both of which are typical of central nervous system astrocytes. Glial cells in other autonomic ganglia do not contain glial fibrillary acidic protein (Jessen & Mirsky 1983). Electron microscope studies also show that enteric neuronal satellite cells resemble glial cells or astrocytes of the central nervous system more than Schwann cells of other peripheral ganglia or nerve trunks (Gabella 1971, Gabella 1972a, Cook & Burnstock 1976b, Gabella 1981b, Komuro et al. 1982b). The glial cells partly surround nerve cell bodies and axons in the ganglia, leaving bare large areas of neuronal membrane at the surfaces of ganglia. There is a marked contrast in relationships of glial cells to axons in small mammals (for example, guinea-pig or rat) and large mammals (such as cat or human). In small mammals, glial cell processes fail to penetrate all the interstices between nerve cell bodies and between axons in the neuropil (Gabella 1972a, Komuro et al. 1982b). In fact, many nerve processes are in direct membrane-to-membrane contact with each other; the glial cells only separate them into groups and rarely form a sheath around an individual axon. In contrast, in enteric ganglia of human and monkey, axons are separated from one another by intervening glial cell processes (Baumgarten et al. 1970). It is noteworthy that Auerbach (1864) had recognized the differing relations of neurites and supporting cells between humans and some other mammals. Glial cells in nerve strands of the myenteric plexus of small mammals give rise to radiating lamellae which divide the axons into large bundles, and up to 600 neurites may be associated with one glial cell (Gabella 1981b). The ratio of numbers of glial cells to nerve cells increases with species size, in myenteric
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ganglia from 1.1 in mice to 4.5 in sheep and in submucosal ganglia from 0.6 to 1.5 (Gabella & Trigg 1984). Glial cell nuclei can readily be distinguished from those of neurons (Fig. 1.10). The glial nuclei have conspicuous clumps of chromatin, particularly adjacent to the nuclear envelope, and the nuclear surface often displays deep invaginations. The most prominent distinguishing feature of the majority of enteric glial cells is the numerous 10 nm gliofilaments. The cytoplasm also contains smooth and rough endoplasmic reticulum, numerous free ribosomes, mitochondria, lysosomes, and microtubules. Groups of gliofilaments criss-cross the cell bodies and run parallel to the long axes of the cell processes. The gliofilaments appear to be anchored to dense aggregations of material adjacent to the surface membrane. There are some points of apposition between glial cells and between glial cells and neurons where there are small areas of cytoplasmic density inside each plasma membrane. Gap junctions between enteric glia are not often seen, but dye filling indicates that they are coupled to each other (Hanani et al. 1989). The structural similarities and functional differences between regions may have an evolutionary basis A nervous system embedded in or surrounding the gut tube is found throughout the animal kingdom, and is seen for example in simple animals such as Hydra (Hansen et al. 2002, Shimizu et al. 2004). It is thought that the ganglia that form the primitive brains of helminths, and eventually the brains of higher animals, as well as chains of ganglia that provide body segments, were derived from the nervous system around the gut tube (Bullock & Horridge 1965). However, at the same time that primitive brains in the form of cephalic ganglia evolve, there is development of a recognizable enteric nervous system (Fig. 1.12), for example in insects (Ganfornina et al. 1996), leeches (Ábrahám & Minker 1958), octopus (Alexandrowicz 1928), and snails (Campbell & Burnstock 1968, Röszer et al. 2004). Perhaps the most primitive enteric nervous system is that of the marine polyp Hydra (Shimizu et al. 2004). In this species the nodes of the plexus contain only single nerve cells. Nevertheless, when the enteric nervous system in Hydra is ablated, peristalsis, mixing movements and defecation are severely compromised (Shimizu et al. 2004). Although there is a primitive gut tube with little regional specialization in typical invertebrates, the mammalian gut exhibits distinct regional specializations. In turn, the enteric nervous system has evolved to control the functions of the different regions of the mammalian digestive system. So, although the general arrangement of the myenteric ganglia looks similar in all regions of the tubular digestive tract, reflecting a common origin, its functions show important specializations. In the striated muscle part of the esophagus, the
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A
Fig. 1.12 The enteric nervous system in the octopus. A: A ganglion of the superficial plexus. This plexus occurs in the connective tissue at the surface of the intestine, a similar location to the subserous plexus in mammals. B: Cross-sectional drawing of the enteric plexuses of cephalopods. Ganglionated plexuses are found at the serosal surface of the intestine and in the external muscle. Cm: circular muscle; Imp: intramuscular plexus; Lm: longitudinal muscle; lp: lamina propria; Epi: epithelium lining the gut tube. From Alexandrowicz (1928).
enteric nervous system is extensive, but it has a limited control over organ function compared to the enteric nervous system of the small intestine. In fact, the motor functions of the striated muscle esophagus are largely controlled through motor pattern generators in the brainstem (Chapters 3, 5), although enteric neurons may have modulatory roles (Chapter 2). By contrast, the enteric nervous system of the small intestine has substantial roles in the intrinsic control of its movements, fluid exchange with the lumen, and local blood flow. It is as if the enteric nervous system of the striated muscle part of the esophagus has retained a structural complexity that is excessive to the needs of the organ. In the stomach, the enteric nervous system has evolved to influence gastric acid secretion and movement, while considerable influence
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over these functions is also exerted through the vagus. In fact, the sophisticated intrinsic reflex control of organ function that occurs in the small and large intestines is less well developed in the stomach (see Chapter 5). Development of the enteric nervous system The enteric nerve cells are not generated in the gut, they arise from precursor cells that migrate from the neural crest. One of the first investigations of the origin of enteric ganglia was that of Kuntz (1910) who examined the precursor and developing ganglia of the sympathetic chains, sympathetic pre-vertebral ganglia, cardiac plexuses, and the gastrointestinal tract in pig embryos of different ages. He concluded that the enteric neuron precursors reach the developing gut by migration along the vagus. This conclusion is essentially correct (Gershon et al. 1993, Young & Newgreen 2001, Gershon 2002). Experimental confirmation of an origin from the vagal neural crest was obtained by Jones (1942) and Yntema and Hammond (1954) who removed parts of the developing neural crest from early chick embryos. Yntema and Hammond (1954) found that removal of rostral regions, but not removal of caudal regions, prevented the development of enteric ganglia throughout the gut, and concluded that vagal level neural crest cells were the source of enteric ganglia. There was initial disagreement about a sacral source in birds: one investigator reported that removal of sacral level neural crest diminished the formation of ganglia in the hindgut (Jones 1942), and it was soon after argued that this observation was mistaken (Yntema & Hammond 1954). Ablation of part of the developing chick is a dramatic intervention that might interrupt signaling in the embryo even if the enteric neurons had a non-vagal origin. An elegant solution was devised by Le Douarin who transplanted quail neural crest into chickens from which the equivalent region was removed (Le Douarin & Teillet 1973, 1974). Quail cells have a nuclear marker that is absent in chick, and these experiments showed that vagal neural precursors migrate to the gut, to form enteric neurons and glial cells throughout its length, and that the precursors from sacral sources contribute to neural populations in the hindgut. In mammals there is also a small contribution of sacral crest cells to the colonization of the rectum (Kapur 2000). In both species, the entry of cells of sacral origin does not occur until vagal cells have arrived. Moreover, if the vagal source is eliminated, the progeny of sacral cells that enter the gut of the chick remain in the hindgut, the rest of the gastrointestinal tract being uncolonized (Burns et al. 2000). In mammals, most recent work has been conducted in mouse, using excellent markers of the early partly differentiated neurons. A wave of migrating cells flows down the gut, from the foregut to the end of the hindgut (Fig. 1.13) in the period between embryonic days 9.5 and 14.5 (Kapur et al. 1992, Young et al. 1998). As the wave advances, maturing
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Fig. 1.13 The developing enteric nervous system. The intestine of an embryonic day 10 mouse is shown, with neurons revealed by staining for the transcription factor Phox2b. At this stage, myenteric neurons extend throughout the foregut and the furthest migrated nerve cell is in the midgut (arrow), but no cells have yet appeared in the hindgut. From Young et al. (1998).
cells are halted in their progress and subsequently form ganglia. Thus behind the wave front there is a maturing enteric nervous system. Substantial advances have been made in identifying the signaling mechanism involved in determining the migration and maturation of enteric neurons and glial cells, primarily through the investigation of transgenic animals, but also through the investigation of the genetic basis of disorders of enteric nervous system development (Gershon 2002, Newgreen & Young 2002a,b, Young et al. 2004). One of the first clues came from genetic analysis of Hirschsprung’s disease, a disorder in which enteric nerve cells fail to colonize the gut completely. Most commonly, enteric ganglia are missing from the rectum and sigmoid colon, but the absence of enteric ganglia is sometimes more extensive (Newgreen & Young 2002a, Swenson 2002). About half of the cases are attributable to abnormalities of a cell recognition and signaling complex that consists of the membrane inserted receptor-tyrosine kinase, Ret, and its partner molecules, glial-derived neurotrophic factor (GDNF) and GFR 1 (Gershon 2002, Newgreen & Young 2002a). Nerve cells only occur in the first part of the foregut (that will become the esophagus) of mice in which the c-ret gene that encodes Ret or the gene for GDNF is knocked out (Schuchardt et al. 1994, Moore et al. 1996, Pichel et al. 1996). The reciprocal defect, a failure of the development of esophageal ganglia, occurs when the transcription factor Mash-1 is inactivated (Guillemot & Lo 1993). Mash-1 knockout also prevents the development of a subset of neurons in the gastrointestinal tract beyond the esophagus, so although this region is colonized, it is not normal (Gershon 2002). Another regulatory system that controls the development of the enteric nervous system is the endothelins (ETs) which bind to the cell surface receptors
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ETA and ETB (Newgreen & Young 2002a). In the enteric nervous system, the important ET form appears to be ET3, and the receptor is ETB. Current evidence is consistent with the theory that ET3 is produced by the developing mesenchyme, and that the ETB receptor is on enteric neurons (Kapur et al. 2004). Mutations in the ET and ET receptor genes cause about 5% of Hirschsprung’s disease and distal bowel aganglionosis in the piebald and lethal spotting mouse and the spotting lethal rat. Thus defects in endothelin or its receptor results in incomplete colonization of the bowel. It has been suggested that the defects cause premature differentiation of the precursors (Hearn et al. 1998, Wu et al. 1999), thus limiting the availability of proliferating and migrating neuron and glial cell precursors to colonize more distal gut. However, the complexity of the interactions of neurons and their environment during the development of the enteric nervous system suggests a number of other possible mechanisms (Gershon 2002). Transcription factors that initiate enteric neuron differentiation have been identified. If the genes for either Phox2b or Sox10 are knocked out, no neurons enter the gut (Newgreen & Young 2002a). Other genes that influence the development of the enteric nervous system have been identified, and, although their roles are not yet accurately defined (Gershon 2002, Newgreen & Young 2002a, Young et al. 2004), it is likely that they soon will be in this rapidly advancing field. The precursors of enteric neurons must supply the 15 to 20 individual types that are present in the enteric nervous system. Therefore signaling mechanisms must exist that influence the final differentiation, the density of neurons, the connections that they make, the placement of myenteric neurons between the external muscle layers, and the positioning of other neurons in the submucosa. The molecules that have so far been identified in directing the proliferation, migration, and differentiation are those whose deficiency or manipulation cause substantial changes in enteric neuron number or migration; the control of the final differentiation, positioning, and connectivity of enteric neurons has yet to be unravelled. Development of enteric neurons of the submucosal plexus, pancreas, and gall bladder The biliary system and pancreas develop as outgrowths from the gut, and the similarities in the structure of the neurons within these organs would suggest that they derive from enteric neural precursor cells. In mice, the submucosal plexus is colonized 2–3 days after the myenteric plexus at the same level of the intestine, and might therefore be assumed to derive from the myenteric plexus. In human, the delay in formation of the submucosal plexus is greater, lagging behind the myenteric plexus by 2–3 weeks (Wallace & Burns 2005). These assumptions have been tested experimentally, leading to the
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observation that myenteric neural precursor cells migrate from the myenteric plexus to the submucosa and pancreas under the influence of netrins and the netrin receptor, DCC (Jiang et al. 2003). Maturation of enteric neurons and development of function Development of function in the enteric nervous system begins in fetal life and continues for some time post-partum. This period varies considerably between species, amongst other things depending on the development program of the species, and the maturity at birth. In human, there is informative data from studies on aborted fetuses, fetuses in utero and from investigation of pre-term infants (Milla 1993). Neural crest cells enter the human foregut at about week 4, are observed in the developing stomach at 6 weeks, and the myenteric plexus is identifiable in the small intestine by about 8–9 weeks, the submucosal plexus is present 2–3 weeks later, and by around 12–14 weeks the gut has developed its adult form and pattern of enteric ganglia (Milla 1993, Wallace & Burns 2005). Cyclic regular electrical activity and propulsive activity occurs in the small intestine in human, sheep, and dog at about 70–80% of the full term of gestation (Buéno & Ruckebusch 1979, Milla 1993), which suggests that the enteric nervous system in the small intestine is functional at this time. In the lamb, mature migrating myoelectric complexes (MMC; Chapter 5) were seen at 10 days before birth, whereas in the dog they commenced 15 days postnatal. For premature human infants born at 34 weeks or less, there is a lack of co-ordinated activity of swallowing and esophageal motility, but within about a further week the infant sucks, and the motility of the stomach and small intestine shows features of the mature pattern, including the presence of the MMC and postprandial patterns of movement that are qualitatively similar to the adult (Berseth 1989, Milla 1993). The MMC has a shorter cycle time in the fetus and for the first few weeks after birth. In mice, which are considerably less mature at birth, movement of fluid in an anal direction is independent of enteric neurons to the end of gestation, but if the neurons are lacking the mice die soon after they are born (Anderson et al. 2004). Effective neuromuscular transmission from excitatory and inhibitory motor neurons to the muscle of the small intestine is seen for rabbit at E17 (embryonic day 17; gestation 29–35 days, usually 31 days) and in the mouse at E17 (Gershon & Thompson 1973). Consistent with the maturation of the function of the enteric nervous system in utero, evidence of neuronal differentiation, such as the expression of neurotransmitter systems, occurs early. In mice at embryonic day 10 (E10), enzymes that reduce NADPH were seen in neurons adjacent to the stomach (Branchek & Gershon 1989). The NADPH reductase activity is probably due
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to the presence of nitric oxide synthase (NOS), the synthesizing enzyme for the enteric transmitter nitric oxide (NO). At E12, the enteric neuropeptide NPY was observed in enteric neurons. Other neural markers that appear during fetal life include choline acetyltransferase, 5-HT, CGRP, enkephalin and VIP (Pham et al. 1991). The early appearance of NADPH reductase/NOS is consistent with the observation that some neurons immediately behind the advancing front of migrating neurons express NOS (Young et al. 2002). By electron microscopy, identifiable synapses between enteric neurons are first seen in the small intestine at E38 (term is 55–60 days) in the guinea-pig (Gershon et al. 1981), indicating that nerve circuits were beginning to be assembled 60–70% through gestation. In mouse intestine (gestation 21 days), synapses with a mature appearance were first detected at E16.5 (Vannucchi & Faussone-Pellegrini 2000), although primitive synapse-like structures were transiently present between 12.5 and 16.5 days. Structural studies confirm the functional data that maturation of the enteric nervous system continues beyond birth (Matini et al. 1997). Changes in enteric neurons with aging Considerable loss of enteric neurons may occur in animals and human towards the ends of their lifespan. Between 6 and 24 months of age in the rat, the numbers of myenteric neurons stained by the NADH diaphorase method declined by 40% in the jejunum and 63% in the colon (Santer & Baker 1988). This method is general, and was thought at the time to stain all neurons. It is a histochemical method that detects NAD:NADH oxidoreductase of the respiratory chain and should detect metabolically active neurons. The guinea-pig small intestine harbors 2.75 million NADH diaphorase positive neurons in the young adult (3–4 months old), which falls to 1.1 to 1.6 million in guinea-pigs aged 26–30 months (Gabella 1989). Moreover, the nerve cell bodies in guinea-pigs became smaller and their surfaces were more angular. In contrast, in rats maintained on a restricted diet, no decline in myenteric neuron numbers was observed between 4 and 24 months (Johnson et al. 1998b). Given these contrasting results, a comparison of rats on a free-feeding diet and a food-restricted diet were compared (Cowen et al. 2000). Rats on an unrestricted diet ate an average of 47 g of rat food per day and lost about 50% of myenteric neurons, whereas rats restricted to 25 g per day had the same numbers of neurons at 4 and 24 months. The loss was selective for cholinergic neurons (neurons that stained for choline acetyltransferase) over NOS neurons. There is also evidence for enteric neuron loss with age in human (Wade & Cowen 2004), although it is not known whether this is diet related.
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Summary and conclusions The enteric nervous system is composed of thousands of small ganglia (most being contained in the myenteric and submucosal nerve plexuses), the nerve fibers that connect the ganglia, and nerve fibers that supply the muscle of the gut wall, the mucosal epithelium, arterioles, and other effector tissues. The ganglia contain nerve and glial cells, but not connective tissue elements, and in many respects are similar in structure to the central nervous system. Nerve fiber bundles within the enteric nervous system consist of the axons of enteric neurons, axons of extrinsic neurons that project to the gut wall, and glial cells. The myenteric plexus forms a continuous network, extending from the upper esophagus to the internal anal sphincter. The ganglionated submucosal plexus is present in the small and large intestines, but is absent from the esophagus and contains only very few ganglia in the stomach. Large numbers of neurons are contained in the enteric nervous system, about 200–600 million in human. This is far more neurons than occurs in any other peripheral organ and is similar to the number of neurons in the spinal cord. Neuronal plexuses occur around the gut tube from the simplest animals, such as Hydra, to the most advanced mammals. In mammals, the general structures of the enteric ganglia are comparable throughout the gut, although their functional roles have diverged. The enteric nervous system originates from neural crest cells that colonize the gut during intra-uterine life. It becomes functional in the last third of gestation in human, and continues to develop following birth. There is some loss of enteric neurons in mammals of advanced age.
2: Constituent neurons of the enteric nervous system
Enteric neurons have been classified by their shapes, their physiological properties, specific histochemical, and immunohistochemical staining, and other distinguishing features, including the structures that they innervate, the transmitters they utilize, and the connections that they receive. One of the major advances of the last 15–20 years has been success in correlating most of these features, so that characterizing profiles of all functionally identified neurons in the guinea-pig ileum have now been established (Table 2.1; Fig. 2.1). Enteric neurons can be grouped by their functions as intrinsic primary afferent neurons, interneurons, and motor neurons. Intrinsic primary afferent neurons (IPANs) are sometimes referred to as “intrinsic sensory” neurons, a terminology that is discussed below. Although extensive data on each neuron type has only been obtained for the guinea-pig, the enteric nervous system performs the same functions in all mammalian species, and therefore all species have essentially the same functionally defined neurons. For example, all mammals have excitatory and inhibitory motor neurons that innervate the muscle of the gastrointestinal tract. There are sufficient data to identify in other species the equivalent neurons (orthologs) that have been identified by function in the guinea-pig. One clue is shape. It is generally true that neurons that serve the same functions in different species have the same shape (Peters et al. 1991), a principle that appears to apply to the enteric nervous system (Brehmer et al. 1999a, Furness et al. 2004b). Examples of neurons that have the same shapes and serve the same functions in different species are motoneurons in the ventral horns of the spinal cord, cerebellar Purkinje cells, pyramidal cells in the cerebral cortex, retinal ganglion cells, and dorsal root ganglion cells. It is thus justified to use morphology as a first indicator of the equivalence of neurons in different species. Other clues are chemical properties related to the transmitters that neurons utilize, and projections to targets. In relation to shape, it should be noted that there is a tendency for equivalent neurons to have more elaborate dendritic trees in larger animals, and less elaborate dendrites in smaller mammals (Purves & Lichtman 1985, Purves et al. 1986, Tabatabai et al. 1986). 29
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Fig. 2.1 A: The types of neurons in the small intestine of the guinea-pig, all of which have been defined by their functions, cell body morphologies, chemistries, key transmitters, and projections to targets. The numbers adjacent to the neurons correspond to the numbers in Table 2.1, which lists each of the neuron types by their functions and provides data on their chemistries and the percentages of their cell bodies in the myenteric or submucosal ganglia. LM: longitudinal muscle; MP: myenteric plexus; CM: circular muscle; SM: submucosal plexus; Muc: mucosa. Note that the muscularis mucosae (MM) is very thin in the guinea-pig small intestine and the neurons that innervate this muscle have not been identified in this species. In other species they are in the submucosal ganglia. B: Types of neurons in the small intestine of the pig. In many cases the equivalent neurons have been identified in the pig and guinea-pig, and they have been given the same numbers. A, adapted from Furness et al. (2004a); B, modified from Timmermans et al. (2001).
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For example, in the enteric nervous system, Dogiel type I neurons (Fig. 2.1) have considerably less elaborate dendrites in mice (Nurgali et al. 2004) than in larger mammals, such as guinea-pigs, pigs, dogs, and humans (Dogiel 1899, Stach 1980, Furness & Costa 1987). In general, enteric neurons are smaller, simpler in form, and easier to classify, in small mammals (e.g. in guinea-pigs) and become more elaborate, and more difficult to classify, in larger species of animal, including humans (Brehmer et al. 2004a). Table 2.1 includes data on the chemical coding of the different classes of enteric neurons in the guinea-pig small intestine. Most of these data were gathered over a period of around 20 years, from about 1975 to 1995, and have been summarized and reviewed in a number of papers (Furness et al. 1984, 1995b, Costa et al. 1996, 2000). Subsequently, CART (cocaine and amphetamine-regulated transcript peptide) has been identified as a constituent of a minority of myenteric neurons, and of the VIP-containing secretomotor/vasodilator neurons (Ellis & Mawe 2003). These neurons also contain corticotrophin-releasing factor (CRF) (Liu et al. 2005). Another peptide that, like CART, is involved in regulation of feeding, orexin, is found in enteric neurons (Kirchgessner & Liu 1999a). Orexin occurred in IPANs in the myenteric and submucosal plexusus, and in about 25% of VIP-immunoreactive submucosal neurons. Two further chemical markers for IPANs have also been identified recently, these being the neuronal nuclear protein, NeuN, which occurs in both the nucleus and cytoplasm of IPANs (Chiocchetti et al. 2003) and binding sites for isolectin B4 from Bandiera simplificifolia (Hind et al. 2005), which binds specifically to α-D-galactose end-groups of glycoproteins (Laitinen 1987). Both cytoplasmic NeuN and IB4 binding appear to be specific to IPANs, and to label the entire population of these neurons in the guinea-pig ileum. Shapes of enteric neurons The first and most enduring classification of enteric neurons by their shapes was that of Dogiel (Figs 2.2, 2.3, 2.7). He provided a comprehensive description of neuron morphologies in the myenteric and submucosal plexuses of the intestine from human, guinea-pig, rabbit, rat, dog, and cat (Dogiel 1895b, 1899). The majority of his illustrations are of guinea-pig and human samples. He described three neuron types, now generally referred to as Dogiel types I, II, and III. Between the publication of his first paper and the more comprehensive publication of 1899, La Villa (1898) published observations that revealed cell types in the rabbit intestine (Fig. 2.4) that corresponded closely to those described and depicted by Dogiel.
<1% N/A
*Motor neurons to gut endocrine cells
26%
Dogiel type I
Dogiel type I
Dogiel type III (filamentous) Dogiel type II
4%
ChAT/ TK/ orexin/ IB4/ NeuNcyt/ NK3 receptor, most calbindin ChAT/ BN/ VIP/ NOS/ CCK/ ENK N/A
ChAT/ SOM
ChAT/ 5-HT
ChAT/ Calretinin/ TK/ ENK ChAT/ NOS/ VIP ± BN ± NPY
Large Dogiel type I Dogiel type I Dogiel type I
NOS/VIP/GABA
2%
Intestinofugal neurons (3)
Descending interneurons (secretomotor and motility reflex) (9) Descending interneurons (migrating myoelectric complex) (10) Myenteric intrinsic primary afferent neurons (IPANs) (2)
~ 2% Inhibitory longitudinal muscle motor neurons (5) Ascending interneurons (local reflex) (1) 5% Descending interneurons (local reflex) (8) 5%
Short: ChAT/ TK/ ENK/ GABA Long: ChAT/ TK/ ENK/ NFP Short: NOS/ VIP/ PACAP/ ENK/ NPY/ GABA Long: NOS/ VIP/ PACAP/ dynorphin/ BN/ NFP ChAT/calretinin/ TK
Chemical coding
Small Dogiel type I (small simple neurons) Dogiel type I
Dogiel type I
16%
25%
Dogiel type I
12%
Excitatory circular muscle motor neurons (6) Inhibitory circular muscle motor neurons (7)
Excitatory longitudinal muscle motor neurons (4)
Proportion Shape
Functional definition
Table 2.1 Types of neurons in the enteric nervous system. (a) Myenteric neurons.
For example, neurons innervating gastrin cells
Primary transmitter ACh, co-transmitter VIP
Primary transmitters TK and AChCGRP and ACh in other species
Several co-transmitters with varying prominence: NO, ATP, VIP, PACAP Primary transmitter ACh, co-transmitter TK Primary transmitter ACh, ATP may be a cotransmitter Primary transmitters ACh, 5-HT (at 5-HT3 receptors) Primary transmitter ACh
Primary transmitter ACh, co-transmitter TK
To all regions, primary transmitter ACh, cotransmitter TK Several co-transmitters with varying prominence: NO, ATP, VIP, PACAP
Transmitter/comments
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Uni-axonal with thin dendrites Dogiel type II
29% 1%
Dogiel type I Dogiel type I
N/A N/A
Transmitter/comments
NOS/VIP
ChAT/ TK
ChAT/ TK/ orexin/ IB4/ NeuNcyt/ calbindin
Pharmacology of transmission appears to be similar to other enteric inhibitory muscle motor neurons
Calbindin-IR, seen with some antisera only. Primary transmitter ACh, may be TK contribution. CGRP and ACh in other species Primary transmitter ACh
ChAT/ NPY/ CCK/ SOM/ CGRP/ Primary transmitter ACh. A small proportion of dynorphin these have cell bodies in myenteric ganglia VIP (NOS?) Possibly displaced myenteric interneurons
VIP/ CART/ CRF/ GAL/ PACAP/ Primary transmitter VIP. A small proportion of NMU. In humans, these these have cell bodies in myenteric ganglia neurons may be cholinergic ChAT/ calretinin/ dynorphin Primary transmitter ACh
Chemical coding
This table lists the neuron types that are found in the guinea-pig small intestine, some of their defining characteristics, and percentages of occurrence in each of the ganglionated plexuses. Also listed are three types of motor neuron found in other parts of the tubular digestive tract, marked by asterisks*. The numbers in parentheses are the identifying numbers for the neurons in Fig. 2.1. ACh, acetylcholine; BN, bombesin (the mammalian form also referred to as GRP, below); CCK, cholecystokinin; ChAT, choline acetyltransferase; CART, cocaine and amphetamine-regulated transcript peptide; CGRP, calcitonin gene-related peptide; CRF, corticotrophin-releasing factor; ENK, enkephalin; GABA, gamma amino butyric acid; GAL, galanin; GRP, gastrin-releasing peptide (mammalian bombesin); 5-HT, 5-hydroxytryptamine; IB4, isolectin B4; MMC, migrating myoelectric complex; NeuNcyt, cytoplasmic immunoreactivity for the neuronal nuclear protein NeuN; NFP, neurofilament protein NK, neurokinin; NOS, nitric oxide synthase; NPY, neuropeptide Y; PACAP, pituitary adenylyl cyclase activating peptide; SOM, somatostatin; TK, tachykinin; VIP, vasoactive intestinal peptide.
*Excitatory motor neurons to the muscularis mucosae *Inhibitory motor neurons to the muscularis mucosae
Type IV
15%
Cholinergic secretomotor/ vasodilator neurons (13) Cholinergic secretomotor (non-vasodilator) neurons (14) Uni-axonal neurons projecting to the myenteric plexus (15) Submucosal intrinsic primary afferent neurons (IPANs) (11) 11%
Stellate
45%
Non-cholinergic secretomotor/ vasodilator neurons (12)
Dogiel type I
Proportion Shape
Functional definition
(b) Submucosal neurons
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Dogiel type I neurons Dogiel described type I cells as flattened, slightly elongated, with stellate or angular outlines, when viewed in wholemounts (Fig. 2.2). Cell bodies were between 13 and 35 μm in length and 9–22 μm in width. The cells had from 4 to 20 or more dendrites and one axon. A defining feature of these neurons is their lamellar dendrites that are flattened in the plane of the myenteric plexus (Dogiel 1899, Lawrentjew 1929), which in most cases extend for short distances from the cell body (see Fig. 2.2). Lamellar dendrites give rise to several short subsidiary expansions (see, for example, Fig. 2.2f). In addition, some neurons have mushroom-shaped protrusions from their cell bodies that extend above and below the plane of the myenteric plexus (Costa et al. 1982, Furness et al. 1989b). Dogiel wrote that the axons of type I cells run out of the ganglia and continue through up to four other ganglia before entering the circular muscle coat. This is correct for many of the Dogiel type I neurons, but not all Dogiel type I neurons have axons that enter the circular muscle.
Fig. 2.2 Examples of Dogiel type I neurons as defined and drawn by Dogiel (1899). Neurons A, C, E, F, and H are from the myenteric plexus of the guinea-pig small intestine, neurons D and G are from the myenteric plexus of human small intestine and neuron B is from dog gall bladder.
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Modern methods, utilizing cell filling and immunohistochemistry, as well as improved silver impregnation techniques, have fully confirmed Dogiel’s description of type I neurons as a distinct neuron type in all species studied, including cat, dog, guinea-pig, human, mouse, pig, rabbit, rat, and sheep (Brehmer et al. 1999a). These neurons do not belong to a single functional class; some are inhibitory motor neurons to the muscle, some are excitatory motor neurons and some Dogiel type I neurons are interneurons. Axons occasionally bifurcate close to their origin, and often give rise to short spines in their initial parts, usually the first 100 μm. The axons usually arise from the cell body, but sometimes emerge from a dendrite. Despite Dogiel’s clear description of type I neurons as having short, broad, flat dendrites, some authors have included in Dogiel’s type I classification neurons that have short, fine or tapering dendrites that clearly belong to his type III or to the type IV neurons that are described below (compare Figs 2.2, 2.5, and 2.9). At least some of this drift in terminology derives from the work of Hill (1927) in which she defines cells as type I which she states are probably identical with type III. Subsequent authors have illustrated cells as type I which belong to Dogiel’s type III classification or the later defined type IV category (for example, van Esveld 1928, Iwanow 1930, Sokolowa 1931, Murat 1933, Cavazzana & Borsetto 1948). Dogiel type II neurons Dogiel type II neurons (Fig. 2.3) are the most prominent neurons in myenteric and submucosal ganglia of the small intestine and colon, but are very rare in the stomach. They have large round or oval cell bodies and they are numerous, usually 10–25% of the neuron population in the myenteric ganglia of small and large intestines. Dogiel described them as having 3–10 dendrites and one axon. It is possible that Dogiel believed that neurons could only have one axon. However, it is now recognized that the principal processes of these neurons are all axons, both on morphological and on functional grounds (Stach 1981, Hendriks et al. 1990). In some cases, Dogiel type II neurons have a single process that branches into subsidiary axons at a short distance from the cell body; these are usually called pseudounipolar neurons. Dogiel and later investigators generally depict the cells as having a smooth surface. However, when they are examined optically at high resolution, or by electron microscopy, the cell surfaces are seen to be irregular, for example being grooved by nerve fiber bundles at the cell surface (Pompolo & Furness 1988). Dogiel reported that the cells had major diameters of 22–47 μm and minor diameters of 13–22 μm, which is in the range that current observers also report.
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Fig. 2.3 Examples of Dogiel type II neurons as defined and drawn by Dogiel (1899). Neurons A and B are from the myenteric plexus of the guinea-pig small intestine, neuron C is from the myenteric plexus of human small intestine and neuron D is from the myenteric plexus of guinea-pig large intestine.
The morphologies and projections of Dogiel type II neurons have been thoroughly determined by intracellular dye filling, retrograde tracing, immunohistochemistry, and electron microscopy. Dye filling shows that the long processes of these neurons within the myenteric plexus run primarily in the circumferential direction (Bornstein et al. 1991b). The long processes issue large numbers of fine varicose branches in the same and adjacent ganglia (Fig. 2.5). Projections to adjacent ganglia were already described by Dogiel (1899) and Iwanowa (1958), who repeated Dogiel’s methylene blue staining technique, illustrated branching varicose processes similar to those later
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C B
Fig. 2.4 Neurons in the myenteric plexus of the rabbit intestine, drawn by La Villa (1898). Cell bodies corresponding to Dogiel’s descriptions are readily recognized, for example a type I neuron (neuron C) and a type II neuron (B).
revealed by dye filling of the neurons (Fig. 2.6). Electron microscopy demonstrates that the varicose branches in myenteric ganglia provide synaptic connections with other neurons, including other Dogiel type II neurons (Pompolo & Furness 1988). Dogiel (1899) found that some processes projected to the mucosa from the cell bodies of Dogiel type II neurons in submucosal ganglia. Because he worked with wholemounts of the separated layers, he could not examine projections to the mucosa from myenteric neurons, although it is now known that the myenteric Dogiel type II neurons also project to the
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Fig. 2.5 An IPAN (AH/Dogiel type II neuron) that had been injected with biocytin after being characterized electrophysiologically. A and B show micrographs of the region of the ganglion that contains the cell body at two different magnifications. C is a camera lucida drawing of the neuron and its processes within the ganglion containing its cell body; the varicosities on the fine processes that are a prominent feature (see micrograph in B) have not been drawn. Arrows point to expansion bulbs, where axonal processes were broken by removal of the circular muscle. Calibration: 40 μm, applies to all. Reproduced from Bornstein et al. (1991).
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Fig. 2.6 Drawing of the cell body and terminals of a Dogiel type II neuron in the myenteric
plexus of the cat small intestine, from a preparation stained with methylene blue, reproduced from Iwanowa (1958). The branching, varicose appearance of the terminals is similar to that seen with more modern techniques (e.g. Fig. 2.5).
mucosa in all species where this has been examined – guinea-pig (Furness et al. 1990b, Song et al. 1991, 1994, Neunlist & Schemann 1997, Lomax & Furness 2000, Vogalis et al. 2000), pig (Brehmer et al. 1999b, Hens et al. 2000), human (Hens et al. 2001), and mouse (Furness et al. 2004b). Indeed, it is probable that all myenteric Dogiel type II neurons project to the mucosa, although this has only been directly shown for the guinea-pig small intestine (Song et al. 1994). The processes that run to the mucosa provide collaterals that innervate neurons in the submucosal ganglia (Furness et al. 1990b). A feature of Dogiel type II neurons that is revealed by electron microscopy is their content of large numbers of mitochondria, and numerous lysosomes (Pompolo & Furness 1988). Because of their size and these ultrastructural features they can be distinguished from other neurons even without using immunocytochemical methods. Although the majority of terminals of myenteric Dogiel type II neurons are in ganglia close to the cell bodies, in a band that is circumferentially oriented and about 2 mm wide in the guinea-pig, some neurons have long axons that project anally for distances from 2 to 40 mm in most cases, but occasionally up to 100 mm (Brookes et al. 1995). Functional studies indicate that these long anally-directed processes form synapses in myenteric ganglia (Johnson et al. 1996). Some Dogiel type II neurons in the duodenum have projections to ganglia of the sphincter of Oddi (Kennedy & Mawe 1998). Dogiel type II neurons in submucosal ganglia have also been examined after dye filling, and, like their myenteric counterparts, they supply terminals that innervate neurons in adjacent ganglia (Lomax et al. 2001, Reed & Vanner 2001, Furness et al. 2003a). They also project to the mucosa (Dogiel 1899, Kirchgessner et al. 1992). The projections travel very little distance along the intestine; all submucosal Dogiel type II neurons that were labeled from the mucosa were within about 2 mm oral or anal to the labeling site (Song et al. 1992).
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A subclass of Dogiel type II neurons with short dendrite-like processes (dendritic type II neurons) has also been described (Stach 1989, Bornstein et al. 1991b, Brookes et al. 1995). However, there is no substantive functional data that defines the short processes as dendrites. No physiological data on the short processes has been obtained, and it is not known whether they receive synapses, although synapses are observed on the cell bodies and initial parts of axons (Pompolo & Furness 1988). Some of the dendrite-like processes give rise to varicose processes that appear to be axons that innervate other nerve cells (Nurgali et al. 2003b). Thus, whether the short processes are dendrites or initial segments of axons remains undetermined. The short processes, in fact all processes of Dogiel type II neurons, are not immunoreactive for the dendritic marker MAP2 (Murofushi et al. 1989, Scheuermann et al. 1991). There are only a few indications that Dogiel type II neurons with short dendrite-like processes may be different from other Dogiel type II neurons. In the guinea-pig small intestine, dendritic neurons have a different pattern of projections to other Dogiel type II neurons (Bornstein et al. 1991b, Brookes et al. 1995). The dendritic neurons supply terminal fields to fewer local ganglia, they rarely supply terminals to their own ganglion, and they have long anally projecting axons. Thus dendritic neurons may have a different role in the circuitry than do other Dogiel type II neurons, even though no electrophysiological differences have been detected. Between 80 and 90% of Dogiel type II neurons in guinea-pig myenteric ganglia are immunoreactive for the calcium-binding protein calbindin (Iyer et al. 1988, Song et al. 1991). However, dendritic neurons cannot be distinguished from adendritic neurons on the basis of immunoreactivity for calbindin in the small intestine or colon (Brookes et al. 1995, Nurgali et al. 2003b). Dogiel type III neurons Type III cells were described by Dogiel as having from 2 to 10 dendrites that became thinner and branched as they were followed from the cell body. The dendrites are depicted as relatively short, compared with the processes of type II cells, and they end within the ganglion of origin. The dendrites are generally smooth, but sometimes have varicosities. Some type III cells with thick dendrites were also seen by Dogiel (Fig. 2.7). The axon begins from a small conical protrusion of the cell body, or from a dendrite. The axons are initially smooth and were seen entering fiber bundles, but could not be followed to their terminations. Some short side branches sometimes arose from the initial segment of the axon. It is difficult to be sure which of the neurons revealed by current methods correspond to Dogiel type III neurons,
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Fig. 2.7 Examples of Dogiel type III neurons as defined and drawn by Dogiel (1899). Both
neurons are from the myenteric plexus of the guinea-pig large intestine. The upper neuron is similar to neurons that were later identified as filamentous interneurons in the small and large intestine of the guinea-pig.
but these are probably best matched by neurons with long branched processes (filamentous neurons) that have been described in the guinea-pig (Portbury et al. 1995b, Lomax et al. 1999). Most of the neurons illustrated by Dogiel, and both the examples of Dogiel type III neurons, are from the guinea-pig (Fig. 2.7). Dogiel type III neurons have also been identified in the pig small intestine, where many of them project anally within the myenteric plexus and another population projects from the intestine to prevertebral ganglia (Stach 1982a, Timmermans et al. 1993). Extension of Dogiel’s classifications: types IV, V, VI and VII Methods developed in the last 20 years, particularly methods of injection of individual neurons with marker dyes, and immunohistochemical methods, have greatly improved the ability to reveal neuron shapes. These methods have confirmed the existence of Dogiel type I and II neurons in many species of mammal, and have revealed type III in more limited examples. The classification of enteric neurons by shape has been further developed by several authors, most notably Stach and Brehmer (Stach 1989, Stach et al. 2000, Brehmer et al. 1999a, 2002, 2004a). These authors have identified neurons that are designated types IV to VII and mini-neurons (Fig. 2.8). With the exception of mini-neurons, I have used Stach and Brehmer’s terminology in this book. Mini-neurons (also called small neurons or simple neurons; Brehmer et al. 1999a) have features that liken them to Dogiel type I neurons (Furness
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Fig. 2.8 Type IV, V and VI neurons. The type IV neuron is from the submucosal plexus of the
guinea-pig ileum and is reproduced from Furness et al. (2003a). The type V and VI neurons are from the myenteric plexus of the pig intestine. Micrographs kindly supplied by Professor Werner Stach. Calibrations: 20μm.
et al. 2000), and I have referred to them in this book as small Dogiel type I or small simple neurons. Small neurons that do not fit readily with Dogiel’s classifications were noted by several other investigators (Stöhr 1952, Gunn 1968, Stach 1989). Type IV neurons have a single axon that projects vertically (that is, through the circular muscle towards the mucosa) and short tapering dendrites that branch, but less extensively than dendrites of type III neurons (Stach 1982b, 1989). They often have an eccentrically placed nucleus and dendrites that are grouped at one side or one pole of the cell body. They were first identified in the pig, where some at least are secretomotor neurons (Brehmer et al. 1999b). I had originally classified similar neurons in the guinea-pig small intestine as type III, but later studies placed them in the type IV category (Brehmer et al. 1999a, Furness et al. 2003a). In the guinea-pig, as in the pig, these are secretomotor neurons. The definition of type V neurons also comes from studies of pig intestine, where these neurons form characteristic clusters (Stach 1985). They are uniaxonal neurons with long branched dendrites and the nucleus is often eccentrically placed (Stach 1989). Type V neurons have also been described in human myenteric ganglia (Brehmer et al. 2004b), where they generally have a stem-like process from which the axon and branched dendrites both arise. Type VI neurons have a single axon and fine dendrites that arise from its proximal part and from the cell body (Stach 1989). In the pig, these include
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neurons that project from the submucosal ganglia to the myenteric plexus. Two rare neuron types, type VII and giant neurons, have been described in dog, pig, and human intestine (Stach et al. 2000). Cell physiological classifications of enteric neurons Neurons in the small and large intestines The classification of enteric neurons by their electrophysiological properties is useful in defined circumstances, but is limited by the differences that are encountered between species, and by the fact that the properties of neurons can be altered by the actions of neurotransmitters, hormones, and other agents, for example inflammatory mediators such as histamine and prostaglandins. The first intracellular recordings from enteric neurons, and hence the first descriptions of their passive and active electrical properties that provided a basis for classification, were reported in the 1970s (Nishi & North 1973a, Hirst et al. 1974, Wood & Mayer 1978). Nishi and North separated the neurons into three groups: type 1 neurons had relatively high input resistances and fired action potentials throughout a depolarizing pulse, type 2 neurons exhibited only one action potential in response to a 50 ms depolarizing pulse (that is, they showed rapid accommodation) and type 3 neurons were inexcitable. It is quite likely that the inexcitable neurons were neurons that were damaged by impalement, or were glial cells. With the better recording methods that are now available it is very rare to encounter inexcitable neurons. The terms S and AH (Fig. 2.9) to distinguish neurons in the small intestine of the guinea-pig
Fig. 2.9 Action potentials generated in an S neuron (A) and in AH neurons (B, C, and D) by intracellular current pulses. The action potential in the S neuron is followed by a fast afterhyperpolarization, whereas that in the AH neuron is followed by a fast after-hyperpolarization and, after a partial restitution of the membrane potential, by a slow after-hyperpolarizing potential (AHP). The slow AHP is better seen in C and D where a greater timescale is shown. The AHP in D lasts about 10 s. The recordings are from neurons in the guinea-pig small intestine. Reproduced from Hirst et al. (1974).
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were introduced in 1974 (Hirst et al. 1974). The classification remains useful, although the reasons for calling them S and AH are no longer well justified. Historically, S stood for synaptic, the S neurons exhibiting large-amplitude, fast synaptic potentials when their inputs were stimulated, and AH stood for after-hyperpolarizing, in recognition of the very prominent after-hyperpolarizing potentials that follow action potentials in these neurons. AH neurons in the guinea-pig ileum usually do not exhibit fast excitatory post-synaptic potentials (EPSPs) and, when they are recorded, the EPSPs have very small amplitudes (Hirst et al. 1974, Iyer et al. 1988, Bornstein et al. 1994, Brookes 2001). On the other hand, slow EPSPS do occur in AH neurons and can trigger action potentials (Wood & Mayer 1978). The axons of enteric neurons are fine and unmyelinated, and, as this would predict, have slow conduction speeds. Measurements from fiber bundles give speeds of 0.3–0.7 m/s (Kosterlitz & Lydon 1971, Yokoyama et al. 1977). Antidromic action potentials that were recorded in the cell bodies of Dogiel type II neurons at 34°C travelled at speeds of 0.23 m/s (Hendriks et al. 1990). S neurons S neurons exhibit brief action potentials that are followed by short duration after-hyperpolarizing potentials (AHPs), lasting 20–100 ms. These action potentials are blocked by tetrodotoxin. S neurons have a variety of shapes, which include Dogiel type I and filamentous morphologies; they never have Dogiel type II morphology in the guinea-pig intestine, where all S neurons have a single axon. They exhibit fast EPSPs, and compound fast EPSPs of sufficient amplitude to generate action potentials can be evoked in all S neurons. These EPSPs are referred to as “compound,” because they are the summed effects of transmission from several axons. Slow EPSPs are also recorded from S neurons (see Chapter 4). S neurons were classified as tonically firing on the basis of the continuity of action potential firing in response to 50 ms intracellular depolarizing pulses (Wood 1989). However, if pulses of 500 ms duration are used, there is a reduction or cessation of firing after about 200–250 ms (Kunze et al. 1997). The limitation of firing appears to be due to the presence of an outward rectifier current. Brookes et al. (2001) provide evidence that different types of neurons which fall into the S category have different properties. For example, ascending but not descending interneurons had a prominent A current. Specialization of a class of S neurons was also shown by Smith et al. (1999), who reported highly excitable S neurons, with high input resistances, which occur in the corners of myenteric ganglia. These appear to be longitudinal muscle motor neurons. However, the literature on electrophysiological subclassification of S neurons is incomplete.
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There are some S neurons that exhibit slow AHPs. These include filamentous interneurons in the small and large intestines (Song et al. 1997b, Clerc et al. 1998a, Lomax et al. 1999). Some neurons in the rectum that exhibit a prolonged AHP are uniaxonal and receive fast EPSPs (Tamura 1992). AH neurons The major channels and currents of these neurons have been extensively investigated (Fig. 2.10; Table 2.2). The action potentials of the AH neurons in the guinea-pig small intestine are large, 75–110 mV in amplitude, when measured with intracellular electrodes, of greater duration than those of S neurons, and have an inflection (hump) on the falling phase. They are normally followed by two separate phases of hyperpolarization, an early and a late AHP (Fig. 2.10). The slow AHP can last from about 2 to about 30 s
Fig. 2.10 The resting currents, action potential, and after-potentials that occur in the cell bodies
of AH neurons, derived from analysis of neuron properties in the guinea-pig small intestine. The potentials are shown schematically, not to scale. Upward arrows, outward currents; downward arrows, inward currents. Soma action potentials are caused by the opening of voltage-activated sodium conductances (gNaV) and a calcium conductance (gCaV) that is carried by N-type Ca2+ channels. The major component of the inward Na+ current is blocked by tetrodotoxin (TTX). There is also a TTX-resistant sodium conductance, gNaV (non-TTX), which may make a minor contribution. The calcium current outlasts the sodium current and is responsible for the hump on the falling phase of the action potential. The action potential is terminated by decline in the Na+ and Ca2+ conductances and activation of at least two K+ conductances, a BK current and a delayed rectifier current, that also contribute to an early after-hyperpolarizing potential (early AHP). This is followed by a late AHP that is carried by a calcium-dependent potassium conductance (gKCa), the ion channels for which are IK channels. In about 80% of AH neurons, the hyperpolarization of the late AHP triggers a depolarizing non-selective cation current, Ih, carried by HCN channels. The Ih current reduces the amplitude of the late AHP. An after-depolarizing potential (ADP), due to Ca2+ activation of a mixed cation conductance (gCAN), occurs between the early and late AHPs. The termination of the early AHP may be contributed to by the A current that is prominent in AH neurons. Adapted from Furness et al. (2004a).
ATP activated K
After-depolarizing current, IADP
Chloride
Delayed rectifier BK current Background K
A-type K+ current, IA
Inward rectifier
Hyperpolarization activated cation current, Ih
After-hyperpolarizing current, IAHP
Action potential current, ICaV
Niflumic acid block Niflumic acid block (high concentration) Tolbutamide block
Ca2+ activated cation channel (CAN) KATP
Sensitive to cytoplasmic ATP
4-AP block, TEA-resistant. Increased by 100 μM Zn2+ (effect competed by H+) TEA (10 mM) block TEA and charybdotoxin block One component H+ sensitive
Blocked by Cs and Ba
CaV2.2, N-type Ca channel HVA current (minor contribution from R-type Ca channel) KCa3.1, IK channel Ca2+ sensitive, voltage insensitive, apamin resistant HCN family, cationic Voltage sensitive, persistent, channel on and off τ between 50 and 300 ms Kir family Rapidly activated by hyperpolarization negative to EK KV4.x family Depolarization activated, fast inactivating but could be persistent around –55 mV ? Depolarization activated KV 1.x Ca2+ and voltage sensitive KCa1.1 Uncharacterized K2P family Voltage insensitive, may include several family members Cl– Ca2+ sensitive
Rugiero et al. 2003
Zholos et al. 2002
Reference
Liu et al. 1999
Bertrand & Galligan 1994; Starodub & Wood 2000a Vogalis et al. 2002b
Galligan et al. 1989b; Rugiero et al. 2002 Hirst et al. 1985a; Galligan et al. 1989b; Starodub & Wood 2000c Hirst et al. 1985a Kunze et al. 2000 Ngugen et al. 2005
ωGVIA, ωMVIIA and ωMVIIC Rugiero et al. 2002 but no ω−agatoxin GIVA block Charybdotoxin and Hirst et al. 1985b; Neylon clotrimazole sensitive et al. 2004 Cs and ZD7288 block Galligan et al. 1990; Xiao 2004
NaV family, unknown which Voltage sensitive, fast inactivating TTX blocks α-subunit NaV1.9 (NaN, SNS-2) Voltage sensitive, slow inactivating TTX resistant window current
Pharmacology
Action potential current, TTX sensitive INaV Subthreshold current, TTX-R INaV
Characteristics
Channel
Current
Table 2.2 Currents and channels of intrinsic primary afferent neurons. Channel identification in bold from IUPHAR Compendium of Voltage-Gated Ion Channels (ed. WA Catteral, KG Chandy, GA Gutman, International Union of Pharmacology, 2002)
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(Hirst et al. 1974, 1985b). All Dogiel type II neurons in the guinea-pig ileum are AH neurons. However, most Dogiel type II neurons in the pig intestine lack the late AHP, although they do have a hump on the action potential (Cornelissen et al. 2000). Whether this lack of a late AHP is a consequence of the state of excitability that the neurons happened to be in is not known; as I discuss below, the AHP can be inhibited by many endogenous compounds to which AH neurons are exposed. Action potentials in AH neurons. Two inward currents underlie the action potential in the soma of AH/Dogiel type II neurons, a tetrodotoxin (TTX)sensitive Na+ current (INaV) and a TTX-insensitive Ca2+ current (ICaV) (North 1973, Hirst et al. 1985a, Rugiero et al. 2003). A TTX-resistant Na+ current (TTX-R INaV) also occurs in these neurons (Rugiero et al. 2003), but there is no evidence that this contributes in any significant way to the action potential in physiological conditions. The properties of TTX-R INaV are consistent with it being due to the expression of NaV1.9, and transcripts corresponding to this channel, but not to NaV1.8, were obtained by single cell reverse transcriptase polymerase chain reaction (RT-PCR) of AH neurons (Rugiero et al. 2003). In the presence of TTX, the Ca2+ current is still sufficient for action potential generation in the soma. The ICaV is responsible for the inflection (hump) on the repolarizing phase of the action potential. The hump on the action potential persists in the presence of L-type calcium channel blockers (North & Tokimasa 1987, Kunze et al. 1994). The identity of the high voltage activated (HVA) Ca2+ channels has been investigated in experiments in which the contribution of Ca2+ to the action potential was enhanced by adding tetraethylammonium (TEA). The Ca2+ plateau that occurs in the presence of this K+ channel blocker and nicardipine was almost blocked by ω-conotoxin GVIA or ω-conotoxin MVIIA, both blockers of N-type channels, but not by ω-agatoxin GIVA which is a blocker of P/Q-type channels (Rugiero et al. 2002). There may be a small contribution of R channels to the ICaV (Rugiero et al. 2002). Consistent with N-type channels being the predominant type of HVA channel of AH neurons, immunoreactivity for the α1B (N-type) calcium channel subunit is localized to Dogiel type II neurons, whereas only weak α1B (P/Q-type channel) immunoreactivity was found (Kirchgessner & Liu 1999b). The action potentials in the long processes of the AH neurons (process action potentials) differ from the action potentials in the cell bodies. Conduction of process action potentials is blocked by TTX, indicating that a TTXsensitive Na+ current is the major action potential current in the processes (North & Nishi 1976). If there is an active Ca2+ current in the processes, it is not sufficient to sustain a regenerative action potential. Moreover, the process action potential is not followed by a substantial slow AHP, because, unlike in the soma region, the frequency of generation of process action potentials
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Fig. 2.11 Blockade of the soma action potentials in an AH neuron by the slow after-hyperpolarizing potential (AHP). An axonal process of the cell was stimulated by brief electrical pulses at 1 Hz. The first three pulses gave rise to action potentials in the cell body. Thereafter, the AHP blocked the generation of a high proportion of action potentials and, in their place, proximal process potentials, the small upward deflections, were seen in the soma. Even though the process was stimulated at 1 Hz, and process action potentials followed the stimulation faithfully, the soma fired action potentials at an average frequency of about 0.2 Hz. Reproduced from North and Nishi (1974).
is not limited by the occurrence of the AHP (Fig. 2.11), and the excitability of the processes is unaltered when a slow AHP is observed in the soma (Hirst et al. 1974, North & Nishi 1974). The channels that carry the AHP current, IK channels, occur in the soma and only the initial parts of the AH neuron processes, which is consistent with the AHP being absent from the processes, except close to the soma (Furness et al. 2003c). Properties of the TTX-sensitive Na+ current have been analyzed in cultured myenteric AH neurons of adult guinea-pigs (Zholos et al. 2002). The current was activated between –50 and –40 mV and peaked at –10 mV. Inactivation kinetics were fast and the steady state half-inactivation potential was –56 mV. The time constant of inactivation decreased as the holding potential depolarized, with a maximum of 161 ms at –70 mV and a minimum of 2.3 ms at –30 mV. Recovery from inactivation was also rapid with time constants between 7 and 44 ms for holding potentials of –100 and –60 mV respectively. The early AHP in AH neurons. The early AHP is continuous with the falling phase of the action potential. It lasts 20–100 ms and is similar to the undershoot that is seen in most neurons, including enteric S neurons, neurons in the central nervous system and neurons in autonomic ganglia. The currents of the early AHP have been investigated in some detail, and include contributions from a TEA-sensitive delayed rectifier (Hirst et al. 1985a, Zholos et al. 1999) and large conductance Ca2+- and voltage-sensitive K+ channels (BK channels). If the resting potential is more negative than about –60 mV, the contribution of a 4-aminopyridine-sensitive A current is detected (Hirst et al. 1985a). The after-depolarizing potential. Following the early AHP there is a depolarization that is followed by the slow AHP (Figs 2.9, 2.10). This gives the
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misleading impression that the onset of the slow AHP current is delayed. However, when the underlying channel activity is recorded it is clear that the current onset is rapid, within a few milliseconds (Vogalis et al. 2002a). The transient depolarization (ADP) that occurs between the early and late AHPs is due to a Ca2+-activated depolarizing cation current (Vogalis et al. 2002b). The physiological significance of the ADP is not known. The current is blocked by niflumic acid, but at higher concentrations than are needed to block Ca2+activated Cl– currents. The late AHP and its influence. The excitability of AH neurons is critically dependent on the degree of activity of the late AHP current, both at rest and following an AHP. When IAHP is activated following the action potential, the resultant hyperpolarization can be sufficient to reduce subsequent action potentials that arrive at the cell soma to non-regenerative proximal process potentials (Fig. 2.11). A consequence is that action potentials that would normally cross the cell and provide inputs to interneurons and motor neurons (Figs 2.19, 3.3, 3.5) are prevented from doing so. Thus the cell body acts as a gate in the circuit (Wood & Mayer 1979a, Furness & Costa 1987). The AHP and its effect in gating the outputs of the IPANs is important in determining the properties of peristaltic reflexes (Chapter 5). Ca2+ entering during the action potential is necessary for generation of the late AHP. The Ca2+ dependence of the K+ current underlying the late AHP was first established by ion substitution (Hirst & Spence 1973, Hirst et al. 1974). While addition of TTX has no effect, the absence of external Ca2+, or addition of calcium channel blockers such as Co2+, Mn2+, or Mg2+, suppress the IAHP following an action potential (Morita et al. 1982a, Hirst et al. 1985b, North & Tokimasa 1987). As mentioned above, enhancing the Ca2+ contribution to the action potential by adding TEA allows IAHP to be amplified without altering its onset kinetics, and this enhances the AHP (Hirst et al. 1985a,b, North & Tokimasa 1987). IAHP increases progressively when the cell is dialyzed with caffeine included in a patch pipette (Rugiero et al. 2002), indicating that release of Ca2+ from intracellular stores can contribute to the opening of K+ channels and the generation of the AHP. Moreover, the AHP is substantially reduced by ryanodine and when Ca2+ stores are depleted by long-term application of caffeine (Hillsley et al. 2000, Vogalis et al. 2001). Therefore, the activation of the channel depends on Ca2+-induced Ca2+ release from intracellular stores via a ryanodine-sensitive receptor (Fig. 2.12). The repolarization of the AHP is dependent on removal of Ca2+ by mitochondria, because block of mitochondrial Ca2+ uptake or of mitochondrial respiratory enzymes greatly prolongs the AHP (Vanden Berghe et al. 2002). IAHP is very weakly voltage sensitive and is not affected by apamin. These properties indicate that the channels responsible are not conventional BK
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Fig. 2.12 Pathways for control of IK channels of intrinsic primary afferent neurons. The opening probabilities of channels are increased by intracellular Ca2+ and reduced by channel phosphorylation, through the actions of both PKA and PKC. PKC is linked to the so-far unidentified transmitter of sustained slow excitation (the SSPE) and to the primary transmitters of slow EPSPs, the major one being a tachykinin acting through NK3 receptors. The action potential admits Ca2+ through the soma membrane. This Ca2+ triggers Ca2+-induced Ca2+ release from intracellular stores, which opens IK channels. Ca2+ also increases the activity of protein phosphatase 2B (calcineurin), which dephosphorylates the channels and permits their activation. The IK current generates the late after-hyperpolarizing potential (AHP). Removal of Ca2+ by mitochondria is important for terminating the AHP.
(highly voltage-sensitive) or SK (apamin-sensitive) channels. Recent evidence shows that they are IK channels (Vogalis et al. 2002a, Neylon et al. 2004), which were formerly not thought to be expressed in neurons (Sah & Faber 2002). These channels have been localized to Dogiel type II neurons, and the channel protein and mRNA has been detected in extracts of external muscle including myenteric ganglia from human, rat, and mouse (Furness et al. 2003b, 2004b, Neylon et al. 2004). The IAHP current contributes to resting conductance. Charybdotoxin (20 nM), which blocks the IK channels responsible for IAHP, depolarized AH neurons from –57 to –49 mV and increased input resistance from 190 to 260 MΩ in experiments using intracellular recordings (Kunze et al. 1994). Consistent with this, AH neurons depolarize by 4–15 mV in Ca2+-free solution, and their input resistance increases by approximately 15% (North & Tokimasa 1987). Block of IAHP with clotrimazole causes a 10 mV depolarization (Nguyen et al. 2005). IK channels have consensus sites for protein kinase A (PKA) and protein kinase C (PKC) binding, and phosphorylation by either of these kinases
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causes a reduction in their opening probability (Wulf & Schwab 2002, Del Carlo et al. 2003, Vogalis et al. 2003). AH neurons of the guinea-pig small intestine contain PKA and several isoforms of PKC (Poole et al. 2003). Consistent with this, the PKC stimulant phorbol dibutyrate (PDBu; 1 nM–1 μM) causes excitability increases, membrane depolarization, and increased input resistance of AH neurons in a concentration-dependent manner (Kawai et al. 2003). The catalytic subunit of PKA, added to inside-out patches from AH neurons, reduces the opening probability of the IK channels, and in whole cell recording, its application quickly suppresses the IAHP (Vogalis et al. 2003). The IK channels are probably controlled by a dynamic equilibrium between phosphorylation and dephosphorylation (Fig. 2.12). The major dephosphorylating enzyme for IK channels in AH neurons is calcineurin (protein phosphatase 2B) and block of calcineurin reduces IAHP substantially in a period of 10–20 min (Vogalis et al. 2004), indicating that there is ongoing phosphorylation of the channel at rest. Hyperpolarization-activated cation current of AH neurons. A hyperpolarization-activated non-specific cation current (Ih) occurs in AH neurons (Galligan et al. 1990, Rugiero et al. 2002). This current is active at rest even in AH neurons in a low state of excitability (Rugiero et al. 2002). It is persistent in its activation range, and thus contributes to membrane conductance at potentials more negative than about –40 mV. It has slow activation and inactivation kinetics (Rugiero et al. 2002, Nguyen et al. 2005). The regulation of this current has been little studied in AH neurons, but in other cell types, such as cardiac muscle cells, its regulation through cyclic AMP plays an important role in determining cell excitability. In AH neurons in guinea-pig, mouse, and rat, the prominent Ih channel type is HCN2 (Xiao et al. 2004). Ih is increased by IAHP-induced hyperpolarization, and is involved in the restoration of the membrane potential as the slow AHP declines. Ih activation can generate a sustained depolarization (Galligan et al. 1990, Rugiero et al. 2002). Other currents in AH neurons. AH neurons also exhibit an A-type K+ current (IA) (Hirst et al. 1985b, Galligan et al. 1989b, Nguyen et al. 2005). This current is inactivated at resting potential in most neurons, but in AH neurons IA may have a low level of activation at rest (Starodub & Wood 2000b). IA is deinactivated by hyperpolarization below about –60 mV (Hirst et al. 1985a). It limits firing rates by slowing the return of the membrane potential after hyperpolarization, for example, following the hyperpolarization caused by the fast AHP (Fig. 2.10). Chloride currents occur in AH neurons, but there is no evidence to suggest that they make a significant contribution to resting conductance (Bertrand & Galligan 1994, Starodub & Wood 2000a). Activation of GABAA receptors, which are ligand-gated chloride channels, depolarizes
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AH neurons (Cherubini & North 1984), confirming that the chloride equilibrium potential in these neurons is positive to resting membrane potential. KATP channels also occur in AH neurons (Liu et al. 1999). These channels are opened by lowering intracellular ATP and were deduced to be present by the hyperpolarization of the neurons caused by lowering bath glucose, and the block of this effect by tolbutamide (Liu et al. 1999). Immunoreactivity for the KATP channel complex was also detected in AH/Dogiel type II neurons. The cytoplasm of Dogiel type II neurons has about four times the density of occurrence of mitochondria compared to other myenteric neurons (Pompolo & Furness 1988), which suggests that they are highly metabolically active, and that they may require the protective role of the KATP channels, for example if there is a lowering of O2 tension in the intestine. Influence of recording conditions. The electrophysiological properties of AH neurons are influenced by the recording conditions. When background synaptic transmission is suppressed, they exhibit the late AHP that identifies them as AH neurons in the terminology of Hirst (Hirst et al. 1974), but when AH neurons are acted upon by neurotransmitters or hormones, the late AHP can be suppressed or obliterated (Wood & Mayer 1979a, Morita & North 1985, Clerc et al. 1999). AH neurons in a state of low excitability do not fire more than 1–2 action potentials in response to a 500 ms intracellular depolarizing pulse, they have a resting membrane potential of around –60 to –65 mV, and they can be defined by the presence of the late AHP. The resting conductance is low, with input resistance of about 200 MΩ measured by intracellular electrodes and about 500 MΩ measured with whole cell patch recording. Background K+ and to a smaller extent Na+ and Cl¯ currents contribute to the resting conductance. Some contribution to the background conductance is made by the hyperpolarization-activated cation current, Ih. Whole cell patch recording estimates the contribution of Ih at about 20% of the cell conductance (Rugiero et al. 2002). The IAHP current contributes about 30% of the conductance of AH neurons that are in a low state of excitability (Nguyen et al. 2005). Neurons in other regions: stomach, esophagus, gall bladder, and pancreas In the gastric corpus, almost all myenteric neurons have S characteristics, and do not have a prolonged hyperpolarization after the action potential (Schemann & Wood 1989a,b). Rare neurons with AH characteristics are found in the antrum (Tack & Wood 1992). This is consistent with the rarity of Dogiel type II neurons in the stomach. There is one report of electrophysiological investigation of esophageal neurons (De Laet et al. 2002). These neurons were in myenteric ganglia of
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the striated muscle part of the rat esophagus. All the neurons had an action potential with a hump on the falling phase, which was mediated by a Ca2+ current through N channels. However, all except one neuron were uni-axonal with short dendrites. Thus the relation between electrophysiology and morphology for esophageal neurons appears to be different to neurons in the intestine. Neurons on the gall bladder appear to be all of the S-type and most of the neurons have a simple morphology, with very short or no apparent dendrites and a single axon, although a few had long dendrite-like processes (Mawe 1990, Bauer et al. 1991). No neurons with AH electrophysiological characteristics or Dogiel type II morphology were encountered. This is consistent with gall bladder neurons being under vagal control, and with an apparent absence of intrinsic gall bladder neuronal reflexes (Mawe 1998). The electrophysiological properties and shapes of neurons with cell bodies in the pancreas have been investigated in cat and guinea-pig (King et al. 1989, Sha et al. 1996, Liu & Kirchgessner 1997). In both species, the neurons have a single axon and short filamentous dendrites. They have S-type electrophysiological properties. Fast EPSPs that are recorded from the neurons are blocked by hexamethonium and action potentials are blocked by tetrodotoxin. There is no evidence for AH/Dogiel type II neurons in pancreatic ganglia. Functionally defined enteric neurons Enteric neurons have been placed in physiologically defined groups by combining data from studies of the properties of enteric reflexes, the morphological identification and projections to targets of those neurons that are present, and correlation of neurochemical (primarily immunohistochemical) and pharmacological properties of enteric neurotransmitters and their receptors (Table 2.1). The model species has been the guinea-pig. From these data has come a complete definition of the types of neurons in the small intestine, and thorough, but not yet quite so complete, definitions of neuron types in the colon (Lomax & Furness 2000) and in the stomach (Schemann et al. 1995, Pfannkuche et al. 1998, Michel et al. 2000) (Fig. 2.13). In many cases the corresponding neurons have been identified in other species (see above). Motor neurons Muscle motor neurons: smooth muscle esophagus, stomach, small and large intestine, and gall bladder Excitatory and inhibitory neurons innervate the longitudinal and circular smooth muscle and the muscularis mucosae throughout the digestive tract.
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Fig. 2.13 Types of neurons, as defined by their functions, cell body morphologies, chemistries, and projections, in the colon of the guinea-pig. There are many similarities with the ileum (Fig. 2.1). Similar motor neurons and intrinsic primary afferent neurons (neuron types 2–7 and 11–14) are found in each region (see Table 2.1 for definitions of the neurons). The major differences are in the types of interneurons that are present in the two regions. LM, longitudinal muscle; MP, myenteric plexus; CM, circular muscle; SM, submucosal plexus; MM, muscularis mucosae; Muc, mucosa; PVG, prevertebral ganglia.
These are uni-axonal neurons with S-type electrophysiology. The primary transmitters of the excitatory neurons are acetylcholine and tachykinins. The inhibitory neurons have multiple transmitters, including nitric oxide (NO), VIP, and ATP, which are released by each neuron (Chapter 4). The relative roles of the several inhibitory transmitters vary between regions and species (Furness et al. 1995b). Neurons innervating the circular muscle (circular muscle motor neurons). The majority of neurons that innervate the circular muscle have their cell bodies in the myenteric ganglia. In fact, in the guinea-pig they are all in myenteric ganglia, as shown by electron microscopic analysis following ablation of the myenteric plexus (Wilson et al. 1987). In other species, including rat (Ekblad et al. 1987, 1988), dog (Sanders & Smith 1986, Furness et al. 1990a), pig (Timmermans et al. 2001), and probably human, a component of circular muscle innervation comes from submucosal ganglia. In large mammals the submucosal supply to the muscle comes primarily from the outer submucosal plexus.
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Quantitative ultrastructural studies, which are supported by numerous physiological and immunohistochemical investigations, indicate that there are only two types of neurons that innervate the circular muscle (Llewellyn Smith et al. 1988). These are the inhibitory motor neurons, which are best revealed by their immunoreactivity for the synthesizing enzyme for NO, nitric oxide synthase (NOS) or by immunoreactivity for VIP, and excitatory motor neurons revealed by immunoreactivity for tachykinins or the vesicular acetylcholine transporter. Enteric inhibitory and excitatory motor neurons supply both the sphincter and non-sphincter circular muscle of the gastrointestinal tract. NOS can be readily revealed histochemically, and shows the shapes of the cell bodies of the inhibitory motor neurons very clearly. These are Dogiel type I neurons in most species (Ward et al. 1992, Timmermans et al. 1993, Furness et al. 1994, Costa et al. 1996, Nurgali et al. 2004), including human (Porter et al. 1997). However, this does not apply in the pig, where NOcontaining neurons have a variety of shapes, and anally projecting neurons have type III and type VI shapes (Timmermans et al. 1994, Brehmer & Stach 1997). Dogiel type I morphology is also revealed by retrograde labeling of motor neurons from the muscle in both guinea-pig and human (Brookes et al. 1991b, Wattchow et al. 1995, 1997), and by intracellular dye injection combined with immunohistochemical identification of the neurons (Bornstein et al. 1984a, Katayama et al. 1986). The distances that circular muscle motor neurons project along the intestine have been mapped by placing small crystals of the tracer dye DiI on the muscle and waiting for it to be transported back to the cell bodies of the neurons (Brookes et al. 1991b, Wattchow et al. 1995). In the small intestine of the guinea-pig, retrogradely labeled neurons were distinguished as excitatory motor neurons (immunoreactive for choline acetyltransferase) and inhibitory motor neurons (immunoreactive for VIP) (Brookes et al. 1991b). The experiments showed that the majority of excitatory motor neurons innervate the muscle between 5 mm oral and 2 mm anal to the cell body in the myenteric plexus, and the majority of inhibitory motor neurons innervate the circular muscle 0–6 mm anal to the cell body. Lesion studies provide essentially the same data, excitatory motor neurons innervating the muscle adjacent and just oral to the nerve cell bodies and inhibitory motor neurons innervating the muscle just anal (Costa et al. 1981, Costa & Furness 1983, Daniel et al. 1987, Ekblad et al. 1987, 1988, Williamson et al. 1996). The projections of inhibitory neurons to the muscle have also been mapped using electophysiological techniques (Bornstein et al. 1986). These studies showed that the inhibitory motor neurons project anally and give rise to terminals that run circumferentially. The distances that neurons project was estimated by stimulating the descending nerve pathways in the presence of hexamethonium. This indicated that the majority of neurons projected up to about 8 mm anally,
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but some projected as far as 30 mm anally. Electrophysiological analysis of the projections of excitatory neurons show that the majority project for short distances, less than 3 mm, in an oral direction, with a few projecting as far as about 10 mm (Smith et al. 1988). Similar data come from experiments that examined projections in the mouse colon (Okasora et al. 1986). Excitatory neurons projected about 2 mm in the oral direction, and inhibitory neurons projected about 10 mm anally. Circular muscle motor neurons have been filled with dye and their processes traced into the muscle (Nurgali et al. 2004) (Fig. 2.14). The pattern of branching in the muscle was similar to that deduced when populations of stained neurons were examined (Cajal 1895, Furness & Bornstein 1991). The axon from each neuron branches extensively in the circular muscle and most neurons innervate a band of circular muscle that is 0.5–2 mm wide and runs around the intestinal circumference. Physiological studies and counts of neuron numbers indicate that the neurons overlap extensively in their fields of supply, and that several hundred neurons supply motor units that are about 2–3 mm wide (Chapter 3). Neurons innervating the longitudinal muscle. The cell bodies of motor neurons that supply the longitudinal muscle are in the myenteric plexus of small animals. In the pig, and probably in other large mammals, the majority of the cell bodies are in the myenteric plexus, but some longitudinal muscle motor neurons have cell bodies in the outer submucosal plexus. Processes have been directly traced from filled neurons in the guinea-pig, where each neuron innervates a small patch of muscle about 2 mm long and 1 mm wide (Furness et al. 2000) (Fig. 2.15). The longitudinal muscle motor neurons are surprisingly numerous. In the guinea-pig small intestine they are about 25% of nerve cells in the myenteric ganglia (Table 2.1). As has already been reviewed in Chapter 1, in small mammals the branching axons from these neurons form a meshwork at the inner face of the longitudinal muscle, whereas in larger mammals they enter the muscle coat and their branches run parallel to the muscle fibers, similarly to the branches of circular muscle motor neurons. Motor neurons of the muscularis mucosae. Similarly to other smooth muscle of the wall of the gastrointestinal tract, the muscularis mucosae is innervated by excitatory and inhibitory motor neurons (Onori et al. 1971, Gallacher et al. 1973, Angel et al. 1984). The cell bodies of neurons innervating the muscularis mucosae of the cat colon, which was investigated by Onori et al. (1971), were probably in the submucosa, because the muscle attached to the mucosa was contracted by DMPP, a stimulant of nicotinic receptors on the cell bodies of enteric neurons. In the small intestine and colon of the dog, removal of myenteric ganglia, allowing time for axon degeneration, did not
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Fig. 2.14 Representations of circular muscle motor neurons published between 1895 and 2004. A: Drawing of motor neurons made by Cajal (1895) from his examination of preparations in which all neurons were stained (similar to Fig. 1.8). B: Representation made in 1991 from studies of preparations in which most neurons were stained. Reproduced from Furness et al. (1991). C: The appearance of a single circular muscle motor neuron that was filled with a marker dye via an intracellular microelectrode. Reproduced from Nurgali et al. (2004).
change the innervation of the muscularis mucosae, which also suggests that the innervation derives from nerve cells in submucosal ganglia (Furness et al. 1990a). In the esophagus and stomach, where there are few or no submucosal nerve cells, the innervation must arise from nerve cells in myenteric ganglia, at least in small mammals. In larger mammals, including human, submucosal ganglia in the stomach may be a source of innervation of the muscularis mucosae (for example, Colpaert et al. 2002), but this has not yet been resolved.
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Fig. 2.15 The processes arising from the single axon of a motor neuron innervating the longitudinal muscle of the guinea-pig ileum (a tertiary plexus neuron). The neuron had been filled with a marker dye through an intracellular electrode placed in the cell soma. This image was created from a camera lucida drawing. The ganglia and connecting strands of the myenteric plexus are shown in gray. The axonal branches do not follow the main strands of the myenteric plexus; they run in the tertiary plexus (see Chapter 1).
Muscle motor neurons that supply the striated muscle of the esophagus Vagal motor neurons, with their cell bodies in the medulla oblongata of the brainstem, are the principal neurons through which the movements of the striated muscle portion of the esophagus are controlled (Chapter 5). The endings of these neurons form conventional motor endplates on the striated muscle cells (Samarasinghe 1972, Rodrigo et al. 1985). However, about a third of the endplates have an additional innervation from myenteric neurons, through which vagal excitation is modulated (Chapter 5). The distal, smooth muscle part of the esophagus and the lower esophageal sphincter are innervated by enteric neurons. Secretomotor and secretomotor/vasodilator neurons controlling fluid exchange Exocrine fluid secretion, such as that from the salivary glands, sweat glands, and pancreas, relies on supply of water and electrolytes from the blood. Be-
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cause of this, exocrine secretion is coupled to vasodilation. Coupling of fluid secretion and vasodilation also occurs in the intestine (Chapter 6), where secretion and vasodilation are controlled together. Transport of water and electrolytes across the mucosa, independent of blood flow, can be studied in isolated sheets of mucosa, or of mucosa plus underlying tissue, using the Ussing chamber. Hubel (1978) seems to have been the first to stimulate nerves electrically in such preparations. He found that activation of mucosal nerves generated an ionic current across the mucosa of the rabbit small intestine. This phenomenon has been further investigated in the small intestine of the rabbit (Hubel & Callanan 1980), human (Hubel & Shirazi 1982), and guinea-pig (Cooke et al. 1983a,b, Keast et al. 1985a,c). Analysis of the currents has shown them to be due to the active secretion of chloride ion, which is accompanied by sodium and water secretion. These secretory responses are blocked by tetrodotoxin. The final motor neurons that regulate these functions have been fully identified only for the small intestine of the guinea-pig, where there are three types of enteric secretomotor and secretomotor/vasodilator neurons (Furness et al. 2003a). These are: VIP-containing, non-cholinergic, secretomotor/vasodilator neurons that innervate both the mucosal epithelium and small arterioles in the gut wall; calretinin-containing, cholinergic secretomotor/vasodilator neurons; and NPY-containing cholinergic secretomotor neurons that do not also innervate arterioles (Fig. 2.16). The motor neurons that innervate the mucosa have been identified through a combination of immunohistochemical studies, investigations of their terminals in the mucosa after lesioning nerve pathways from which they could originate, retrograde tracing studies and pharmacological analysis of the control of secretion and vasodilation. VIP is a potent stimulant of secretion in the intestine (Barbezat & Grossman 1971, Schwartz et al. 1974), and nerve fibers containing VIP provide a rich innervation of the mucosa throughout the small and large intestines and in the gall bladder of the many mammalian species that have been examined (Furness et al. 1988). The VIP innervation of the mucosa is not altered following sympathetic and vagal denervation (Larsson et al. 1976), or following the removal of the myenteric plexus from a segment of intestine, and allowing time for the axons that are thus disconnected from their cell bodies to degenerate (Costa & Furness 1983, Keast et al. 1984). These experiments indicate that the VIP fibers in the mucosa arise from cell bodies in the submucosal plexus, and that VIP immunoreactivity is an effective marker of secretomotor neurons. Although the VIP neurons are not immunoreactive for the synthesizing enzyme for acetylcholine, choline acetyltransferase (ChAT) (Furness et al. 1984), there are two components of transmission to the mucosa, a cholinergic component and a non-cholinergic component (Cooke & Reddix 1994). Thus there must be separate cholinergic secretomotor neurons. In the guinea-pig, NPY and calretinin neurons, which are both ChAT-immunoreactive, provide the
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cholinergic innervation (Furness et al. 1985, Brookes et al. 1991a). In rat, the chemical coding of the neurons is different, with NPY immunoreactivity occurring in the VIP neurons (Ekblad et al. 1984). Both VIP and calretinin axons, arising from submucosal nerve cells, innervate arterioles in the gut wall, indicating that the VIP (non-cholinergic) and calretinin (cholinergic) neurons could have dual secretomotor and vasomotor functions (Costa & Furness 1983, Brookes et al. 1991a, Song et al. 1997a, Li et al. 1998). Experiments in which single neurons or the surfaces of submucosal ganglia were stimulated and the resulting changes in the diameter of submucosal blood vessels measured provide functional evidence for the presence of both cholinergic and non-cholinergic vasodilator neurons (Neild et al. 1990, Vanner & Surprenant 1991, Kotecha & Neild 1995). Direct evidence that the same neurons supply the mucosa and blood vessels was first obtained in experiments in which galanin and VIP were co-localized, and the projections of the neurons were determined in nerve lesion studies (Furness et al. 1987b). This was confirmed in elegant experiments in which the same neurons were simultaneously labeled by retrograde transport of different dyes applied to the mucosa and to submucosal arterioles (Jiang et al. 1993). Retrograde labeling by dye applied to the mucosa indicates that the secretomotor and secretomotor/vasodilator neurons project for only short distances along the small intestine, in most cases less than 1 mm in guinea-pig (Song et al. 1992) and 3–4 mm in human (Porter et al. 1999). Similar data has been obtained by tracing the axons from cell bodies (Furness et al. 1985, 2003a, Evans et al. 1994). The neurons branch within the submucosa and each provides innervation to an estimated eight villi (Song et al. 1992). Along with information on cell morphology (Furness et al. 2003a), the data on the branching patterns and co-localization of marker substances permit a reconstruction of the motor neurons to the mucosa (Fig. 2.16). In the colon, but not in the ileum, the mucosa receives a polarized innervation by descending non-cholinergic and ascending cholinergic pathways of the submucosal plexus (Chapter 3). Gastric vasodilator neurons Gastric acid secretion and blood flow are enhanced when the vagus nerve is stimulated and these effects are reduced by muscarinic antagonists. In most experiments, it is not possible to determine whether vasodilation is due to a direct vascular action of cholinergic neurons in addition to a functional hyperemia consequent on the increased secretion (Jodal & Lundgren 1989). However, Thiefin and colleagues showed that centrally administered thyrotropin-releasing hormone (TRH) stimulated a vagal pathway in the rat that caused gastric vasodilation after acid secretion was blocked by omeprazole (Thiefin et al. 1989). The blood flow increase in the absence of secretory
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Fig. 2.16 Secretomotor neurons of the small intestine. A: Non-cholinergic secretomotor/ vasodilator neuron. These secretomotor neurons project to multiple villi and also innervate the mucosal glands (crypts of Lieberkuhn) and submucosal arterioles. In all species these neurons contain VIP. B: Cholinergic secretomotor/vasodilator neuron. These neurons project selectively to the glands and bases of the villi in the small intestine, in addition to innervating arterioles. C: Cholinergic secretomotor neuron. These neurons innervate the glands and the villi, but not blood vessels.
change was antagonized by atropine. There is also evidence for non-cholinergic gastric vasodilator neurons that use VIP as a transmitter (Ito et al. 1988), but whether these are vasodilator alone or, more likely, secretomotor/vasodilator neurons has not been determined. Innervation of gastrointestinal veins and lymphatic vessels The veins within the walls of the stomach and intestine are very sparsely if at all innervated, but there is a sympathetic innervation of the mesenteric veins, which is quite dense for the larger veins, including the hepatic portal vein (Furness 1971). The sympathetic nerves constrict the large mesenteric veins and are involved in controlling the volume of blood held by the veins, which can be diverted to other regions of the body (Folkow et al. 1964). The larger extramural veins have a sparse innervation by tachykinin-immunoreactive
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nerve fibers, but the veins within the gut wall are not innervated by these fibers (Furness et al. 1982). It is presumed that the tachykinin fibers are afferents. Mesenteric veins also receive a sparse innervation by VIP-immunoreactive fibers (Della et al. 1983). Lymphatic vessels in the mesentery and in the wall of the gastrointestinal tract are sparsely innervated by sympathetic, noradrenergic nerve fibers, by sensory nerve fibers with tachykinin immunoreactivity, and by VIP-immunoreactive nerve fibers (Furness 1971, Guarna et al. 1991, Shimoda et al. 1998). Gastric secretomotor neurons that stimulate acid output Some secretomotor neurons govern gastric acid secretion. These neurons are cholinergic and act on the parietal cells through muscarinic receptors. Projection studies indicate that the secretomotor neurons have cell bodies in the myenteric plexus close to the regions of mucosa that they innervate (Pfannkuche et al. 1998). For a complete description of the roles of these neurons and their relation to hormonal control of gastric acid secretion, see Chapter 6. Motor neurons to enteric endocrine cells A variety of endocrine cells reside in the mucosa of the gastrointestinal tract, and because the mucosa is densely innervated, these cells have nerve fibers in close proximity, but it is not clear in all cases whether the endocrine cells are functionally innervated. The best documented motor neurons innervating enteric endocrine cells are those controlling release of gastrin, which is under the influence of vagal and of intrinsic gastric pathways (see Chapter 6). The final motor neurons in both paths are in the gastric myenteric plexus (Ekblad et al. 1991, Furness et al. 1991). Transmission from the final secretomotor neurons is mediated at least in part by gastrin-releasing peptide (Makhlouf et al. 1989). Hormone release from other entero-endocrine cells is also likely to be under neural control. Peptide YY is released from the distal small intestine by electrical vagal stimulation, and there is also evidence of vagal reflex control of its release (Onaga et al. 2002). The release is attenuated by atropine. The basal release of motilin is reduced by atropine and by tetrodotoxin, and stimulated by muscarinic agonists, suggesting that motilin cells receive an excitatory cholinergic input (Poitras et al. 1997). There is also evidence for nerve-mediated release of 5-HT from enteric endocrine cells. Innervation of lymphoid tissue (Peyer’s patches), lymphocytes, and mast cells Lymphoid aggregations of the gastrointestinal tract, Peyer’s patches, have surrounding nerve fibers, but these do not enter the lymphoid tissue proper, or only
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very rarely (Krammer & Kuhnel 1993, Kulkarni-Narla et al. 1999). This suggests that there is no direct neural influence on lymphocytes within intestinal lymph nodes. On the other hand, receptors for transmitters of enteric neurons occur on lymphocytes that are scattered in the connective tissue (lamina propria) of the mucosa (Heel et al. 1997, Renzi et al. 2000), and there are close approaches that suggest functional innervation of isolated lymphocytes within the connective tissue of the mucosa (Ichikawa et al. 1992). There are also close appositions between axons and mast cells in the mucosa (Stead et al. 1989). Enteric interneurons Interneurons have been identified in all gut regions, and possibly vary between regions more than other neuron types. For example, the ileum and colon contain the same, or very similar, motor neurons and intrinsic primary afferent neurons, but their complements of interneurons are quite different (compare Figs 2.1A and 2.13). Within the myenteric plexus, the interneurons form chains of like neurons that run both orally and anally (Pompolo & Furness 1993, Portbury et al. 1995b, Young & Furness 1995). In the guinea-pig small intestine, three classes of descending interneurons and one class of ascending interneuron occur. Detailed studies of synaptic connections indicate that the chains formed by two of the types of descending interneuron interconnect (Mann et al. 1997). The ascending interneurons appear to be involved in local motility reflexes, as are two types of descending cholinergic neurons, those which contain NOS and those containing 5-HT (Chapter 3). Another type of descending interneuron, the ACh/SOM interneurons, might be linked to the passage of the migrating myoelectric complexes (MMC) along the intestine. These are Dogiel type III neurons with numerous branching, tapering, filamentous dendrites (Portbury et al. 1995b); they receive very few connections from IPANs (Stebbing & Bornstein 1996, Pompolo & Furness 1998), although responses of the neurons to distension have been recorded (Bornstein et al. 1991a, Thornton & Bornstein 2002). MMCs are waves of excitatory activity that are conducted anally along the gut that are mediated through the intrinsic neural pathways of the small intestine (Chapter 5). MMCs begin in the stomach and duodenum and pass along the full length of the small intestine, but are not conducted along the colon. It is thus pertinent that the ACh/SOM descending interneurons are found in both the ileum and the duodenum but not in the proximal or distal colon. The ACh/5-HT interneuron is possibly involved both in secretomotor and motility reflexes (Chapters 3, 6). Each of these three types of descending interneurons also provides innervation to submucosal ganglia. However, the ascending interneurons do not provide collaterals to submucosal ganglia; these ascending neurons and their terminals are
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immunoreactive for calretinin, but there are no calretinin-immunoreactive nerve terminals in submucosal ganglia (Brookes et al. 1991a). A small number of submucosal neurons in the guinea-pig ileum (fewer than 1%) have single axons that project to myenteric ganglia, but do not have collaterals to the mucosa or to submucosal arteries (Song et al. 1998). Thus these are interneurons connecting between the plexuses. They are immunoreactive for VIP. A small proportion (<1%) of submucosal neurons has NOS immunoreactivity (Furness et al. 1994). It is feasible that these are the same neurons, and that they represent “displaced” myenteric NOS/VIP interneurons. It has previously been suggested that a small number of neurons in the myenteric ganglia that have the phenotypes of submucosal secretomotor neurons are “displaced” secretomotor neurons (Brookes & Costa 2002). The idea that small numbers of neurons can be displaced from their normal position in layered structures seems to apply in other parts of the nervous system, for example in the retina, where “displaced” amacrine cells are described in the ganglion cell layer. It should be pointed out that in larger animals there are numerous NOS-immunoreactive neurons in the submucosal ganglia, some of which are almost certainly inhibitory neurons that provide innervation of the muscularis mucosae and external muscle. Intrinsic primary afferent neurons (intrinsic “sensory” neurons) The intrinsic reflex pathways that are involved in the control of physiological functions in the small intestine and colon, that is, gut movement, blood flow, and secretion, are activated through intrinsic neurons that respond to several stimuli, such as distension, luminal chemistry, and mechanical stimulation of the mucosa (Chapter 3). The intrinsic afferent neurons have now been positively identified as AH neurons with Dogiel type II morphology in the small intestine of the guinea-pig, and analogous neurons have been found in other species and regions. About 100 years ago, several authors showed that enteric motility reflexes could be elicited in segments of intestine that had no neural connections with the central nervous system and it was therefore assumed that primary afferent neurons were contained in the gut wall (Chapter 3). However, it was discovered at about the same time that reflexes, notably cutaneous vasodilator reflexes, could be initiated via axon collaterals even when the axons bearing the collaterals were disconnected from their cell bodies (Langley 1903). This led to a controversy about whether IPANs exist, which has only been resolved by their positive identification in the last decade. I will return to the question of whether some enteric reflexes are mediated through axon reflexes later in this chapter and in Chapter 3.
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Intrinsic primary afferent neurons: Why use this terminology? Cervero (1994) has provided a valuable summary of the conceptual differences between the afferent innervation of tissues and organs and their sensory innervation. As he points out, the activation of visceral receptors (such as arterial chemoreceptors, stretch receptors in the lung, and osmoreceptors in the liver) evokes no sensation, and therefore the term afferent is most appropriate to designate the neurons that carry this information. On the other hand, some neurons innervating the viscera do give rise to sensation, such as pain and warmth, and these are properly called sensory neurons (or sensory afferents; Grundy & Scratcherd 1989). Sherrington (1900) and Langley (1903) had earlier made the same distinction. Sherrington used it when referring to the production of pain by activation of fibers that normally do not cause sensation. He wrote: “The impulses from visceral fibers on the central nervous system appear hardly at all to elicit conscious sensations. When abnormally they do so, it is as though particular afferent nerves, which are not in the strict meaning of the term sensory nerves, can on occasion become sensory.” The important difference between visceral afferent neurons in general, and the subset of visceral afferents that mediate conscious sensation from the viscera, extends to the intrinsic afferent neurons within the intestine, and when the first clues about the identities of these neurons were being gathered, Kirchgessner and Gershon (1988) were careful to call them intrinsic primary afferent neurons (IPANs). Thus, when the IPANs are referred to as sensory, it should be remembered that they do not actually convey any sensation from the intestine. Afferent pathways, for example those that enter the central nervous system via the dorsal roots, involve neurons that connect in series. The first neurons in such series are referred to as primary, and neurons with which they communicate as secondary and higher-order afferent neurons. In accord with this nomenclature, the neurons within the enteric nervous system that detect the state of the intestine are referred to as primary afferent neurons because they are the first neurons of the intrinsic reflex pathways of the intestine. Afferent neurons carry information towards reflex centers or to integrating nerve circuits, whereas efferent neurons carry information away from reflex or integrating centers. Included among efferent neurons are motor neurons to muscles and glands. Matters are not quite as simple as this, because there are many examples of neurons that have more than one function. Good examples are motor (efferent) neurons to skeletal muscle. These neurons have collaterals that connect with neurons within the spinal cord (Renshaw 1946, Eccles et al. 1954). It would obscure their main function if these were to be called interneurons instead of being called motor neurons. A further example is spinal primary afferent neurons which, through transmitter release from
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their peripheral endings, cause vasodilation and plasma extravasation (Lewis 1927). Thus these primary afferent neurons, including some that supply the gut, also have efferent effects (Holzer et al. 1991, Holzer & Maggi 1998). IPANs probably also have efferent effects when they release transmitter onto the mucosal epithelium (see Chapters 3, 6). IPANs receive excitatory synapses and provide outputs to other neurons; in this context they can be considered to be interneurons. Another example of primary afferent neurons that receive synapses to their somas is trigeminal primary afferent neurons that have their cell bodies in the trigeminal mesencephalic nucleus (Honma et al. 2001). So, like the IPANs, these trigeminal primary afferent neurons could be considered to be interneurons. What properties are expected of primary afferent neurons? Primary afferent neurons transduce and encode information about the chemical environment and physical state of the tissues that they innervate, and convey this information to integrative circuitry through which the functional states of organs can be modified. A fundamental property is that the primary afferent neurons respond to adequate (that is, physiologically appropriate) stimuli in a manner that codes the intensities, durations, and patterns of stimuli. IPANs react to stimuli that are at the threshold for eliciting enteric reflexes, for example they respond to puffs of nitrogen gas that cause minimal movement of the mucosal villi (Gershon & Kirchgessner 1991) and have a low level of activity even when no deliberate stimulus is applied (Kunze et al. 1997). The activity of IPANs has been detected by intracellular and patch electrodes and by activity-dependent dyes. With these methods, IPANs have been identified that are responsive to mechanical distortion of the mucosa, to distortion of their processes within the myenteric plexus and to various chemicals applied to the mucosa (Gershon & Kirchgessner 1991, Kirchgessner et al. 1992, Bertrand et al. 1997, 1998, Kunze et al. 1998, 1999, 2000). As expected of primary afferent neurons, the responses of IPANs are graded with stimulus strength (Kunze et al. 1998), as are the reflexes that are evoked by their activation. In addition to transducing adequate physiological stimuli, IPANs can function as nociceptors, since their activation by noxious stimuli triggers protective responses (see below). Primary afferent neurons: intrinsic and extrinsic The state of the gastrointestinal tract is detected by three types of cells: primary afferent neurons; entero-endocrine cells and immune cells (Fig. 2.17). Each of these detecting systems is more extensive than those of non-digestive organs (Furness et al. 1999). About 20% of neurons in the enteric nervous
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Fig. 2.17 Three types of afferent signals that originate from the gastrointestinal tract, carried by hormones, the immune system, and neurons. Endocrine messages are carried by hormones released from entero-endocrine cells in the mucosal epithelium. The hormones enter the circulation, and can act at remote sites. They also act locally, on nerve endings, on the epithelium, and on cells of the immune system. Immune messages are conveyed by circulating lymphocytes that are activated by antigens presented to them from the lumen. Immune cells and cells of tissue defense, such as mast cells and macrophages, also release substances that act locally within the gut wall. Signals are also conveyed by neurons whose receptive endings are in the lamina propria, beneath the mucosal epithelium, in the muscle and in enteric ganglia. Some afferent neurons have cell bodies in the gut wall (IPANs and intestinofugal neurons) and the cell bodies of others are in extrinsic ganglia (extrinsic primary afferent neurons, see Fig. 2.18). Adapted from Furness et al. (2004a).
system, which contains about half a billion neurons in humans (Chapter 1), are intrinsic primary afferent neurons, and more than 50 000 nerve processes that reach the gut through the vagus and splanchnic nerves are also afferents. Furthermore, the gastroenteropancreatic endocrine system contains thousands of entero-endocrine cells, many of which react to their local environment, and from which more than 20 identified hormones are released. Finally, the gut immune system detects immunogens and contains 70–80%
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Fig. 2.18 The afferent neurons of the digestive tract. Two classes of intrinsic primary afferent neuron (IPAN) have been identified: myenteric IPANs that respond to distortion of their processes in the external muscle layers and, via processes in the mucosa, to changes in luminal chemistry, and submucosal IPANs that detect mechanical distortion of the mucosa and luminal chemistry. Extrinsic primary afferent neurons have cell bodies in dorsal root ganglia (spinal primary afferent neurons) and vagal (nodose and jugular) ganglia. Spinal primary afferent neurons supply collateral branches in prevertebral (sympathetic) ganglia and in the gut wall. Intestinofugal neurons are parts of the afferent limbs of entero-enteric reflex pathways. LM, longitudinal muscle; CM, circular muscle; MP, myenteric plexus; SM, submucosa; Muc, mucosa. Nerve endings in the mucosa are activated by hormones, most prominently 5-HT, released from entero-endocrine cells (arrows). Adapted from Furness et al. (2004a).
of the body’s immune cells. There is extensive interaction between these three systems, and physiological responses commonly involve actions through neurons, endocrine cells, and immune cells. Two broad classes of primary afferent neurons are associated with the gut: IPANs with cell bodies, processes, and synaptic connections in the gut wall and extrinsic primary afferent neurons (Figs 2.18, 2.19). Extrinsic primary afferent neurons have cell bodies in nodose and jugular ganglia (vagal afferents) or in dorsal root ganglia (spinal afferents). In addition, signals are
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Fig. 2.19 Diagram of a myenteric intrinsic primary afferent neuron. The IPANs are multipolar
Dogiel type II neurons. Action potentials initiated by physiologically appropriate stimuli and by noxious stimuli can traverse the cell bodies (transcellular conduction) or can be conducted to output synapses via an axon reflex (axon reflex conduction). Conduction across the cell body is modified by the synaptic inputs that it receives. The myenteric IPANs make synaptic connections with other neurons in the myenteric and submucosal ganglia. Adapted from Furness et al. (2004a).
carried by intestinofugal neurons that have cell bodies in the gut, but send processes to neurons outside the gut wall (see below and Chapter 5). Monitoring and control of the digestive system by neurons is hierarchical. The enteric nervous system is capable of generating appropriate local reflexes and also participates in reflexes between organs, for example, between the duodenum and stomach, in this case to regulate gastric emptying. Reflexes between parts of the digestive tract (entero-enteric reflexes) also travel via the prevertebral ganglia, thus bypassing the central nervous system (Chapter 5). Furthermore, signals are conveyed from the digestive organs via extrinsic primary afferents to the central nervous system, which trigger reflexes that act back on the digestive system. Some afferent signals to the CNS mediate coordination with other body systems, and some relate to sensations including discomfort, nausea, pain, and satiety. Characteristics of intrinsic primary afferent neurons IPANs that have been identified to date have Dogiel type II morphology. Interestingly, when Dogiel (1899) described these neurons, he guessed that they would be primary afferent neurons. Myenteric IPANs have axons that supply
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terminals around several types of nerve cells in myenteric ganglia, including other IPANs, interneurons, and motor neurons, and also provide inputs to submucosal ganglia. They receive synapses on their somas and on the initial parts of their axons (Pompolo & Furness 1988). Submucosal Dogiel type II neurons connect with other neurons in the submucosal ganglia (Furness et al. 2003a) and with neurons in myenteric ganglia (Kirchgessner & Gershon 1988, Song et al. 1998). In the guinea-pig small intestine, IPANs have AH electrophysiological characteristics, although some AH/Dogiel type II neurons without a prominent AHP have been reported (Clerc et al. 1998a). Neurons with the same shapes in the pig (Cornelissen et al. 2000) and mouse (Nurgali et al. 2004) often do not exhibit an AHP. As explained earlier in this chapter, the AHP can be suppressed by neurotransmitters and hormones, and whether the absence of the AHP means that the mechanism to generate it is missing cannot be decided without further experimentation. IPANs that respond to luminal chemicals Intracellular records from nerve cell bodies in the guinea-pig small intestine show that IPANs respond to chemicals, such as inorganic acids and shortchain fatty acids at neutral pH, applied to the luminal surface of the mucosa of the small intestine (Kunze et al. 1995, Bertrand et al. 1997). Activation of submucosal and myenteric neurons by glucose applied to the mucosa has also been reported (Kirchgessner et al. 1996). Neurons respond briskly at frequencies of up to about 20/s, and the responses were still observed by recording from the cell bodies of IPANs after synaptic transmission had been blocked (Kunze et al. 1995, Bertrand et al. 1997). Thus the effect on IPANs was direct, not indirect through an intervening synapse. IPANs sensitive to stretch or distortion at the level of the myenteric plexus IPANs possess mechanosensitive ion channels that allow them to transduce distortion of their processes into action potential firing (Kunze et al. 1998, 2000). In experimental conditions, distortion can be applied directly to the processes or can be caused by muscle movements, because the muscle is adherent to the processes via collagen fibers. From experiments performed on longitudinal muscle–myenteric plexus preparations from the small intestine, with some residual attached circular muscle (Kunze et al. 1999), the mechanism through which these neurons are activated by muscle movement has been deduced (Fig. 2.20). It appears that the muscle pulls on connective tissue elements, which are in turn connected to IPANs. The ensuing distortion of
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Fig. 2.20 Mechanism of activation of mechanosensitive IPANs by contraction of the wall of the intestine, which has been deduced from experiments in which recordings were made from the neurons during the application of stretch to the tissue. This opens stretch-activated channels (SACs) in the muscle membrane, which results in muscle contraction. The contracting muscle distorts the IPAN processes. If the muscle SACs are blocked by gadolinium, or if contraction is prevented by isoprenaline or nicardipine, excitation of IPANs by stretch is diminished. On the other hand, when the muscle is contracted by opening L-type Ca2+ channels with BK 8644, IPANs are activated. The muscle pulls on IPAN processes through connective tissue which can be weakened by dispase, preventing IPAN activation in response to stretch. The distortion of processes of IPANs opens Gd3+-insensitive neuronal SACs, which cause depolarization and action potential initiation. IPANs can also be activated experimentally by direct distortion of their processes. Adapted from Furness et al. (2004a).
the IPANs opens mechanosensitive channels that cause local depolarizations (generator potentials) and initiation of action potentials. Pressure, applied with a fine probe to the processes of IPANs within the myenteric plexus evokes generator potentials that are recorded at distances of 0.1 to 0.5 mm from the cell body (Kunze et al. 2000). Generator potentials and action potentials initiated by pressing the myenteric processes of IPANs, as well as action potentials generated by stretching the muscle, were not affected by lowering Ca2+ in the external solution (Kunze et al. 1999, 2000), indicating that the neurons were directly (not synaptically) activated and that the mechanosensitive channels are not Ca2+ channels. In addition, action potentials generated in IPANs by contracting the muscle were unaffected by gadolinium, which blocks mechanosensitive channels in intestinal muscle cells (Kunze et al. 1999). The neurons fire action potentials phasically, when their processes are directly deformed, or at the beginning of stretch applied to the longitudinal muscle (Kunze et al. 2000). During maintained stretch, the muscle contracts more or less rhythmically, and IPANs continue to discharge (Kunze et al. 1998). The rate of discharge is proportional to the degree of distension. However, the discharge of action potentials was abolished if the muscle contraction was prevented by muscle relaxants, either isoprenaline (an agonist of β-receptors for catecholamines) or nicardipine. This indicates that active tension in the muscle contributes to the excitation of the tensionsensitive IPANs. The involvement of the muscle is interesting, because it has
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long been known that intestinal muscle cells are directly sensitive to stretch and respond to it by contracting (Bülbring 1955). This reaction of the smooth muscle may be integral to the response of IPANs during sustained stretch. When the tissue was exposed to a proteolytic enzyme, to weaken connective tissue links between the muscle and the IPANs, activation of IPANs by muscle contraction was prevented (Kunze et al. 1999). The IPAN cell bodies are hyperpolarized when a small area of soma membrane is stretched (Kunze et al. 2000). This occurs through the opening of BK-type Ca2+-activated K+ channels on the cell soma, which appear to be directly distortion-sensitive. Myenteric nerve cell bodies are deformed by muscle movement (Gabella & Trigg 1984), and it has been speculated that distortion of IPANs when the intestine contracts may reduce their excitability, as part of a protective mechanism that limits the strength of reflex contraction of the intestine (Kunze et al. 2000). There is evidence that neurons other than Dogiel type II neurons are mechanosensitive. Kunze et al. (1999) recorded from one Dogiel type I neuron that responded directly to stretch. In flat sheet preparations of guinea-pig colon in which the circular muscle was maintained attached to the myenteric plexus, stretching the circular muscle triggered reflexes via mechanosensory neurons that were activated even when the smooth muscle was paralyzed by nifedipine (Spencer et al. 2002, 2003). Excitatory junction potentials oral and inhibitory junction potentials anal to the stimulus were synchronized only in preparations longer than 7 mm, and they were not triggered in preparations 3 mm long. From these results, it was deduced that Dogiel type II neurons were not the mechanosensitive neurons through which the reflexes were initiated, because these would be expected to have outputs to motor neurons in short preparations, and because, by extrapolation from the small intestine, their activity was predicted to be blocked by nifedipine (Spencer et al. 2002). Spencer et al. (2002) suggested that the mechanosensory neurons were ascending interneurons. However, the projections of Dogiel type II neurons, their connections and their responsiveness to mechanical distortion in the colon have not been determined by direct recording. Prior to the studies that identified AH/Dogiel type II neurons as IPANs, experiments using extracellular recording techniques had identified mechanosensitive neurons in the myenteric plexus. It was found that some myenteric neurons fired action potentials when the myenteric plexus was pressed with a fine probe, but that responses were not elicited by pressing on the adjacent muscle (Wood 1970, 1973, Ohkawa & Prosser 1972). The mechanoreceptive neurons responded to pull on the muscle, 0.5–2 mm from the recording electrode and to contractile activity in the circular muscle (Wood 1970). In these experiments it was not possible to determine the morphologies of the neurons, but these mechanoreceptive neurons have properties that are similar to the
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AH/Dogiel type II neurons from which intracellular records have been obtained in response to tension in the external muscle or pushing on the adjacent myenteric plexus where their processes are found (Kunze et al. 1999, 2000). Mucosal mechanoreceptors Secretomotor reflexes can be elicited from preparations that contain the mucosa and submucosa, but not the external muscle or myenteric ganglia, implying that there are IPANs in submucosal ganglia (Chapter 3). Additional evidence comes from experiments in which c-Fos immunoreactivity was detected in submucosal nerve cells after the mucosa had been distorted by puffs of nitrogen gas that were ejected from a pipette (Kirchgessner et al. 1992). The c-Fos expression was abolished by tetrodotoxin, but not by the nicotinic receptor blocker hexamethonium, suggesting that it was produced in the cell bodies of IPANs that had processes in the mucosa. Styryl dyes, which are taken up by the endings of active neurons and transported back to the cell bodies, have also been used to identify IPANs that are mucosal mechanoreceptors (Kirchgessner et al. 1996). Distortion of the villi by puffs of nitrogen gas caused styryl dye labeling in neurons of numerous submucosal ganglia and in a few myenteric ganglia, in the presence of hexamethonium, to block fast excitatory synaptic transmission. Fibers in myenteric ganglia, presumed to be the axons of submucosal nerve cells, were labeled. These results suggest that cell bodies of mucosal mechanoreceptors are in submucosal ganglia and project to the myenteric plexus (Fig. 2.18). This projection has been confirmed by structural studies (Kirchgessner & Gershon 1988, Song et al. 1998). Distension stimuli can activate both mucosal mechanoreceptors and distension-sensitive neurons (myenteric IPANs). This explains why both myenteric and submucosal neurons were revealed by styryl dyes after distension (Kirchgessner et al. 1996). Whether cell bodies of stretch-sensitive neurons are present in the submucosal ganglia, as well as in myenteric ganglia, has not yet been determined. The axons of submucosal IPANs project more or less vertically to the underlying mucosa and thus the recording of direct responses of submucosal IPANs to mechanical stimulation of the mucosa has not been reported. However, recordings have been made from second-order submucosal neurons (Pan & Gershon 2000). These respond to mechanical stimulation of the mucosa with fast and slow EPSPs. Roles of IPANs as nociceptors In the small intestine and colon, protective secretory and motility responses are initiated by irritants that are included in enemas, by bacterial products, and by parasitic infestations (see Chapters 5 and 6). The intestine exhibits
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exaggerated secretomotor reflexes, which are manifested as diarrhea, when there are excessive levels of bacteria and bacterial toxins in the gut lumen (Lundgren et al. 2000, Lundgren 2002). Powerful propulsive reflexes are also triggered by bacteria or parasites in the gut lumen to expel these organisms (Sukhdeo & Croll 1981, Mathias et al. 1982, Collins 1996, Vallance et al. 1997). Thus the IPANs appear to act as detectors of noxious conditions in the gut, as well as reacting to benign stimuli to elicit enteric reflexes. Binding of the plant lectin IB4 appears to be specific to nociceptive neurons, but it does not bind to non-nociceptive neurons, in dorsal root and nodose ganglia (Stucky & Lewin 1999, Gerke & Plenderleith 2001). Enteric IPANs, but not other enteric neurons, also have high affinity binding sites for IB4 (Hind et al. 2005). Intermediates in the activation of IPANs The mucosal epithelium separates the nerve endings of IPANs from the luminal environment. Therefore luminal chemicals can only influence the nerve endings if they provoke a signal from epithelial cells that affects the nerve endings or if the chemical stimulants cross the epithelium. There is evidence that 5-HT (5-hydroxytryptamine, serotonin), which is a potent stimulant of the endings of IPANs, is an intermediate in enteric reflexes elicited by changes in the chemical contents of the lumen or by mucosal distortion (Kirchgessner et al. 1992, Bertrand et al. 1997, 2000, Pan & Gershon 2000). 5-HT is released when the mucosa is mechanically stimulated and the reflex responses are antagonized by drugs that block 5-HT receptors (Bülbring & Crema 1959, Neya et al. 1993, Foxx Orenstein et al. 1996, Grider et al. 1996). Furthermore, mechanical stimulation of the mucosa causes c-Fos induction in IPANs in submucosal ganglia, and this induction is blocked by an antagonist of 5HT receptors (Kirchgessner et al. 1992). Recent data indicate that ATP could be also involved in communicating excitation from entero-endocrine cells to the mucosal endings of IPANs (Bertrand & Bornstein 2002). Other hormones that are contained in gut endocrine cells, such as cholecystokinin (CCK) and motilin, are released by nutrients and act on neurons, but have not been tested for their possible roles as intermediates in enteric reflexes. Polymodal nature of IPANs IPANs in myenteric ganglia that respond to chemicals applied to the mucosa can also respond to mechanical stimulation of the mucosa (Bertrand et al. 1997, 1998). About 60% of myenteric IPANs responded to chemicals (acid, base, or fatty acid) applied to the mucosa (Bertrand et al. 1997) and about 80% responded when the intestine was stretched in the circumferential direction (Kunze et al. 1998). Thus, some IPANs are activated by both mucosal
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chemical stimulation and stretch. Although these data indicate that individual IPANs are polymodal, it is possible that subgroups of IPANs respond more strongly to one type of stimulus than to another. The intestine is never likely (other than in experimental conditions) to be exposed to only a single stimulus; it must normally encounter signals from chemicals in the lumen, from the bulk of its contents, and from its own contractile activity. Synaptic inputs to IPANs Although it was originally thought that AH/Dogiel type II neurons did not receive any synaptic input (Hirst et al. 1974), inputs were only stimulated at low frequencies for short periods of time in that study. It was soon discovered that slow EPSPs could be elicited in AH neurons by short bursts of high-frequency stimulation (Wood & Mayer 1978). Much later, it was found that they also respond to low-frequency activation of synaptic inputs when stimulation is maintained for several minutes (Clerc et al. 1999). It has been confirmed that fast excitatory synaptic inputs to these neurons are rarely seen in the guineapig small and large intestine (Bornstein et al. 1994) or in the mouse colon (Furukawa et al. 1986, Nurgali et al. 2004) and, when recorded, fast EPSPs are of low amplitude (Iyer et al. 1988, Bornstein et al. 1994, Tamura et al. 2001). In contrast, fast EPSPs are reliably recorded in Dogiel type II neurons of some other regions and species, such as the pig (Cornelissen et al. 2001). There may be an increase in the occurrence and amplitudes of fast synaptic events in IPANs during inflammation of the intestine (Linden et al. 2003). Synaptic interactions between IPANs Both physiological and ultrastructural studies indicate that IPANs synapse with other IPANs (Pompolo & Furness 1988, Kunze et al. 1993). They also receive many other synaptic inputs whose origins have not been determined (Pompolo & Furness 1988). The axons of IPANs give rise to very dense networks of varicose terminals, which appear to surround all nerve cell bodies in their own and adjacent ganglia (Furness et al. 1990b, Bornstein et al. 1991b). Electron microscope studies show that these terminals form synapses on nerve cell bodies, including the cell bodies of IPANs (Pompolo & Furness 1988). Moreover, when the cell bodies of two IPANs were impaled with microelectrodes, and action potentials were evoked by stimulus pulses passed through one electrode, the other IPAN responded with slow EPSPs (Kunze et al. 1993). Pharmacological experiments support the conclusion that one origin of slow EPSPs in IPANs is synapses made on them by other IPANs. In the guinea-pig small intestine, IPANs are immunoreactive for tachykinins, and
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antagonists of tachykinin receptors partially block the slow EPSPs in these cells (Neunlist et al. 1999, Alex et al. 2001, Johnson & Bornstein 2004). The conclusion that slow EPSPs in IPANs arise from other IPANs is consistent with experiments in which the axons of ascending and descending interneurons were severed (Bornstein et al. 1984b). In areas between lesions, where the severed axons of interneurons had degenerated but endings of IPANs remained, slow EPSPs of usual amplitude were recorded in the cell bodies of IPANs. Thus, data from several experimental approaches all indicate that the IPANs form interconnected networks, and, because transmission at the connections between IPANs is excitatory, these networks would be self-reinforcing (Wood 1994, Bertrand et al. 1997, Kunze & Furness 1999). Intestinofugal neurons Intestinofugal neurons (Fig. 2.18) are neurons that have cell bodies in the gut wall and send their processes to prevertebral ganglia, where they form synapses with post-ganglionic sympathetic neurons (Kuntz 1938, Szurszewski & Miller 1994). Chapter 5 contains a more complete description of these neurons and discussion of their roles. They have been called IFANs (intestinofugal afferent neurons) because they carry afferent signals from the gut (Szurszewski et al. 2002). In small mammals the cell bodies of intestinofugal neurons occur in myenteric ganglia; the majority of these neurons have Dogiel type I morphology, but some have Dogiel type II morphology. In pigs, some of the intestinofugal neurons are in the submucosal ganglia (Barbiers et al. 1993). This may also apply to other large mammals. These neurons are in the afferent limbs of nerve pathways that carry entero-enteric reflexes, that is, reflexes that are initiated from the intestine, and which act back on the stomach or intestine (Szurszewski & Miller 1994). The sympathetic neurons that are innervated by intestinofugal neurons are motility inhibiting and secretomotor inhibiting neurons. The roles of intestinofugal neurons and of entero-enteric inhibitory reflexes in controlling motility are reviewed in Chapter 5. An unusual group of intestinofugal neurons, which have been little investigated, are neurons with cell bodies in the rectum that project to the spinal cord (Neuhuber et al. 1993), and similarly rare neurons that project from the gastric myenteric plexus directly to the dorsal vagal complex in the brainstem (Holst et al. 1997). Neurons in human intestine with equivalence to those investigated in laboratory animals Pharmacological investigations of neurotransmission and immunohistochemical studies of innervation patterns both indicate that muscle motor
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neurons and secretomotor neurons in human are similar to those in other mammals, and that human equivalents of IPANs can be identified. The interneurons are more difficult to study, but do appear to be similar to those of other mammals. The muscle layers of the human intestine are densely innervated by nerve fibers showing immunoreactivity for tachykinins (Ferri et al. 1983, Llewellyn Smith et al. 1984), which are markers of excitatory muscle motor neurons, and for VIP and NOS (Ferri et al. 1983, De Giorgio et al. 1994), which are contained in inhibitory motor neurons. Double-labeling shows that tachykinins and VIP are in different fibers innervating the muscle (Wattchow et al. 1988). Furthermore, nerve cell bodies which are immunoreactive for these markers occur in myenteric ganglia (Llewellyn Smith et al. 1984, Porter et al. 1997). Retrograde transport shows that the majority of neurons that project to the muscle in humans have Dogiel type I morphology (Wattchow et al. 1995) and that the tachykinin-immunoreactive neurons project orally, whereas the VIPimmunoreactive fibers project anally (Wattchow et al. 1997), as they do in other mammals. It has also been shown that the orally projecting neurons are immunoreactive for choline acetyltransferase (ChAT), and are thus presumed to be cholinergic, and the anally projecting neurons are nitric oxide synthase (NOS) immunoreactive (Porter et al. 1997). Thus, the muscle motor neurons in humans appear to be analogous to those in other mammals, the excitatory motor neurons being Dogiel type I cholinergic neurons with tachykinin immunoreactivity and the inhibitory motor neurons being anally projecting Dogiel type I neurons with NOS and VIP immunoreactivity. These deductions are supported by pharmacological investigation of enteric reflexes in human intestine (Grider 1989a), which identify cholinergic motor neurons in the ascending reflex pathway, and VIP neurons in the descending pathway. As in other mammals, the human intestinal mucosa is densely innervated by VIP-immunoreactive nerve fibers (Ferri et al. 1982a, 1983). Most of these probably come from submucosal neurons, over 40% of which are VIP immunoreactive and which are labeled by retrograde transport from the mucosa (Porter et al. 1999, Hens et al. 2001). VIP is a stimulant of secretion in the human intestine (Krejs 1982). These data, considered together, suggest that VIP-containing secretomotor neurons occur in human intestine. In the small intestine, most submucosal VIP neurons stain for the vesicular acetylcholine transporter, whereas these form largely separate populations in the colon (Anlauf et al. 2003). In the rectum, the human VIP neurons stain for ChAT, again suggesting that they are cholinergic (Schneider et al. 2001). Thus ACh and VIP may be co-transmitters in the small intestine and rectum, but perhaps not in the colon, where separate cholinergic and non-cholinergic secretomotor neurons could occur.
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Brehmer (2004b) has investigated the identity of IPANs in the human small intestine. He was able to identify the Dogiel type II neurons in myenteric ganglia stained for neurofilament proteins. Most of these neurons were immunoreactive for somatostatin, tachykinins, and calretinin. Many of the neurons projected a process towards the mucosa, which is consistent with earlier work that showed that myenteric Dogiel type II neurons were retrogradely labeled after dye was applied to the mucosa of the human intestine (Hens et al. 2001). In other species, Dogiel type II neurons are also immunoreactive for tachykinins and calcium-binding proteins. Thus, although the information is incomplete, it seems likely that neurons in the human intestine have similarities in their primary transmitters and principal chemical markers to those in other mammals. Summary and conclusions Essentially all functional types of enteric neuron have been characterized by their shapes, neurochemistries, and cell physiological properties in the guinea-pig, and their orthologs in human and several other species have been identified. There are 15 types of neuron in the small intestine, slightly more in the colon, and slightly fewer in the stomach and esophagus. The neurons can be placed in four functional classes, intrinsic primary afferent neurons (IPANs, also referred to as intrinsic sensory neurons), interneurons, motor neurons, and intestinofugal neurons. IPANs occur in the small and large intestine where they are neurons with multi-axonal Dogiel type II morphology. They have endings in the mucosa that respond to physiological stimuli, and other processes that form synapses with interneurons and motor neurons, as well as with other IPANs. Structural and electrophysiological evidence suggests that there are very few IPANs in the stomach, but it may be that they have a different morphology and have escaped identification. IPANs in guinea-pigs have very distinctive electrophysiological characteristics and are called AH neurons for their electrophysiological features, which include a prominent after-hyperpolarizing potential (the AH or AHP) that follows the action potential. Interneurons are mono-axonal neurons, almost all of which have what is called S-type electrophysiological properties. They project small distances along the gut wall (in comparison with its length) and form chains of like neurons. They connect with each other and with motor neurons. Motor neurons include excitatory and inhibitory muscle motor neurons, secretomotor neurons, secretomotor/vasodilator neurons (neurons whose axons divide to supply both intramural arterioles and the secretory epithelium), and neurons that innervate entero-endocrine cells. Motor neurons are mono-axonal, and almost all have S-type electrophysiological properties.
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Intestinofugal neurons have cell bodies in enteric ganglia, but their axons project to sympathetic ganglia and in some cases to the sacral spinal cord and brainstem. Those that project to sympathetic ganglia form parts of the afferent limbs of entero-enteric reflexes. Enteric neurons contain many substances that provide them with a neurochemical signature. These signature molecules include the primary transmitters of the neurons, the synthesizing enzymes for the transmitters, molecules that have transmitter or hormonal roles at other sites in the body, but are not necessarily transmitters of the enteric neurons that harbor them, and other markers, such as calcium binding proteins. Thus a chemical code can be found that defines enteric neurons, and distinguishes neurons of each functional type, but the codes commonly differ between gut regions or between species. The part of the chemical code that relates to the primary transmitters of the neurons is conserved.
3: Reflex circuitry of the enteric nervous system
Evolution of ideas about enteric circuitry Structural and functional data that were gathered from the middle of the 19th to the beginning of the 20th century led to the hypothesis that motility and secretomotor reflexes are mediated through the intrinsic ganglia that were well known from the work of Remak, Meissner, Auerbach and other mid-19th century anatomists. In relation to secretion, Pye-Smith hypothesized in 1874 that “the nerves immediately inducing secretion are probably the ganglia contained in Meissner’s plexus, the short afferent fibers passing from them to the mucous membrane, and the short secreting fibers passing from them to the secreting glands” (Pye Smith et al. 1874). In the case of motility reflexes, these were already assumed to be dependent on the enteric ganglia by Lister, who wrote in 1858 “To sum up …, it appears that the intestines possess an intrinsic ganglionic apparatus which is in all cases essential to the peristaltic movements, and, while capable of independent action, is liable to be stimulated or checked by other parts of the nervous system” (Lister 1858). An experimental proof of the assumption of independent action had to await the investigations by Langley and Magnus (1905), who demonstrated that motility reflexes occur in segments of the intestine after the nerves leading to them are cut and time is allowed for their degeneration. A number of investigators tried to draw conclusions about the circuitry from histological examination of the enteric nervous system, but these studies were confounded by the huge numbers of intertwining axons that occur in myenteric ganglia. Nevertheless, some researchers were able to follow processes of enteric neurons, and found evidence of connections between them (Kuntz 1922) and of the projections of myenteric neurons to the muscle and the mucosa (Cajal 1911, Müller 1920). Physiological observations were also used to make deductions concerning the organization of the circuits (e.g. Magnus 1904b, Fleisch & von Wyss 1923, Hukuhara et al. 1958, Hukuhara & Miyake 1959). However, progress was very slow and experiments were often ambiguous.
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A conjunction of new methods in the 20 years from about 1975 meant that more progress was made in this period than in more than a century of previous effort, both in the identification of individual neurons (see Chapter 2), and in the analysis of the circuits to which they belong. This chapter deals with nerve circuits that control functions of the small and large intestines, esophagus, and stomach of mammals. The structure and physiology of the enteric nervous system has been investigated in other vertebrate classes, including fish, amphibians, and birds (Olsson & Holmgren 2001). The authors conclude that the enteric nervous system and control of gastrointestinal function is amazingly similar across vertebrate classes. I have also included in this chapter a brief description of the sympathetic innervation of the gastrointestinal tract, and of the entero-enteric reflex pathways that connect parts of the gastrointestinal tract via sympathetic prevertebral ganglia. Motility controlling circuits of the small and large intestine Motility is a broad term, encompassing several movement behaviors of the intestine, including propulsion (peristalsis), mixing movements, and the migrating complexes of the small intestine (Chapter 5). However, almost all the functional experiments that provide useful information about the neuronal connections within enteric nerve circuits come from studies of propulsive reflexes. When it is considered that each of the patterns of movement must be derived from an integrated activity of the same sets of neurons, analysis of propulsive reflexes is as good a starting point as any other to deduce the organization of the circuits. The simplest stereotyped reflexes that are related to propulsion are the polarized contraction of the circular muscle oral to a physiological stimulus and the relaxation that occurs on the anal side (Mall 1896, Bayliss & Starling 1899). By a physiological stimulus I mean a stimulus whose nature and intensity mimics a naturally occurring stimulus. When an artificial bolus is introduced into the lumen, the relaxation and contraction move the bolus along the intestine; a relaxation is observed before the bolus arrives at the recording point and the contraction is observed after it passes (Fig. 3.1). Analysis of the reflex pathways can be simplified by applying a localized (stationary) stimulus to mimic the stimulus that would be made by the intestinal contents, while avoiding propulsion of the contents (Costa & Furness 1976). Useful information about the pharmacology of reflexes can also be obtained by filling a segment of intestine with fluid and measuring the intraluminal pressure changes that occur when the intestine propels the fluid (Fig. 3.2). This approach is generally known as the Trendelenburg method (Trendelenburg 1917), of which there are many variants. Although these techniques are
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Fig. 3.1 Record of the movements of the circular and longitudinal muscle of the dog small intestine, in vivo, as a bolus passes, published by Bayliss and Starling (1899). At time A, a bolus was inserted 12 cm oral to the recording point. At the times marked by the dots the bolus was 7.5, 5 and 2.5 cm oral to the recording point. Between B and C it was at the recording point. Note that both muscle coats relaxed as the bolus approached, and contracted together after it passed. These records demonstrate the presence of descending inhibitory and ascending excitatory reflexes.
attributed to Trendelenburg, similar methods were used by earlier investigators (Mall 1896, Gayda 1913). The observation that propulsive reflexes occur in segments of intestine that are removed from animals and placed in an artificial nutrient solution (Mall 1896), and that they also occur in the intestine in vivo after all connections with the central nervous system are severed (Bayliss & Starling 1899), indicates that the nerve circuits for their generation are in the bowel wall. Langley realized that the severed ends of extrinsic neurons did not immediately die, and that it was possible that stimulation of these disconnected endings could initiate a reflex response. He therefore conducted experiments in which he cut the nerves leading to the intestine in a living animal, waited until the endings that were disconnected from their cell bodies of origin had died, and then removed the intestine. This procedure is known as degenerative section of nerves. Propulsive reflexes were recorded in isolated segments of rabbit colon after this degenerative section of the extrinsic nerves (Langley & Magnus 1905). Propulsive reflexes, which appear no different from those in normally innervated intestine, have also been recorded following degenerative section of extrinsic nerves of the guinea-pig ileum (Bülbring et al. 1958b, Furness et al. 1995a), guinea-pig colon (Crema et al. 1970), dog duodenum (Thomas & Baldwin 1971), cat jejunum (Bülbring et al. 1958b), and the smooth muscle part of the cat esophagus (Roman 1982). Moreover, intestinal transit and the passage of the MMC are unaffected after intestinal autotransplantation, which effectively provides extrinsic denervation of the intestine (Sarr et al. 1989, Iwanami et al. 2003). Interestingly, it is also possible to generate reflex responses that appear to depend on extrinsic axons, which remain viable after isolation of the intestine in vitro (Grider & Jin 1994).
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Fig. 3.2 The Trendelenburg method of recording reflexes in the intestine. A: Drawing of the apparatus from Trendelenburg’s paper of 1917. A segment of intestine is set up as a blind sac. The sac fills from a pressure head, and reflexly contracts to expel the fluid. The movement of fluid into and out of the segment and changes in length of the segment are recorded. B: A modification of the method, based on Bülbring et al. (1958a), in which fluid flows through the segment and is collected. Length changes, intraluminal pressure, and amount of fluid expelled are recorded.
Polarized reflexes are elicited by mechanical or chemical stimulation of the mucosa (Hukuhara & Miyake 1959, Smith & Furness 1988), and by distension (Bayliss & Starling 1899). Intrinsic primary afferent neurons (IPANs) that respond to each of these stimuli have been identified (Chapter 2). Reflex
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responses to distension occur in segments of intestine from which both the mucosa and the submucosa have been removed (Magnus 1904b, Costa & Furness 1976, Yokoyama & Ozaki 1980, Tsuji et al. 1992, Spencer et al. 2002), implying that sensitive parts of IPANs that respond to stretch are in the external musculature, and the cell bodies of interneurons and motor neurons, sufficient to maintain the reflexes, are in the myenteric plexus (Fig. 3.3). This does not mean that sensitive endings of IPANs in the mucosa do not normally contribute, merely that activation of stretch-sensitive processes of IPANs in the external layers is sufficient to generate the reflexes. In the intact intestine of human and other large mammals, many motor neurons to the external muscle are in submucosal ganglia (Chapters 1, 2) and these submucosal neurons almost certainly contribute to motility reflexes. It has been demonstrated in both dog and guinea-pig that, when the myenteric plexus is
Fig. 3.3 The circuitry for propulsive reflexes in the intestine. A: Simplified representation of the connections of motility-controlling circuits, illustrating the passage of axons between the layers. Motility reflexes are initiated through IPANs, some with cell bodies in myenteric ganglia and some in submucosal ganglia. The IPANs connect with chains of interneurons that run orally and anally within the myenteric plexus and directly with motor neurons (shown in panel B). Some IPAN processes run for long distances in the anal direction. The interneurons connect with motor neurons that innervate the muscle layers. In many species, cell bodies of motor neurons also occur in submucosal ganglia. (Continued.)
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Fig. 3.3 (Continued.) The circuitry for propulsive reflexes in the intestine. B: Reflex pathways depicted in a short segment of intestine to illustrate circumferential projections of neurons. The first parts of descending inhibitory and ascending excitatory reflex pathways for propulsive reflexes are depicted. Intrinsic primary afferent neurons (IPANs) are circumferentially oriented and form self-reinforcing networks. They provide outputs to ascending and descending interneurons and monosynaptic connections to motor neurons. One type of descending interneuron (bottom part of diagram), which is involved in conducting the migrating myoelectric complex along the small intestine, receives few inputs from IPANs, but connects with motor neurons.
severed between the point of initiation of a reflex and the point of contraction or relaxation of the muscle, the response in the muscle is prevented, but the reflexes persist after the submucosal plexus is severed between points of stimulus and response (Hukuhara et al. 1958, Costa & Furness 1976, Smith & Furness 1988) (Fig. 3.4). In cats, severing the myenteric plexus, with the submucosal plexus remaining intact, also prevented propulsive reflexes (Cannon 1912). Thus the pathways for the propulsive reflexes travel along the intestine in the myenteric plexus (Fig. 3.3). The presence of the cell bodies of interneurons and motor neurons of intrinsic reflexes in the myenteric plexus was confirmed in experiments in which synaptic activity was recorded from the neurons when stimuli that elicit motility reflexes were applied to the intestine (Hirst & McKirdy 1974a, Hirst et al. 1975, Bornstein et al. 1991a, Smith et al. 1992, Thornton & Bornstein 2002). As reviewed in Chapter 2, interneurons form synaptically connected chains that run along the intestine in the myenteric plexus and the IPANs make synapses with each other at which slow EPSPs are recorded (Fig. 3.3). Most of the final motor neurons in the reflex pathways innervate the smooth muscle close to their cell bodies, although a minority of circular muscle motor neurons run for considerable distances along the intestine in the guinea-pig, up to 30 mm for the descending inhibitory neurons and about 10 mm for the ascending excitatory motor neurons (Chapter 2).
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Fig. 3.4 The effect of lesioning
the myenteric plexus on the conduction of a reflex along the intestine. In the left-hand panel, contraction of the circular muscle oral to a stimulus (upper trace) and relaxation anal (lower trace) are recorded. The mucosa between the recording points was mechanically stimulated at the downward deflection of the marker trace (stimulus). Before repeating the stimulus and obtaining the records in the right panel, the myenteric plexus was transected just oral and just anal to the stimulus site. This prevented the reflex responses. Recordings are from the dog jejunum, time marker indicates 6 s intervals. Reproduced from Hukuhara et al. (1958).
The interneurons of motility reflexes The majority of the detailed quantitative data that exist on the identification of interneurons come from studies of the guinea-pig small intestine, and, unless it is indicated otherwise, the quantitative information that is alluded to below applies to this region. The general principles of organization of the enteric circuits apply to all mammals. Ascending interneurons In the guinea-pig small intestine there is only one type of ascending interneuron. These have medium-sized to large cell bodies with Dogiel type I morphology and are immunoreactive for calretinin and ChAT. All calretinin-immunoreactive nerve terminals in the myenteric ganglia arise from these neurons (Brookes et al. 1991a, Pompolo & Furness 1993), so calretinin immunoreactivity can be used to analyze the connections made by the neurons. Approximately 12% of the synapses on other calretinin-immunoreactive interneurons are calretinin immunoreactive, indicating that the interneurons form interconnected chains (Pompolo & Furness 1995). Many other synaptic inputs to the ascending interneurons (70% of inputs) are calbindin immunoreactive, and the remaining inputs appear to come from the cholinergic descending interneurons with NOS immunoreactivity (Pompolo & Furness 1995). They receive few inputs from the other classes of descending interneuron (ChAT/5-HT and ChAT/SOM interneurons).
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Physiological studies (see above) and pharmacological studies, in some cases using blockers of synaptic transmission in multichambered organ baths (Smith & Furness 1988, Tonini & Costa 1990, Johnson et al. 1996), indicate that ascending interneurons connect with excitatory motor neurons that innervate the circular muscle, although these synapses have not been studied by electron microscopy. Transmission is predominantly cholinergic, through nicotinic receptors (Chapter 4). About 30% of synaptic inputs to calretinin-immunoreactive excitatory longitudinal muscle motor neurons are from ascending interneurons and about 35–40% are from IPANs (Pompolo & Furness 1993, 1995). The inputs from IPANs indicate that there are short reflex pathways that are monosynaptic (they do not involve interneurons). In the guinea-pig colon there are three classes of ascending interneuron, in contrast to the single class seen in the small intestine (Lomax & Furness 2000). Neurons of each of the three classes are ChAT immunoreactive, which is consistent with ascending reflexes in the colon utilizing cholinergic neuroneuronal transmission. It is possible that in addition to interneurons of the local reflexes there are interneurons of pathways originating from pelvic nerves and involved in defecation in the colon. Descending interneurons Most, probably all, descending interneurons are also cholinergic, as indicated by their immunoreactivity for ChAT, but transmission in local reflexes is not purely cholinergic (Chapter 4). In guinea-pig small intestine, the classes of descending interneuron are those immunoreactive for ChAT plus somatostatin (SOM), those immunoreactive for ChAT plus nitric oxide synthase (NOS), VIP and other substances, and those immunoreactive for ChAT plus 5-HT (Chapter 2, Table 2.1). The ChAT/5-HT neurons do not make connections with the inhibitory muscle motor neurons (Young & Furness 1995), and pharmacological studies, which show a lack of effect of receptor antagonists, suggest that 5-HT is not a transmitter in local descending inhibitory reflexes (Yuan et al. 1994); the ChAT/5-HT interneurons are involved in descending excitatory reflexes (see below). Electron microscope studies show that the ChAT/SOM neurons, which have a distinct filamentous morphology, receive very few inputs from IPANs, whereas over 80% of their inputs are from other ChAT/SOM neurons. On the basis of these observations, it has been suggested that ChAT/SOM descending interneurons are a conduit for the passage of migrating myoelectric complexes (MMCs) along the guinea-pig intestine (Pompolo & Furness 1998). This is consistent with electrophysiologic studies show that the majority of ChAT/SOM neurons receive inputs from oral, but only one-sixth receive local inputs that are activated by stimulation circum-
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ferential to the cell body, which would activate the circumferential processes of IPANs (Stebbing & Bornstein 1996). Nevertheless, ChAT/SOM filamentous interneurons do respond to distortion of the mucosa or to local distension (Bornstein et al. 1991a, Thornton & Bornstein 2002). The ChAT/NOS interneurons form descending chains (Young et al. 1995) and pharmacological studies also indicate that the NOS interneurons are involved in descending inhibitory reflexes (Yuan et al. 1995). Moreover, electron microscope and high-resolution confocal studies have shown that a high proportion of synaptic inputs to inhibitory motor neurons are from the ChAT/NOS interneurons (Li & Furness 2000). Thus, the ChAT/NOS neurons are the ones most likely to be directly involved in local descending inhibitory reflexes, although an involvement of other interneurons may occur. The inhibitory motor neurons also receive direct inputs from calbindinimmunoreactive IPANs (Li & Furness 2000), indicating that there are monosynaptic IPAN to motor neuron reflex pathways, as has also been directly shown for reflexes impinging on excitatory motor neurons (see above). Direct inputs to circular muscle motor neurons, confirming the existence of monosynaptic reflexes, have been demonstrated by physiological experiments in which two neurons were simultaneously impaled with microelectrodes. Stimulation of an impaled IPAN caused excitation in a connected motor neuron (Kunze et al. 1993). Although excitation travels along the intestine behind a bolus, and is reinforced by the presence of the bolus, excitation can also propagate anally without this reinforcement in some circumstances, for example when the stimulus is stationary. Descending excitatory reflexes, which are also elicited by distension, are reduced by about 75% by the 5-HT3 receptor antagonist granisetron (Monro et al. 2002), and descending excitation evoked by electrical stimulation of intramural nerves is also reduced by a blocker of 5-HT3 receptors (Jin et al. 1989), implying that the 5-HT containing neurons are involved in this component of propulsive reflexes. Intrinsic secretomotor and vasomotor circuits Distension, mechanical stimulation of the mucosa, and chemicals applied to the mucosa evoke secretomotor and vasodilator reflexes (Thiry 1864, Herrin & Meek 1933, Babkin 1950, Biber et al. 1971, Fasth et al. 1977, Diener & Rummel 1990, Frieling et al. 1992, Vanner et al. 1993, Sidhu & Cooke 1995, Weber et al. 2001, Vanner & MacNaughton 2004). An important physiological activator is intraluminal glucose (Sjövall et al. 1984a, See & Bass 1993), which triggers secretomotor reflexes that return to the lumen water that was absorbed with nutrients (see Chapter 6). The reflex dilation and secretion both occur in the intestine and most of the colon that is chronically extrinsi-
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Fig. 3.5 The pathways of secretomotor and vasodilator reflexes. Secretomotor and vasodilator
reflexes are initiated when intrinsic primary afferent neurons (IPANs) are activated, generally by a stimulus to their mucosal endings. Reflex secretion can follow through three related circuits. It can occur through a mono-neuronal reflex, in which an axon reflex, or a transsomatic reflex, occurs in an IPAN. It can also occur through synaptic connection from IPANs to the cell bodies of secretomotor and secretomotor/vasodilator motor neurons in submucosal ganglia. These reflexes bypass the myenteric plexus. Third, reflex excitation can pass through the myenteric ganglia and back to secretomotor and secretomotor/vasodilator neurons with cell bodies in the submucosal ganglia. Arrows indicate directions of information flow in the reflex circuits.
cally denervated, and both are blocked by tetrodotoxin (Biber et al. 1971, Fasth et al. 1977, Cassuto et al. 1981a, 1983, Sjöqvist 1991, See & Bass 1993, Cooke et al. 1997a, Weber et al. 2001). Thus these reflexes, similar to the motility reflexes discussed above, are mediated through intrinsic nerve circuits (Fig. 3.5). In the distal part of the colon, adjacent to the rectum, the reflexes are mediated through an extrinsic reflex passing via the pelvic nerves (Fasth et al. 1977). The intrinsic reflexes appear to be predominantly local, that is, responses occur close to the stimuli. This makes it possible to analyze the reflexes in small regions of intestine mounted in Ussing chambers. The distances that secretomotor reflexes are conducted along the intestine have been examined in the rabbit small intestine, in vitro and in vivo (Hubel et al. 1991). These experiments indicated that reflex responses were prominent only within the area of stimulation, and did not travel more than about 4 mm along the intestine. Electrical stimulation of nerves in the submucosal layer, and retrograde tracing experiments, are consistent with reflex pathways being primarily local (Neunlist et al. 1998). Electrical stimulation 4 mm oral or anal to the region
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of recording indicated that stimulation from oral is more effective. Lesion studies indicate that secretomotor neurons have cell bodies in the submucosa and innervate the underlying mucosa (Keast et al. 1984), and studies of submucosal IPANs indicate that they make local connections (Pan & Gershon 2000, Furness et al. 2003a). There are three types of secretomotor neuron in the small intestine; almost all cell bodies of these neurons are in submucosal ganglia in small mammals, and probably also in larger mammals (Chapter 2, Fig. 2.16). Studies in the guinea-pig small intestine indicate that two of the final motor neurons for secretion have collaterals that innervate arterioles (Furness et al. 1987b, Jiang et al. 1993, Evans et al. 1994, Li et al. 1998) and thus they are also motor neurons for vasodilation. Neurons of one group are non-cholinergic, VIP-containing neurons and the others are cholinergic neurons that are also immunoreactive for calretinin (see Table 2.1 for chemical coding). It should be noted that in humans submucosal VIP neurons are also immunoreactive for vesicular acetylcholine transporter (VAChT) and ChAT, and are therefore probably also cholinergic (Schneider et al. 2001, Anlauf et al. 2003). There are probably not separate reflexes for secretion and dilation, although a third type of secretomotor neuron does not project to the vasculature (these are cholinergic neurons with NPY immunoreactivity in the guinea-pig, see Chapter 2). Intraluminal application of toxins – cholera toxin (Cassuto et al. 1981b) or heat-stable enterotoxin (Eklund et al. 1985, Hubel et al. 1991) – and infective agents, including Salmonella (Brunsson 1987), Cryptosporidium (Argenzio et al. 1996), and rotavirus (Lundgren et al. 2000) cause secretomotor reflexes (Fig. 3.6). Cholera toxin-induced reflexes were blocked by tetrodotoxin, applied by local intra-arterial injection in cats (Cassuto et al. 1982a), indicating that the effect is neural. Rotavirus-induced secretion in segments of mouse intestine was also reduced by tetrodotoxin and by the nicotinic receptor antagonist mecamylamine (Lundgren et al. 2000). The secretomotor reflex pathways that are stimulated by cholera toxin pass through the myenteric plexus and the reflex is prevented by ablating this plexus (Jodal et al. 1993). Cholera toxin does not readily cross the intestinal epithelium, and therefore probably acts on epithelial cells that provoke the secretomotor reflexes (Lundgren 2002). It causes 5-HT release from enterochromaffin (EC) cells (Beubler et al. 1989, Racke et al. 1996) and secretion initiated by intraluminal cholera toxin is abolished by the 5-HT3 receptor blocker granisetron (Mourad et al. 1995), implying that enterochromaffin cells carry the signal from the lumen to 5-HT-sensitive nerve endings. Rotavirus-induced secretion is also reduced by granisetron (Kordasti et al. 2004). The release of 5-HT is dependent on the activation of voltage-activated, L-type Ca2+ channels (Lundgren 2002), suggesting that the toxin depolarizes the EC cells. The effect of cholera toxin applied intraluminally was prevented by the nicotinic receptor antagonist
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hexamethonium, confirming the presence of a nicotinic synapse in the reflex pathway (Cassuto et al. 1982a). Secretion induced by 5-HT, intra-arterial, was also abolished by hexamethonium (Cassuto et al. 1982b), indicating that the site of effect of 5-HT is proximal to the nicotinic synapse (Fig. 3.6). Secretion that is induced by toxins is probably mediated primarily by VIP released from non-cholinergic secretomotor/vasodilator neurons. Atropine had little effect on the response to cholera toxin (Cassuto et al. 1982a). However, responses to cholera toxin and to rotavirus were reduced by VIP receptor antagonists (Mourad & Nassar 2000, Kordasti et al. 2004).
Fig. 3.6 Circuitry for secretomotor reflexes that are initiated by pathogens or the toxins that they produce. The majority of experimental data have been obtained from experiments with cholera toxin. The toxins depolarize entero-endocrine cells, causing the opening of L-type Ca2+ channels and the influx of Ca2+ coupled to the release of 5-HT. At the same time, the noxious stimuli provoke prostaglandin (PG) synthesis and release from enterocytes. 5-HT and prostaglandins act on the endings of IPANs to initiate action potentials in these neurons. In this response, the main receptor for 5-HT is the 5-HT3 receptor. The IPANs synapse with interneurons in the myenteric plexus (MP), which project to VIP-immunoreactive secretomotor/vasodilator neurons. These neurons release VIP and PACAP. Secretion is caused by elevation of cyclic AMP and opening of Cl– channels. Prostaglandins also cause secretion by their direct action on the enterocytes.
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Reflexes initiated by mechanical stimulation of the mucosa or by distension can be mediated entirely through the submucosal plexus, because they are recorded in preparations consisting only of the mucosa and submucosa (Frieling et al. 1992, Cooke & Reddix 1994, Weber et al. 2001). Consistent with this, IPANs provide innervation of secretomotor and secretomotor/ vasodilator neurons in the submucosal ganglia (Lomax et al. 2001, Reed & Vanner 2001, Furness et al. 2003a). When submucosal IPANs are activated by physiological stimuli they cause both fast and slow EPSPs in other submucosal neurons (Pan & Gershon 2000), which implies a direct innervation of the secretomotor neurons because other submucosal neurons in the guinea-pig (which are secretomotor or secretomotor/vasodilator neurons) do not innervate the submucosal ganglia. Thus short reflexes from IPANs to secretomotor and secretomotor/vasodilator neurons are monosynaptic. Interneurons probably occur in the submucosal plexus of the pig intestine. Submucosal circuits for both motility and secretion control are present in the submucosal plexus of the pig and other large animals (Timmermans et al. 2001), but it has not yet been ascertained which of the interneurons are in secretomotor pathways. 5-HT is an intermediate in transducing the distortion of the mucosa to the activation IPANs and the generation of secretomotor reflexes (Fasth et al. 1977, Sidhu & Cooke 1995, Kellum et al. 1999). Pathways controlling vasodilation (and by implication, secretion) also pass through the myenteric plexus (Reed & Vanner 2003). Detailed studies of these pathways have been made in isolated preparations in which dilation of mucosal arterioles was used as a measure of output from the reflex pathways and in which it is possible to sever intrinsic nerve pathways (Reed & Vanner 2003). Vasodilation was observed further than 1.5 cm from local stimuli. Robust responses were evoked when the mucosa was stroked, and weaker responses were elicited by stretch of the external muscle after the mucosa had been entirely removed. These data suggest that both submucosal and myenteric IPANs are involved in initiating vasodilator and secretomotor reflexes (Vanner & MacNaughton 2004). Severing the myenteric plexus 5–7 mm from the stimulus site completely blocked the response, whereas severing the submucosal plexus was ineffective (Reed & Vanner 2003). The pathways involve synapses in the myenteric plexus, and from their length they can be polysynaptic. Some at least of the synapses are cholinergic, since the responses are blocked by hexamethonium, and the final vasodilator neurons are also cholinergic, causing vasodilation through muscarinic receptors (Reed & Vanner 2003). The long reflexes that pass via the myenteric plexus appear to impinge primarily on the cholinergic secretomotor/vasodilator neurons (Reed & Vanner 2003). Reflexes induced by mucosal stroking in preparations consisting of submucosa plus mucosa are neurally mediated (blocked by tetrodotoxin), but are
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not blocked by an antagonist of cholinergic fast neuro-neuronal transmission, mecamylamine (Frieling et al. 1992, Sidhu & Cooke 1995). The concentration of mecamylamine that was used was shown to block nicotinic receptors in the colon (Sidhu & Cooke 1995). Moreover, the responses to stroking were not reduced by extrinsic denervation, indicating that they are dependent on activation of intrinsic neurons (Cooke et al. 1997a). There are two mechanisms that can explain these observations, the presence of non-nicotinic excitatory transmission (Hu et al. 2003), and the possibility that IPANs can mediate effects non-synaptically. These are not mutually exclusive explanations, and both mechanisms may occur. In both rat and guinea-pig distal colon, reflexes that were elicited by mucosal stroking were reduced by blocking purine P2Y receptors (Christofi et al. 2004, Cooke et al. 2004). Pharmacological and localization studies indicated that the receptors are on submucosal secretomotor neurons, and excitatory synaptic events blocked by purine receptor antagonists occur in submucosal neurons (Hu et al. 2003, Monro et al. 2004). Because the local reflexes are likely to be mediated through IPAN to secretomotor neuron connections (see above), this implies a purinergic transmission from the IPANs. There is also evidence that calcitonin gene-related peptide (CGRP) could contribute to non-cholinergic transmission from submucosal IPANs to secretomotor neurons (Pan & Gershon 2000). In addition, IPANs may themselves release neurotransmitters that cause secretion (Fig. 3.5). IPANs are immunoreactive for tachykinins and their varicose processes are immunoreactive for the vesicular acetylcholine transporter, VAChT (Li & Furness 1998). Thus their mucosal endings have the potential to release both acetylcholine and tachykinins, each of which causes secretion. It has been directly shown that action potentials in one process of an IPAN traverse the cell body to invade other processes (Chapter 2) and the pattern of branching of the neurons indicates that action potentials could be conducted, as an axon reflex, between terminals that branch within the mucosa (Fig. 3.5). The responses to mucosal stroking in the guinea-pig colon were partly reduced by blocking NK1 receptors for tachykinins (Cooke et al. 1997a), which are present in IPANs but not in secretomotor neurons. Thus there is evidence that acetylcholine and tachykinins that are released by axon reflexes, or by mononeuronal reflexes crossing the cell bodies of IPANs, contribute to secretory responses (Fig. 3.5). Assemblies of neurons The densities of occurrence of enteric neurons are high in relation to the sizes of regions that are exposed to physiological stimuli, such as distension or changed nutrient concentrations in the intraluminal fluid. From counts made in the guinea-pig small intestine, it is estimated that there are around 2500
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nerve cells in the myenteric ganglia per mm length of unstretched gut (Kunze & Furness 1999). Among these, the total number of myenteric IPANs is about 650 per mm length. IPANs are about 11% of submucosal neurons, and the ratio of submucosal to myenteric neurons is 0.35 : 1 (Gabella 1987). Thus there are about 1100 submucosal neurons, including about 100 submucosal IPANs, per mm length. A 1 mm length also contains the cell bodies of 400 inhibitory motor neurons, 300 excitatory motor neurons to the circular muscle and about 1000 secretomotor neurons, as well as the cell bodies of about 120 ascending and 120 ChAT/NOS descending interneurons (from data in Table 2.1). A large proportion of the remaining myenteric neurons are longitudinal muscle motor neurons, which account for about 500 nerve cell bodies per mm. There are also small populations of ChAT/SOM (4%) and ChAT/5-HT (2%) descending interneurons. IPANS are activated in groups All IPANs project to the mucosa in guinea-pigs (Song et al. 1991, 1994), and probably in other species (Chapter 2). As mentioned, 1 mm length of guineapig small intestine contains about 650 myenteric and 100 submucosal IPANs, most of which are chemo- and mechanosensitive. Because the stimuli giving rise to intestinal reflexes (luminal chemicals, distension, mucosal distortion) are not spatially confined to sub-millimeter distances, it can be deduced that reflexes are usually initiated by the activation of a large population of primary afferent neurons. Moreover, many IPANs, whether they are directly activated or not, are excited synaptically, through the synaptic connections that IPANs make with each other (Chapter 2). IPANs have considerable overlap in their receptive fields; retrograde tracing indicates that each villus is supplied by the axons of about 65 IPANs with cell bodies in myenteric ganglia (Song et al. 1994). By mapping regions of mucosa from which IPANs can be electrically activated, it was concluded that each projects to a strip of mucosa, about 2 mm in oral to anal width and 7 mm in circumferential length (Bertrand et al. 1998), which contains about 80–120 villi. It is thus concluded that assemblies of several thousand myenteric IPANs respond together when changes in muscle tension and luminal chemistry occur. Mucosal distortion also contributes to activation of submucosal IPANs. The information that is gathered about the state of the intestine must be integrated by the IPANs to provide an appropriate output, but no detailed understanding of how this is achieved has yet been reached. Computer modeling of the neural control of the intestine suggests that the assemblies of IPANs act as low-gain amplifiers, and that the co-ordination of firing of
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the neurons may be necessary to generate the migrating myoelectric complex (Thomas et al. 2004). Assemblies of interneurons Activity is conducted orally and anally along the intestine via interneurons. In the small intestine of the guinea-pig there is one class of ascending interneuron and three classes of descending interneurons (Chapter 2). In addition, a small proportion of IPANs have long anally directed processes. A very high proportion, over 90%, of S neurons (interneurons and motor neurons) respond when the intestine is distended (Hirst et al. 1975) and around 50% respond to mucosal distortion (Bornstein et al. 1991a). It thus appears that large numbers of interneurons are activated by a single physiological stimulus. Assemblies of motor neurons Both physiological and structural studies indicate that the circular muscle of the intestine is innervated by dense arrays of nerve fibers that run circumferentially, more or less parallel to the long axes of the circular muscle cells (Chapter 2, Fig. 2.14). These axons influence the muscle cells as a group, because the cells are electrically coupled to each other. The motor unit of the intestinal circular muscle could thus be regarded as a circumferential strip that is electrically closely coupled, and its innervating neurons. In the guinea-pig small intestine, the minimum widths of contractile rings are about 2–3 mm (Schulze Delrieu et al. 1991). Thus, an annulus about 2–3 mm wide, innervated by axons arising from about 600–900 excitatory and 800–1200 inhibitory motor neurons, could be regarded as a motor unit. The numbers of axons supplying the circular muscle have been counted per muscle cell and per mm length (oral to anal). In the deep muscular plexus, there are 2500 axons per mm length of intestine (Wilson et al. 1987), which represents two-thirds of the fibers that innervate the muscle (Gabella 1972b). Thus there are about 3700 axons innervating 1 mm of intestinal length, corresponding to 7500–11 000 axons in a motor unit 2–3 mm wide. Quantitative electron microscopic analysis indicates that about half the axons are from excitatory neurons, and about half are from inhibitory neurons (Llewellyn Smith et al. 1988). These quantitative data indicate that the processes of individual neurons branch within the circular muscle, which has been confirmed by direct observation of individual neurons that were filled with a marker dye (Nurgali et al. 2004). The ratios of axons detected in single cross-sections of the muscle to corresponding nerve cells in the same length is about 12 for the excitatory neurons and 9 for the inhibitory neurons. The majority of excitatory motor neurons project very small distances along the length of the intestine; about 80% of the cell bod-
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ies of neurons supplying a 2 mm band of intestine are located adjacent to the muscle strip, and the remainder are found up to about 8 mm anally (Brookes et al. 1991b). The axons of inhibitory neurons project anally, so the cell bodies providing a muscle strip lie oral to the strip, about 40% within 2 mm and the remainder spread out for up to 14 mm oral to the strip (Brookes et al. 1991b). In view of the large numbers of axons that innervate a neuromuscular unit, it is not surprising that it is difficult to fractionate, by varying the stimulus strength, the electrical events (junction potentials) in the muscle caused by nerve stimulation (Bornstein et al. 1986). The idea of an annulus as a motor unit is consistent with the observation that the muscle undergoes annular (although sometimes eccentric) contractions and relaxations (Cannon 1902, Bass et al. 1961). The intensity of the contraction or relaxation of the muscle depends on the numbers of neurons that are activated, and the frequency with which they fire. Although a strip 2–3 mm wide might be regarded as the smallest motor unit of the circular muscle of the guinea-pig small intestine, the annular contractions of the small intestine usually occupy 10–40 mm (Hukuhara & Neya 1968, Schulze Delrieu et al. 1991, Waterman et al. 1994, Benard et al. 1997, Ferens et al. 2005). This length is innervated by several thousand motor neurons. The muscle motor neurons are not assemblies in the same sense as assemblies of IPANs and interneurons that are connected to each other synaptically. However, as their activity is determined by transmission from the IPANs and interneurons, they are recruited as populations, and because they innervate electrically coupled smooth muscle, their summed, rather than individual, effects are manifested in muscle contraction and relaxation. Circuits in the esophagus and stomach At the end of Chapter 1 the gross structural similarities between the myenteric plexus of the esophagus, stomach, and intestine are referred to and it was speculated that the similarity is a consequence of the common evolutionary origins of the plexus. However, when we come to consider the functions of the myenteric plexus we are faced with a number of contrasts. Esophagus The myenteric ganglia have essentially the same architecture as myenteric ganglia elsewhere in the digestive tract (Morikawa & Komuro 1999), but the circuitry of this part of the enteric nervous system has not been investigated in any significant detail. The control of motility of the esophagus is discussed in Chapter 5. In essence, the movements of that part of the esophagus where the external
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muscle is striated are controlled by a motor program that is generated in the brainstem and co-ordinates swallowing and esophageal movement. Thus the enteric neurons, although they are prominent in this region, have little role in controlling peristalsis. Variable lengths of the distal part of the esophagus have a smooth muscle wall, and in this region propulsive reflexes can be initiated through the enteric nervous system. The circuitry is probably similar to that in the small intestine (see above), although this has not been adequately investigated. Some of the motor endplates of the striated muscle of the esophagus receive an innervation from enteric neurons as well as from the vagal motor neurons. There is evidence that this enteric innervation inhibits transmission from the vagal fibers, and it may be involved in co-ordinating movement of the striated and smooth muscle regions (Chapter 5). Likewise, vagal innervation of enteric neurons that supply the smooth muscle of the distal esophagus may be involved in co-ordination of the two regions. Myenteric neurons in the esophagus are almost certainly the source of innervation of its muscularis mucosae. Stomach The stomach has two main motility-dependent functions, first to adjust its volume to accommodate changes in the amount of gastric contents and second to push the contents against the pyloric sphincter, which acts to mix the food and to propel small liquid aspirates into the duodenum. The adjustment in gastric volume is mainly achieved by two reflexes, the reflex of receptive relaxation and the accommodation reflex (Chapter 5). Both reflexes are mediated through the vagus and involve control centers within the central nervous system, although there is some evidence for an accommodation reflex through intrinsic reflex circuits (Chapter 5). Vagotomy impairs the reservoir function of the stomach (Roman 1982). The rhythmic movements that propel the contents of the stomach towards the pylorus (gastric peristalsis) seem to be generated in the smooth muscle, whose electrical activity is in turn determined by the interstitial cells of Cajal (Chapter 5). The contractions coincide with waves of depolarization (slow waves) in the muscle; they begin at the proximal part of the corpus and propagate across the antrum to the pylorus. The slow waves are observed whether or not the animal is anesthetized, and also occur in vitro (Kelly et al. 1969). When strips of gastric muscle from the dog or human stomach are examined in vitro, slow waves of sufficient amplitude to generate action potentials and muscle contraction are seen even when the activities of enteric neurons are blocked by tetrodotoxin (El Sharkawy et al. 1978). The intensity of the rhythmic gastric movements is modulated by enteric neurons. In contrast to the intestine (see above), the progress of gastric peristalsis is not dependent
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on the continuity of the myenteric plexus. Thus, cuts through the myenteric plexus do not affect gastric peristalsis (Cannon 1912). The gastric peristaltic activity also persists after combined vagotomy and splanchnic nerve section (Auer 1910). In fact, its frequency (5/min in dogs) was unaffected by complete isolation of the stomach from extrinsic neural influence (Van Lier Ribbink et al. 1989). Moreover, Cannon found that nicotine, which almost completely blocks fast excitatory transmission between enteric neurons, does not block peristalsis in the stomach, whereas it is blocked in the small intestine (Cannon 1911). These data indicate that propulsive waves in the stomach are not dependent on reflexes within the enteric nervous system of the stomach or on extrinsic innervation for their generation. However, regulation of the intensity of the contractions that are triggered by gastric slow waves occurs partly through vagal pathways and partly through local enteric reflexes. Thus, the strengths of the contractions are clearly influenced by enteric neurons, and can also be enhanced by compounds that excite gastric muscle (Armitage & Dean 1966). Enteric motility reflexes do occur in the stomach (Yuan et al. 1997), but they do not have the prominence of intrinsic reflexes of the small and large intestines (Chapter 5). Despite the major regulation of gastric motility being through extrinsic nerves, it is unlikely that the myenteric ganglia of the stomach are simply relays between extrinsic neurons and the muscle, as there is a substantial innervation of the ganglia from intrinsic sources (Grundy & Schemann 2002). Co-ordination of motility, secretomotor, and vasomotor reflexes In the small intestine and colon the same types of reflex stimuli trigger motility and secretomotor reflexes. Thus, it is not surprising that there is a degree of synchrony between changes in motility and changes in secretion (Read et al. 1977, Greenwood & Davison 1987, Mellander et al. 2000). Increases in secretion and blood flow are linked through the activation of final neurons with dual secretomotor and vasodilator functions, which is a physiological necessity because the water and electrolyte that is secreted derives in part from the blood supply (Chapter 6). Secretion into the gut lumen is also coordinated with increased motility during the late part of phase II of the migrating myoelectric complex (Mellander et al. 1993). Circuits connecting the intestine, biliary system, and pancreas Neuronal tracing experiments show that neurons in the gastrointestinal tract project to the sphincter of Oddi, the gall bladder and the pancreas. Retrograde tracer injected into either the gall bladder or the sphincter of Oddi
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labels neurons in the duodenum (Padbury et al. 1993, Kennedy & Mawe 1998). There is a reciprocal innervation, as sphincter of Oddi neurons are labeled from the duodenum (Mawe & Kennedy 1999). Retrograde tracer injected into the pancreas labels nerve cells in the gastric antrum and duodenum (Kirchgessner & Gershon 1990). The presence of a reflex connection was demonstrated by recording fast EPSPs in pancreatic neurons in response to duodenal distension (Kirchgessner et al. 1996). Currently there is little indication of the significance of direct neural connections between the digestive canal and the gall bladder or pancreas. Sympathetic innervation of the gastrointestinal tract The sympathetic and enteric innervations of the gastrointestinal tract have considerable interaction, and an understanding of the sympathetic innervation is thus pertinent to an understanding of enteric control. The organization of the sympathetic pathways and the ways in which the endings of sympathetic post-ganglionic neurons are related to the enteric ganglia and other intramural targets are well established. Moreover, the original discoveries that have framed our understanding have been thoroughly reviewed (Furness & Costa 1974, 1987, Szurszewski 1981, Szurszewski & Miller 1994). Because of this, I will summarize most of the essential information without quoting all the original literature sources, which can be found in the reviews cited above. The post-ganglionic sympathetic neurons of mammals contain norepinephrine, which is their primary transmitter. Norepinephrine can be directly converted to a fluorescent product by reaction with aldehydes, so it is very easy to determine the pattern of innervation of the gastrointestinal tract by norepinephrine axons. The neurons can also be visualized by the immunohistochemical localization of the norepinephrine-synthesizing enzymes tyrosine hydroxylase or dopamine β-hydroxylase. The sympathetic, noradrenergic nerve endings provide their most prominent innervation to the enteric ganglia. Seemingly, the cell body of every enteric neuron has numerous varicose noradrenergic axonal endings close by. The other prominent innervation is of arterioles in the gut wall and of the circular muscle layer of the sphincter regions. Few noradrenergic fibers supply the muscle layers of non-sphincter regions, although there is a definite innervation, which is more prominent in larger mammals (e.g. dogs, humans), than in small mammals (guinea-pigs, rats). Even in the larger mammals the innervation of the muscle is extremely sparse compared to the innervation by enteric motor neurons. Some noradrenergic fibers are found in the mucosa of the stomach, intestines, and gall bladder, but again, very few compared to the innervation supplied by the intrinsic enteric neurons.
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The cell bodies of the sympathetic neurons that supply the gastrointestinal tract are in sympathetic chain (paravertebral) ganglia and in the large prevertebral ganglia of the abdominal and pelvic cavities (the celiac, superior mesenteric, inferior mesenteric and pelvic ganglia). The cell bodies in paravertebral ganglia are mainly those involved in controlling the gastrointestinal blood vessels (vasoconstrictor neurons), whereas there are three classes of neurons in prevertebral ganglia (Fig. 3.7): vasoconstrictor neurons, neurons controlling motility, and neurons controlling secretion (neurons of this last
Fig. 3.7 The sympathetic innervation of the intestine. The three main types of sympathetic neurons that supply non-sphincter parts of the intestine are: 1, vasocontrictor neurons, 2, secretion-inhibiting neurons, and 3, motility-inhibiting neurons. These neurons all receive synaptic inputs from the spinal cord. In addition, the secretion-inhibiting and motility-inhibiting neurons receive inputs from intestinofugal neurons. For further details, see text.
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type also affect blood flow). Although indirect evidence had suggested that the three neuron types, controlling the three functions, were separate (Furness & Costa 1974), this was made much more obvious when it was discovered that the three classes of neuron had different chemical coding. Most of the axons of the noradrenergic neurons reach the gut by following mesenteric nerves, but a few axons of neurons with cell bodies in paravertebral ganglia of the upper thorax follow the vagus nerve. The vasoconstrictor neurons innervate the arteries directly and cause constriction through the release of norepinephrine, that is probably augmented by co-release of ATP, which also causes constriction. The noradrenergic vasoconstrictor neurons also contain NPY, which is released on nerve stimulation and which might make a small contribution to the constriction of the vessels. Noradrenergic neurons that influence motility do so in two ways, by inhibiting activation of enteric neurons and by constricting the sphincters (Chapter 5). Both actions reduce the transit of intestinal content along the digestive tract. The observation that actions of sympathetic neurons inhibit the gut dates from at least as long ago as the mid-19th century, when Pflüger (1857) described the reduction of peristalsis in the rabbit small intestine when the splanchnic nerves were stimulated. Pflüger’s results were confirmed in the following year by Lister (1858). Elegant physiological experiments published between 1965 and 1970 showed that the effect was caused by actions on enteric ganglia, not on the muscle, thus linking the physiological site of action to the histochemically determined distribution of noradrenergic nerve endings in the gut (see Chapter 5). The inhibition of secretion of water and electrolyte by sympathetic neurons has great importance for fluid homeostasis. It is mediated principally through post-ganglionic neurons that inhibit the excitability of VIP-containing secretomotor/vasodilator neurons (Chapter 6). Summary and conclusions The discovery of intrinsic ganglia, and observation of the behavior of the digestive system, led 19th-century scientists to surmise that there are intrinsic reflex circuits controlling movement and fluid secretion in the gastrointestinal tract. These circuits have gradually been unravelled. Motility-controlling circuits in the small and large intestines include IPANs with cell bodies in the submucosal and myenteric plexuses that react to appropriate physiological stimuli. They connect with interneurons that carry the reflex signals for short distances along the intestine, through the myenteric plexus. IPANs also connect directly with motor neurons. Large numbers of IPANs, interneurons, and motor neurons are activated at the same time. The circuits generate a number
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of patterns of movement of the gut wall, including propulsive and mixing movements, which are discussed in Chapter 5. Local stimuli initiate polarized reflexes that cause contractions oral to a point of reflex initiation, and relaxation anal. These simple, stereotyped reflexes have been analyzed in detail to deduce the nerve circuitry. Secretomotor and vasomotor reflex circuits also consist of IPANs, interneurons, and motor neurons. In some cases they pass through only the submucosal plexus, in others through the myenteric plexus. Although there are intrinsic neuronal circuits in the esophagus, where the muscle cells of its wall are striated, and in the stomach, reflex control of these regions is primarily through centers in the brainstem. Sympathetic neurons inhibit muscle movement and secretion of fluid indirectly, by actions on the enteric circuits, but cause vasoconstriction through their direct innervation of blood vessels in the gut wall.
4: Pharmacology of transmission and sites of drug action in the enteric nervous system
Chemical coding and multiple transmitters Extensive immunohistochemical studies demonstrate that each enteric neuron contains several substances whose chemistry and receptor pharmacology are consistent with the compounds being neurotransmitters. Moreover, pharmacological analysis of transmission has revealed numerous instances in which enteric neurons release more than one transmitter. Despite this, the roles of many compounds that are contained in enteric neurons and might be supposed to be transmitters are not known. Immunohistochemical investigation of neuropeptides and neurotransmitter-synthesizing enzymes in the enteric nervous system led to the chemical coding hypothesis, which states that each class of neuron that can be differentiated functionally contains a unique combination of chemical markers (Gibbins et al. 1987, Furness et al. 1987a, 1989a). Chemical codes of neurons in the guinea-pig small intestine are included in Table 2.1 (page 32). Knowledge of some of the coding is quite extensive, an example being the cholinergic secretomotor neurons that are immunoreactive for choline acetyltransferase (ChAT), an essential constituent of cholinergic neurons, and also for six peptides whose roles in these neurons are obscure, these being calcitonin generelated peptide, cholecystokinin, dynorphin, galanin, neuropeptide Y and somatostatin (Furness et al. 1984, Steele & Costa 1990). Of the potential transmitters in enteric neurons, several in each neuron appear to be neurotransmitters, such as ACh and tachykinins in excitatory motor neurons to the muscle and nitric oxide (NO), VIP and ATP in motor neurons that inhibit the muscle. Each is discussed in more detail below. Primary transmitters appear to be conserved across mammalian species. Thus, when a substance is a primary neurotransmitter, it appears to be present in all neurons that have equivalent roles in different species and regions of the gastrointestinal tract. For example, extensive studies in many species indicate that all inhibitory neurons to the muscle contain the synthesizing enzyme for NO, nitric oxide synthase (NOS), and the peptide VIP. However, other 103
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peptides that form the chemical code of these neurons vary. In the guinea-pig small intestine, inhibitory neurons with short projection have the code NOS/ VIP/ENK/GABA/NPY/PACAP, whereas the long-projecting neurons have the code NOS/VIP/BN/DYN/NFP/PACAP (Uemura et al. 1995, Williamson et al. 1996). Some of the long neurons are also NPY immunoreactive. Also in humans a proportion of the VIP-containing neurons that innervate the circular muscle are NPY immunoreactive (Wattchow et al. 1988). No differences in the pharmacology of transmission that relate to these chemical coding differences are known. Another example of species difference is that the noncholinergic secretomotor neurons in guinea-pig do not contain NPY (Table 2.1), whereas they do in the rat (Ekblad et al. 1984). The biological significance of there being substances (most of which are small peptides) with no obvious function in neurons has been discussed elsewhere (Furness et al. 1989a, Bowers 1994, Brookes & Costa 2002). It has been postulated that some substances may have no role (they could be considered developmental or evolutionary leftovers) or that some substances may have roles other than participation in neurotransmission. Transmitters of motor neurons that innervate the smooth muscle of the gut Both smooth muscle cells and the interstitial cells of Cajal (ICC) are innervated by muscle motor neurons, and a significant component of transmission to the muscle is indirect, through control by the motor axons of the excitability of ICC that are connected through electrically conducting gap junctions to the muscle (see below and Chapter 5). Excitatory neurons: acetylcholine The transmitters of excitatory motor neurons and their sites of action are summarized in Fig. 4.1. That ACh is the primary transmitter of excitatory neurons innervating the muscle is unquestioned, so this will be reviewed only briefly. This conclusion is based in the first instance on the observation that excitatory transmission to the muscle is almost fully prevented by broadspectrum antagonists of muscarinic receptors for ACh, such as atropine, and is facilitated by inhibition of acetylcholine esterase (AChE), the ACh metabolizing enzyme (Harrison & McSwiney 1936, Ambache & Edwards 1951, Paton 1955, Paton & Zar 1968). Atropine, or other antagonists of acetylcholine receptors on the muscle, block the contraction of the muscle that occurs during peristaltic reflexes (Trendelenburg 1917, Fleisch & von Wyss 1923, Raiford & Mulinos 1934). Muscarinic antagonists also depress motility of the human intestine (Fink 1959). There are reports of resistance
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Fig. 4.1 Transmission from motor neurons to gastrointestinal smooth muscle. Excitatory motor neurons (A) and inhibitory motor neurons (B) both release transmitter that acts on interstitial cells of Cajal (ICC). The consequent events in the ICC are electrically coupled to smooth muscle cells through gap junctions. Transmitter can also diffuse to and act directly on the muscle cells. A: Excitatory transmission. Excitatory motor neurons release acetylcholine and tachykinins as their primary transmitters. These act through muscarinic and NK1 receptors to excite ICC and through muscarinic and NK2 receptors on the muscle. The neurons also release opioid peptides and GABA that provide a negative feedback inhibition of transmitter release. B: Inhibitory transmission. Inhibitory motor neurons release nitric oxide (NO), ATP, VIP and PACAP, all of which have inhibitory actions on the ICC and muscle cells. The receptor for NO is soluble guanylyl cyclase in the cytoplasm of the innervated cells. ATP acts through P2X and P2Y receptors on ICC and P2Y receptors on the muscle, and VIP and PACAP act through VPAC receptors.
of transmission to block by muscarinic antagonists in vivo, but the lack of inhibition in these cases is primarily due to the metabolism of the muscarinic antagonists by atropinases, or through other mechanisms that remove the antagonists (Ambache 1955, Bogeski et al. 2005). Genuinely resistant transmission, mediated by tachykinins, which are co-transmitters with ACh, is discussed below.
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Release of ACh from neurons innervating the gastrointestinal tract was first detected by Dale and Feldberg (1934), work which contributed to the award of the Nobel Prize that Dale shared with Otto Loewi for the discovery of chemical transmission. They showed that vagal stimulation caused the release of a substance, which mimicked ACh in all respects, into the vascular perfusate of the stomach. ACh release was confirmed by numerous other workers and was shown to be blocked or substantially reduced by tetrodotoxin, botulinum toxin or removal of Ca2+ ions (Furness & Costa 1987), which confirms its neural origin. The presence of the ACh synthesizing enzyme, ChAT, in the intestine was first demonstrated biochemically (Dikshit 1938). Later, when antisera to the enzyme became available, immunohistochemical studies showed that ChAT was located in neurons (Furness et al. 1983), including the excitatory muscle motor neurons (Brookes et al. 1991b). Of the five muscarinic receptor types, two, the M2 and M3 receptors, are the predominant muscarinic receptors of gastrointestinal muscle (Lecci et al. 2002). Activation of M2 receptors is linked to a G-protein coupled inhibition of adenylyl cyclase activity, which reduces the effects of this enzyme in inhibiting muscle contraction, and to opening of non-selective cation channels that cause muscle depolarization and Ca2+ entry. M3 receptor-mediated effects occur through activation of phospholipase C and consequent release of Ca2+ from intracellular stores and protein kinase C activation. Co-transmitters of excitatory motor neurons Immunohistochemical studies and quantitative electron microscopy reveal that ACh and tachykinins are contained in the same excitatory muscle motor neurons, and that there is probably only one type of excitatory neuron, which releases both transmitters (Llewellyn Smith et al. 1988, Brookes et al. 1991b). Consistent with this observation, pharmacological analysis of transmission shows that tachykinins are also excitatory transmitters, although they have a lesser role than ACh (Holzer & Holzer Petsche 1997). The first tachykinin to be identified was substance P, but it is now known that several closely related peptides, substance P and neurokinin A (and possibly neuropeptide γ and neuropeptide K) among them, are produced and probably released together from enteric neurons. Therefore the transmitter is referred to as a tachykinin rather than substance P, which was identified as the only tachykinin transmitter known before about 1985. A substance that lowered blood pressure, later identified as substance P, was originally extracted from intestine and brain (Von Euler & Gaddum 1931), which indicates its abundance in these tissues. It was sequenced from brain (Chang et al. 1971) and, soon after, the same peptide sequence was determined for substance P extracted from the intestine (Studer et al. 1973). A few years later,
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immunohistochemical studies showed that tachykinin immunoreactivity in the intestine was confined to neurons, including neurons that innervate the muscle layers (Pearse & Polak 1975, Schultzberg et al. 1978). There is direct experimental evidence for the involvement of both substance P and neurokinin A in excitatory neuromuscular transmission (Grider 1989b, Lippi et al. 1998). The tachykinin component of excitatory transmission appears to be more prominent, relative to the cholinergic component, at high rates of firing of the neurons (Franco et al. 1979). In the mid 1970s, when substance P was first found in nerve fibers innervating the muscle, good receptor antagonists were not available to test its role in transmission. However, it was well known that some substances, including substance P, desensitize their own receptors, and tachykinin receptor desensitization was shown to abolish the non-cholinergic component of transmission, providing evidence for an excitatory transmitter role of tachykinins (Franco et al. 1979). Soon after, better antagonists and improved methods of assay of tachykinins were developed and the role of tachykinins as co-transmitters of the excitatory neurons was established (Barthó & Holzer 1985, Holzer & Holzer Petsche 1997). The initial experiments used electrical stimulation to elicit responses. However, it was also demonstrated that transmission during reflexes elicited by physiological stimuli had a tachykinin component (Costa et al. 1985). Block of the tachykinin receptors while cholinergic transmission is intact usually causes only a small reduction of excitatory transmission (Barthó & Holzer 1985, Zagorodnyuk et al. 1993), which reinforces the idea that the primary transmitter is ACh. When the strength of the stimulus is increased, especially in the presence of a muscarininc receptor blocker, effective excitatory reflexes can be elicited. These are antagonized by tachykinin receptor blockers (Grider 1989b, Holzer 1989, Giuliani et al. 1993). Two receptors for tachykinins, NK1 and NK2 receptors, mediate the excitatory transmission elicited by electrical or reflex activation of excitatory motor neurons (Maggi et al. 1994, Holzer & Holzer Petsche 1997, Holzer et al. 1998, Lecci et al. 2002). NK1 receptors are located primarily on the ICC and NK2 receptors are prominent on the muscle, although there are also NK1 receptors in the muscle (Sternini et al. 1995, Grady et al. 1996, Portbury et al. 1996a,b, Southwell & Furness 2001). The differences in receptor localization may explain differences in the pharmacology of transmission, brief trains of pulses giving responses that are mainly through NK1 receptors, and longer trains of stimuli eliciting responses through both receptor types (Maggi 2000). The hypothesis underlying this explanation is that the ICC are directly innervated, whereas transmitter needs to diffuse greater distances to affect the muscle. Although ACh and tachykinins are the primary transmitters of the excitatory motor neurons, there may be a contribution to transmission from ATP.
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Depolarizing potential changes (excitatory junction potentials; EJPs) can be elicited in intestinal muscle by stimulation of motor neurons after cholinergic transmission has been blocked by atropine (Zagorodnyuk et al. 1995, Zagorodnyuk & Maggi 1998). The EJPs had a fast and a slow component. The slow component was blocked by a combination of NK1 and NK2 tachykinin receptor antagonists, and the fast component by block of purine receptors (receptors for ATP) with suramin or PPADS. Transmitters of excitatory motor neurons that modulate release of the primary transmitters Opioid peptides and γ-aminobutyric acid (GABA), both of which are found in the excitatory motor neurons, inhibit release of the excitatory transmitters. There is also evidence that ACh may have a feedback effect to inhibit excitatory transmitter release. Opioid peptides (enkephalin and enkephalin-related peptides) are contained in the motor neurons that innervate the muscle of the gastrointestinal tract (Elde et al. 1976, Schultzberg et al. 1980). However, activation of opioid receptors, for example with morphine, has no direct effect on the muscle, but reduces the output of ACh from the motor neurons (Paton 1957, Schaumann 1957, North & Tonini 1977). Opioid receptor antagonists increase the output of ACh when the motor neurons are stimulated, indicating that the release of endogenous opioid peptides from the neurons inhibits release (Waterfield & Kosterlitz 1977). Consistent with this observation, if enteric excitatory reflexes are depressed, for example by atropine or hexamethonium, the depression can be partly reversed by opioid receptor antagonists (Holzer et al. 1998). Moreover, excitatory responses to the stimulation of ascending pathways are enhanced by opioid receptor antagonists (Allescher et al. 2000). The principal receptor type involved in depression of transmitter release is the μ-opioid receptor, with some involvement of κ receptors (Shahbazian et al. 2002). GABA is also present in the motor neurons, although it is in a higher proportion of inhibitory compared to excitatory neurons (Hills et al. 1987, Furness et al. 1989b, Williamson et al. 1996). Like the opioid peptides, GABA has no direct effect on intestinal muscle (Krantis et al. 1980), but it acts through prejunctional GABAB receptors to inhibit the release of ACh (Klienrok & Kilbinger 1983, Ong & Kerr 1983, Minocha & Galligan 1993). The release of acetylcholine in response to electrical stimulation of preparations of longitudinal muscle and myenteric plexus is enhanced by blocking muscarinic receptors and is diminished by muscarinic receptor agonists or by enhancing the availability of endogenous ACh by blocking its metabolism by choline esterases (Kilbinger & Wagner 1975, Somogyi & Vizi 1988). The
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modulation of release could be due to effects on ACh release from cholinergic motor neurons and on ACh release at neuro-neuronal synapses. At least part of the modulation is of release from the motor neurons. This is shown by experiments in which the excitation of the muscle in response to electrical stimulation in the presence of the nicotinic receptor antagonist hexamethonium was investigated (Fosbraey & Johnson 1980). Under these circumstances, nicotinic cholinergic transmission to the excitatory muscle motor neurons is eliminated, and the effects of drugs should be on neuromuscular transmission. With nicotinic receptors blocked, ACh reduced the amplitude of the excitatory response to nerve stimulation without desensitizing the muscle, implying that presynaptic muscarinic receptors reduce ACh release. Transmitters of inhibitory motor neurons In contrast to the identification of the primary transmitter of the excitatory neurons as ACh in the first half of the 20th century, there has been considerable difficulty and substantial debate associated with the identification of the transmitter of inhibitory neurons (Campbell 1970, Furness & Costa 1973a, Makhlouf & Grider 1993, Furness et al. 1995b). It is now recognized that these neurons utilize co-transmitters, including adenosine triphosphate (ATP), nitric oxide (NO) and peptides of the vasoactive intestinal peptide (VIP) family (Fig. 4.1B). During the 1950s and early 1960s it was generally believed that there were two transmitters of peripheral autonomic neurons, acetylcholine and norepinephrine, and that if the excitatory transmitter was acetylcholine the inhibitory transmitter should be norepinephrine. It was also thought that each neuron has a single transmitter. However, soon after compounds that block noradrenergic transmission became available, it was found that they failed to have any significant effect on inhibitory transmission to the muscle of the gut (Burnstock et al. 1964, 1966, Martinson 1965, Campbell 1966). Because the transmission could not be accounted for by either of the known peripheral transmitters it came to be called non-adrenergic, noncholinergic (NANC) transmission. After a short time, evidence was gathered that one transmitter of these neurons was ATP (Burnstock et al. 1970, Burnstock 1972). The major argument in favor of ATP was that it mimicked the effects of nerve stimulation (Burnstock et al. 1972) and that metabolites of ATP were released from the tissue when intramural nerves were stimulated (Rutherford & Burnstock 1978, McConalogue et al. 1996). However, the lack of effective and specific antagonists for muscle receptors for ATP made it difficult to prove the theory (Furness & Costa 1973a). Suramin, an antagonist of the action of ATP on P2 purinoceptors in intestinal muscle, depresses transmission from the inhibitory neurons of the guinea-pig taenia coli (Hoyle et al. 1990), but this
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compound is not very selective (Voogd et al. 1993). The receptors for ATP on the muscle are P2Y purine receptors (Burnstock & Kennedy 1985). High concentrations of PPADs, which block P2Y receptors, antagonize inhibitory transmission (Bian et al. 2000). However, transmission from the neurons is also blocked by the P2X antagonist α,β-methylene ATP (Crist et al. 1992). This may be because the inhibitory transmission to the muscle was via the ICC (see below), which express P2X receptors (Burnstock & Lavin 2002). The effect of ATP on the muscle is mediated by the opening of apamin-sensitive K+ channels (SK channels), and apamin is in many cases an effective antagonist of the ATP-mediated component of inhibitory transmission (MacKenzie & Burnstock 1980, Costa et al. 1986). Apamin does not block inhibitory effects of VIP on the muscle (MacKenzie & Burnstock 1980), and generally does not antagonize the effect of NO, or transmission that is mediated through NO (Rand 1992, Sanders & Ward 1992), a selectivity that augmented the evidence for ATP being a transmitter. Within a few years, the rival theory emerged that VIP was the transmitter (Fahrenkrug 1979). This peptide had been detected immunohistochemically in nerve fibers innervating the muscle. Axons are numerous in both the nonsphincter and sphincter regions throughout the gut, and denervation studies showed that the fibers were of intrinsic origin (Bryant et al. 1976, Larsson et al. 1976). Subsequent studies of the projections and chemical coding of enteric neurons identified VIP as a constituent of inhibitory motor neurons (Chapter 2). VIP was also found to be a potent relaxant of gut muscle (Eklund et al. 1979, Bitar & Makhlouf 1982). The additional evidence that VIP was released when pathways impinging on enteric inhibitory motor neurons were stimulated (Fahrenkrug et al. 1978) strengthened the theory. A contribution of VIP to inhibitory transmission has been demonstrated in some gastrointestinal muscles, but in muscle in other regions it makes little contribution (Makhlouf et al. 1989). Peptide histidine isoleucine (PHI), or its human equivalent, PHM, is derived from the same gene as VIP and has similar effects. Another member of the VIP family, PACAP, also relaxes intestinal muscle and is found in enteric neurons (Sundler et al. 1992, Portbury et al. 1995a). Immunoneutralization and pharmacological antagonism of the PACAP receptors both indicate that it contributes to enteric inhibitory transmission in some regions (Grider et al. 1994, McConalogue et al. 1995, Katsoulis et al. 1996). As VIP, PHI and PACAP are probably contained in the same enteric neurons that innervate the muscle (Portbury et al. 1995a), and as these peptides act through the same (VPAC) receptors, there is likely to be a peptide mixture involved in transmission. Next came evidence that NO is an enteric inhibitory transmitter (Rand 1992, Sanders & Ward 1992). NO is an unusual chemical messenger because it is not stored, but is created on demand when the synthesizing enzyme,
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nitric oxide synthase (NOS), is activated. NO was earlier identified as an endothelium-derived vasodilator, and blockers of NOS were quickly developed. NOS inhibitors and scavengers of NO reduce inhibitory transmission to gastrointestinal muscle in many species (Bult et al. 1990, Li & Rand 1990, Toda et al. 1990), including humans (Maggi et al. 1991), an observation that has been replicated in innumerable studies since the original observations that were published in 1990 and 1991. Moreover, NOS is readily revealed in tissue, either by enzyme histochemistry or by immunohistochemistry, and it was quickly found to be located in inhibitory neurons supplying gut muscle (Bredt et al. 1990, Costa et al. 1992, Ward et al. 1992). NO is membrane permeable, and it crosses the membrane to interact with its receptor, soluble guanylyl cyclase, in the cytoplasm. Thus, within a short time, three quite different substances were in contention as inhibitory transmitters – a phosphorylated purine nucleotide (ATP), a 28 amino-acid peptide (VIP) and its close relatives, and a low molecular weight free radical (NO) (Fig. 4.1B). It is now known that all three contribute, and they are released from the same axons. The transmitters contribute to relaxation in different proportions, depending on region and species (Costa et al. 1986, Crist et al. 1992, Suthamnatpong et al. 1993, Furness et al. 1995b, Katsoulis et al. 1996). The major component of transmission seems to be through a combination of ATP and NO, and antagonism of the action of ATP with apamin plus block of NO production with an NOS inhibitor in most cases blocks the inhibition of the muscle (Lecci et al. 2002). VIP and NOS are found together in the inhibitory motor neurons (Chapter 2), and NOS immunoreactivity is a good marker of the neurons. There is also evidence that carbon monoxide could contribute to inhibitory transmission (Xue et al. 2000). One consequence of there being multiple transmitters of the inhibitory neurons may be that a deficiency in one transmitter can be partly compensated. Animals in which neuronal NOS is eliminated by gene knockout have surprisingly little wrong with them. The mice show evidence of pyloric stenosis, the stomach is dilated, gastric emptying is slowed and the gastric muscle is thickened (Huang et al. 1993, Mashimo et al. 2000). There is little evidence of deficiencies elsewhere in the gastrointestinal tract. Transmitters at neuro-neuronal synapses Three types of post-synaptic event occur at neuro-neuronal synapses in the enteric nervous system: fast excitatory post-synaptic potentials (fast EPSPs), slow excitatory potentials (which have several forms, see below) and slow inhibitory post-synaptic potentials (slow IPSPs). Much of the analysis of transmitter pharmacology has been by examination of synaptic events that were elicited by electrical stimulation of nerve fiber pathways. This confounds the
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evaluation of transmission at individual synapses, because this type of stimulation activates many nerve fibers at the same time, so that the consequent synaptic event will generally represent the combined effect of transmitter released from multiple inputs to the neurons. Moreover, electrical stimulation activates axons both antidromically and orthodromically. Antidromic impulses travel down collaterals of the axons and release transmitter at sites closer to the cell body than the site of stimulation (Bornstein et al. 1986). Thus the responses to electrical stimulation sometimes fail to provide insight into the origin of the presynaptic nerve fibers. Fast excitatory post-synaptic potentials Fast EPSPs were recorded with intracellular microelectrodes from myenteric and submucosal neurons in the 1970s (Nishi & North 1973a, Hirst et al. 1974, Hirst & McKirdy 1975). The fast EPSPs had amplitudes that were graded with stimulus strength up to about 20 mV above resting membrane potential, measurement of exact amplitude being confounded by the initiation of action potentials (Fig. 4.2). The early studies showed that the fast EPSPs were substantially reduced in amplitude or were blocked by hexamethonium or curare, which are blockers of nicotinic acetylcholine receptors (Nishi & North 1973a, Hirst et al. 1974). The anticholinesterase physostigmine, which prevents the metabolism of ACh, prolonged the fast EPSPs (North & Nishi 1974). These observations were perfectly consistent with observations
Fig. 4.2 Fast EPSPs recorded from an S neuron of the myenteric plexus. The downward deflections before the EPSPs are the artifacts of the single stimulus pulses used to evoke them. The strength of stimulus has been increased from A to C. In A, only an EPSP is seen, in B a local response triggered by the EPSP is seen, and in C a full action potential is initiated in the soma of the S neuron. Reproduced from Nishi and North (1973).
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that peristaltic reflexes are blocked by nicotinic receptor antagonists (Bayliss & Starling 1900a, Cannon 1911, Henderson 1928, Feldberg & Lin 1949, Hukuhara et al. 1958) and indicated that ACh, acting at nicotinic receptors, is a major transmitter to enteric neurons. The nicotinic receptor is a ligand-gated ion channel that provides a non-specific (depolarizing) cation conductance. The subunit compositions of nicotinic ACh receptors (nAChRs) of myenteric neurons have been investigated by agonist rank-order potency and immunohistochemistry in guinea-pig. The experiments indicate that the predominant expression is of nAChRs composed of α3, α5, β2, and β4 subunits. These subunits may combine in a homogeneous population of receptors with unique pharmacological properties, or multiple receptors of different subunit composition may be expressed by individual neurons (Zhou et al. 2002). It was not until comparatively recently that a careful examination of the components of fast EPSPs that resist nicotinic receptor block was undertaken (Galligan & Bertrand 1994, Lepard et al. 1997, Lepard & Galligan 1999, Nurgali et al. 2003a, 2004). These experiments indicate that the dominant part of fast transmission is cholinergic, but that it also involves ATP, acting at P2X receptors, and 5-HT, acting through 5-HT3 receptors (Fig. 4.3). Because many fast EPSPs are fully blocked by hexamethonium, these EPSPs are purely cholinergic. In about 60% of neurons with a hexamethonium resistant fast EPSP, the EPSP after nicotinic block was reduced in amplitude or abolished by an antagonist of P2X receptors for ATP, PPADS (Lepard & Galligan 1999). In about 10% of neurons, a 5-HT3 receptor antagonist blocks or reduces
Fig. 4.3 Pharmacological dissection of fast EPSPs in S neurons of the myenteric plexus of the
guinea-pig ileum. A: A fully cholinergic EPSP is blocked by the nicotinic receptor blocker hexamethonium. B: A fast EPSP is partly reduced by hexamethonium. The remaining component is largely blocked by the P2X purine receptor antagonist PPADS. C: A fast EPSP in which the response remaining after the administration of hexamethonium is blocked by the 5-HT3 receptor blocker ondansetron. ACh, acetylcholine. The percentages of synaptic responses of each type are indicated. Reproduced from Galligan et al. (2002).
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fast EPSPs that remain in the presence of hexamethonium (Zhou & Galligan 1999). Thus, at some synapses ATP and/or 5-HT could be co-transmitters with ACh. In some S neurons, the three antagonists in combination do not fully block fast EPSPs, which implies that there is one or more other transmitters of fast EPSPs. Pharmacological and immunohistochemical data suggest that this might include glutamate, acting through AMPA receptors, and that the fast EPSPs seen in a minority of AH neurons might also be glutamate mediated (Liu et al. 1997). Some of the neurons that use ATP as a transmitter project anally in the myenteric plexus (Johnson et al. 1999, Lepard & Galligan 1999, Bian et al. 2000). Fast EPSPs in submucosal neurons had also been assumed to be exclusively nicotinic until they were recently more carefully analyzed (Monro et al. 2002). Blocking nicotinic receptors reduced the amplitudes of fast EPSPs in all submucosal neurons and abolished them in 73%. In about 25% of neurons there was a component of transmission attributable to purine P2 receptors and in about 15% a component was mediated through 5-HT3 receptors. Fast excitatory synaptic inputs to IPANs are rarely seen in myenteric neurons of the guinea-pig small and large intestine (Bornstein et al. 1994) or in the mouse colon (Furukawa et al. 1986, Nurgali et al. 2004) and when recorded, fast EPSPs are of low amplitude (Iyer et al. 1988, Bornstein et al. 1994, Tamura et al. 2001), although in some other regions and species, such as the pig (Cornelissen et al. 2001), presumptive IPANs do exhibit fast EPSPs. Furthermore, fast EPSPs are not observed or have very small amplitudes in submucosal IPANs (Bornstein et al. 1989, Evans et al. 1994). Morphological and pharmacological data indicate that somatic nicotinic receptors, which would be expected to mediate fast synaptic responses, occur in IPANs. First, nicotinic acetylcholine receptors have been detected immunohistochemically on the surface membranes of myenteric IPANs (Dogiel type II morphology, calbindin positive) in the guinea-pig small intestine (Kirchgessner & Liu 1998), and, second, nicotine application leads to a fast depolarization of myenteric IPANs via a direct action (it occurs in the presence of TTX and in low Ca2+, high Mg2+ solution) on post-synaptic nicotinic receptors (Schneider & Galligan 2000). Furthermore, synapses on the neurons are immunoreactive for the vesicular acetylcholine transporter (Li & Furness 1998), which is generally believed to be a reliable marker of cholinergic nerve endings. It is therefore not clear why fast EPSPs are not usually recorded from these neurons in response to synaptic stimulation. It is possible that the nicotinic receptors are not located at the synapses. Myenteric IPANs are also immunoreactive for P2X2 receptors (Castelucci et al. 2002), through which fast EPSPs could be mediated.
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Fig. 4.4 Examples of slow EPSPs in an AH and in an S neuron of the myenteric plexus. In each case the slow EPSP was elicited by a burst of three pulses at 10 Hz at the arrows. The downward deflections are potential changes caused by constant amplitude current pulses. The increased size of the consequent potential changes during the slow EPSPs indicates that cell resistance has increased. Reproduced from Bornstein (1984).
Slow excitatory post-synaptic potentials Slow EPSPs, usually elicited by stimulation of interganglionic connectives at 10–20 Hz for a few seconds, are observed in most enteric neurons (Wood & Mayer 1978, Morita & North 1985) (Fig. 4.4). There is depolarization of the membrane potential, an increase in input resistance and an increase in soma excitability, as exemplified by an increase in the number of spikes elicited by an intracellular 500 ms depolarizing pulse. The effects commonly peak within 10–15 s, last for 1–4 min (Morita & North 1985) and are primarily due to a reduction in resting K+ conductance, including gKCa (North & Tokimasa 1983). This is reported to be combined in some cases with activation of a Cl– conductance (Bertrand & Galligan 1994, Starodub & Wood 2000a). In IPANs there is also a reduction of the late AHP. Slow EPSPs in IPANs are mimicked by the NK3 tachykinin receptor agonist senktide (Schneider & Galligan 2000) and are partially blocked by the NK3 receptor antagonist SR142801 or by the NK1 antagonist SR140333 (Alex et al. 2001, Johnson & Bornstein 2004). These slow synaptic inputs are proposed to come primarily from neighboring IPANs that form a selfreinforcing network and contain tachykinins (Chapter 3). ACh, acting via muscarinic receptors, also elicits slow depolarizing responses in myenteric IPANs, but, as the majority of cells retain stimulation-evoked slow EPSPs in the presence of muscarinic antagonists, ACh is unlikely to be the principal slow neurotransmitter (Morita & North 1985). Slow EPSPs are mimicked both by activators of the adenylyl cyclase (AC)– PKA pathway and by activators of the phospholipase C (PLC)–diacyl glycerol (DAG)–PKC pathway. AC-PKA activators that mimic slow EPSPs include forskolin (Morita & North 1985, Bertrand & Galligan 1995), CCK (Palmer et al. 1987b), histamine (Nemeth et al. 1986), gastrin-releasing peptide (GRP or mammalian bombesin) (Zafirov et al. 1985), PACAP (Christofi & Wood
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1993a), and VIP (Williams & North 1979). Inhibition of their effects by adenosine-induced stimulation of Gαi, a G-protein that is negatively coupled to AC, suggest that these mediators act via stimulation of AC and subsequent increase in intracellular levels of cAMP (Palmer et al. 1987a). Mimics of the slow EPSP that act through the PLC-DAG-PKC pathway include CGRP and tachykinins, such as substance P, acting through NK3 receptors. Although 5-HT causes slow excitation principally through a PKC pathway, it also activates PKA to a small extent (Wood & Mayer 1979b, Pan et al. 1997). Two lines of evidence suggest that the PLC-DAG-PKC pathway, rather than the AC-PKA pathway, mediates the slow EPSP. First, the PLC blocker D609 inhibits the slow EPSP in IPANs (Bertrand & Galligan 1995). Second, adenosine does not inhibit slow EPSPs, suggesting that the receptors primarily mediating slow EPSPs are not coupled to AC by GS (Pan et al. 1997). Thus, receptors coupled to inositol phospholipid hydrolysis and the PKC pathway, such as NK3 receptors (Guard et al. 1988), are probably the principal mediators of these slow excitatory responses in IPANs of the guinea-pig ileum. However, antagonists of receptors for CCK, which acts on IPANs through the AC–PKA pathway, partly block slow EPSPs recorded from IPANs (Schutte et al. 1997). Sustained slow post-synaptic excitation A prolonged synaptic event, sustained slow post-synaptic excitation (SSPE), is observed in IPANs (Clerc et al. 1999, Alex et al. 2001, 2002, Nurgali et al. 2003b). The SSPE outlasts nerve stimulation by many minutes (Fig. 4.5), and successive SSPEs show substantial facilitation. The increased excitation during a facilitated SSPE can be maintained after stimulation for as long as 4 hours. The SSPE is induced by repeated, low frequency (0.5–2 Hz) trains of stimulation of synaptic inputs (Clerc et al. 1999, Alex et al. 2001). It involves increased soma excitability that facilitates with successive stimuli. During the stimulation, the soma excitability of the IPANs slowly increases, they depolarize, and their input resistance increases markedly, often by over two-fold. This magnitude of increase, accompanied by depolarization, implies that K+ currents are inhibited. During the SSPE, late AHPs are reduced in amplitude and duration and can eventually be completely suppressed. In addition, anodal break action potentials occur more often after hyperpolarizing pulses (Clerc et al. 1999) which suggests that a hyperpolarization activated current (Ih, Chapter 2) could be increased in its effect during the SSPE. With sufficient depolarization of the neurons, invasion by antidromic action potentials is suppressed. Spontaneous action potentials were rare and the occurrence of fast EPSPs was not reported during the heightened excitability caused by the SSPE (Clerc et al. 1999).
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Fig. 4.5 Change in neuronal excitability during a sustained slow post-synaptic excitation (SSPE) recorded from an IPAN of the guinea-pig distal colon. Stimulation of synaptic inputs to the neuron at 1 Hz for 4 min (bar) caused depolarization of the membrane potential (MP), increased input resistance (Rin) and increased excitability (measured as the numbers of action potentials elicited by 500 ms depolarizing pulses (AP/500 ms). Increased excitability lasted for more than 1 h. Reproduced from Nurgali et al. (2003).
The suppression of the late AHP suggests that K+ channels are targeted during the SSPE, possibly by phosphorylating enzymes (Chapter 2). Consistent with this, the PKC stimulant phorbol dibutyrate (PDBu), caused somatic excitability increases, membrane depolarization, and increased input resistance in a concentration-dependent manner, mimicking the SSPE (Pan et al. 1997, Kawai et al. 2003). The SSPE is blocked by inhibitors of PKC, but not by inhibition of PKA (Nguyen et al. 2004). PDBu suppressed the late AHP and the effects of PDBu on the late AHP were indistinguishable from those observed during the SSPE (Clerc et al. 1999, Kawai et al. 2003, Nguyen et al. 2004). It seems that PDBu mimics the SSPE in IPANs by closing IK channels that are responsible for the AHP or restricting their opening by Ca2+, and reducing the current carried by these and other K+ channels that are active at rest (Nguyen et al. 2005). Although PDBu mimics the SSPE, the activator of novel PKC isoforms is ineffective, indicating that a conventional PKC isoform is involved in the SSPE (Nguyen et al. 2005). Immunohistochemical investigation indicates that the only conventional PKC that is prominent in the neurons is PKCγ (Poole et al. 2003). An obvious candidate neurotransmitter for the SSPE could be a tachykinin. However, experimental data demonstrate that this is unlikely, because the SSPE persists after block of NK1 and NK3 tachykinin receptors on enteric neurons (Alex et al. 2001, 2002).
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Slow synaptic transmission in S neurons Slow EPSPs have been recorded in identified S neurons in the myenteric and submucosal ganglia. Slow EPSPs in inhibitory motor neurons were blocked by NK1 receptor antagonists (Thornton & Bornstein 2002). However, slow transmission to descending interneurons was not affected by NK1 or NK3 receptor block. These slow EPSPs are antagonized by P2Y purine receptor antagonists (Bornstein et al. unpublished). Pharmacological data indicate that slow transmission to secretomotor neurons in the submucosal is also mediated through purine P2Y receptors (Hu et al. 2003, Monro et al. 2004). The possibility that S neurons exhibit a type of slow transmission elicited by low-frequency stimulation of synaptic inputs, similar to that seen in AH neurons, has been investigated in the guinea-pig small intestine (Alex et al. 2002). A slow depolarization, accompanied by increased excitability, was elicited in only one group of S neurons, inhibitory motor neurons to the circular muscle, by stimulation of synaptic inputs at low frequency, 1 Hz up to 4 min. However, unlike the SSPE of IPANs, depolarization began to decline immediately at the end of stimulation. The excitation and depolarization were blocked by a NK1 tachykinin receptor antagonist. Slow inhibitory post-synaptic potentials Slow IPSPs are observed in a small proportion of myenteric neurons when adjacent internodal strands of the plexus are stimulated (Wood & Mayer 1978, Johnson et al. 1980, Hodgkiss & Lees 1984), but they are far more frequently encountered in submucosal neurons (Hirst & McKirdy 1975). The inhibitory potentials in the two plexuses have quite different properties. In the myenteric plexus, Johnson et al. (1980) recorded IPSPs in about 5% of S neurons and 15% of AH neurons. The IPSP was only evoked by repetitive pulses, it lasted 2–40 s and was associated with a decrease in membrane resistance. Because the reversal potential for the slow IPSP was around –100 mV, close to the potassium equilibrium potential, it was presumed to be the result of an increase in potassium conductance. In submucosal ganglia, transmural stimuli initiated slow IPSPs lasting 1–5 s in about 30–40% of neurons (Hirst & McKirdy 1975, Surprenant 1984). Single stimuli evoked slow IPSPs of up to 18 mV; with pairs or brief bursts of stimuli (20 Hz), hyperpolarizations up to 40 mV in amplitude could be obtained. The IPSP in submucosal neurons is also associated with an increase in membrane conductance to potassium ions. North and Surprenant (1985) have shown that some slow IPSPs in submucosal ganglia are mediated by the release of norepinephrine from sympathetic noradrenergic nerves that supply these ganglia. The toxin for
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adrenergic neurons, 6-hydroxydopamine, irreversibly blocked the IPSP and also depleted norepinephrine from nerve terminals in the ganglia. The IPSP was mimicked by local application of norepinephrine and both the effect of norepinephrine and the IPSP were blocked by antagonists that act at α2 receptors for catecholamines. It had previously been shown that guanethidine, which blocks action-potential initiated release of norepinephrine from nerves, abolishes the slow IPSP (Hirst & McKirdy 1975). However, there is also a non-noradrenergic IPSP in submucosal ganglia because IPSPs can be evoked by transmural stimulation after extrinsic denervation which causes degeneration of noradrenergic endings in the ganglia (Bornstein et al. 1988). Somatostatin is possibly the transmitter of the non-noradrenergic IPSP (Shen & Surprenant 1993). Presynaptic inhibition at neuro-neuronal synapses There is considerable evidence that endogenous transmitters in the enteric nervous system can reduce transmitter release at excitatory synapses. The first evidence of this was a progressive decrease in the amplitudes of fast EPSPs that was observed when presynaptic fibers were stimulated at frequencies of more than about 0.05 Hz (Morita et al. 1982b). This inhibition appears to be due to a feedback inhibition by ACh released from the nerve endings, as it is substantially reduced by blockers of muscarinic receptors. Morita et al. (1982b) also found that the non-cholinergic slow EPSP in both S and AH neurons was enhanced by hyoscine. This implies that the stimuli used to evoke the slow EPSP also stimulated cholinergic neurons that acted presynaptically to reduce the output of the non-ACh transmitter. The principal site of action of norepinephrine, released from sympathetic nerve endings, to inhibit motility in the intestine is within the myenteric ganglia (Chapter 9). Consistent with this, Hirst and McKirdy (1974b) found that stimulation of mesenteric nerves to the intestine reduced or abolished fast EPSPs in myenteric S neurons. The effect is largely due to inhibition of excitatory transmitter release in the myenteric plexus, and to both inhibition of release and to post-synaptic inhibition in submucosal ganglia. Norepinephrine reduces the amplitudes of fast EPSPs in both myenteric and submucosal neurons (Nishi & North 1973b, North & Nishi 1974, Hirst & Silinsky 1975) and recent studies indicate that both cholinergic and non-cholinergic components of fast EPSPs are inhibited by norepinephrine receptor agonists (Lepard et al. 2004). In the same myenteric neurons, norepinephrine does not reduce the effect of acetylcholine (ACh) (Nishi & North 1973b), and in the submucosal neurons the reduction in fast EPSP amplitude is seen in cells in which norepinephrine has no effect on membrane potential (Hirst & Silinsky 1975). These observations indicate that norepinephrine acts presynaptically
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to reduce the release of ACh from synapses impinging on S neurons. Epinephrine and norepinephrine generally have no direct effects on the soma membrane of myenteric S cells, in contrast to the prominent hyperpolarizations recorded in many submucosal neurons. In contrast, hyperpolarization of myenteric IPANs is commonly seen (Morita & North 1981b). 5-HT has two effects on fast EPSPs; it increases their amplitude by an action on 5-HT4 receptors and depresses them through action on 5-HT1A receptors (Galligan et al. 1988, Pan & Galligan 1994). The presynaptic inhibition of the fast EPSP occurs without 5-HT modifying the effect of iontophoretically applied ACh (North et al. 1980). 5-HT also depresses the slow EPSP in myenteric AH and S neurons with a time course similar to its depression of the fast EPSP (North et al. 1980, Johnson et al. 1981, Bornstein et al. 1984b). There is as yet no direct evidence that 5-HT released from nerve endings in the plexus causes presynaptic inhibition. The release of adenosine from endogenous sources, possibly neurons, inhibits the release of excitatory transmitters, both those involved in fast EPSPs and those of slow EPSPs (Christofi & Wood 1993b). The source of adenosine could be the extracellular breakdown of neurally released ATP. Edwards et al. (1976) examined interactions that occur when two sets of transmural electrodes are used to stimulate submucosal pathways at separate points. Their experiments indicate that there are neurons that cause presynaptic inhibition impinging on the cholinergic endings on submucosal S neurons. Sites within the reflex circuitry where specific pharmacologies of transmission can be deduced to occur The identification of transmitters at specific synapses in the circuits has been facilitated by the use of partitioned organ baths (Tonini & Costa 1990). The partitions divide the solutions that bathe adjacent regions of intestine so that the pharmacology of transmission in each region can be separately investigated (Fig. 4.6). It is common to have three compartments – a stimulus compartment from which reflexes are initiated, an intermediate compartment through which reflexes are conducted, and a recording compartment where the responses of the innervated tissue (the circular or longitudinal muscle) are recorded. Bathing solution can also be collected from the compartments in order to detect transmitters that are released into each (Grider & Makhlouf 1986, Grider et al. 1987). Conduction that is dependent on pathways that have synapses in a compartment can be blocked by lowering Ca2+ to 0.1 mM and increasing Mg2+ concentration to 10 mM in that compartment (Johnson et al. 1996). Effects that are not prevented by this synaptic block are due to long axons that cross the compartment without synapsing. Recordings made from neurons during reflexes reveal fast EPSPs, suggesting that fast
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Fig. 4.6 Partitioned organ bath that is used to isolate, pharmacologically, sections of the reflex
pathways in the intestine. The partitions divide the bath into compartments that can be irrigated with solutions of different compositions, or containing different drugs. In this way, the pharmacology of transmission from neurons in different parts of the nerve pathways can be investigated.
transmitters, ACh, ATP and 5-HT, dominate transmission (Hirst et al. 1975, Bornstein et al. 1991a). Recent evidence indicates that slow transmission at synapses on descending interneurons also occurs during the reflexes (Thornton & Bornstein 2002). Ascending reflexes: transmission between interneurons and from interneurons to motor neurons Ascending excitatory reflexes are blocked by nicotinic receptor antagonists in the guinea-pig small intestine (Smith & Furness 1988, Tonini & Costa 1990, Johnson et al. 1996), as well as in other regions and species, including dog, guinea-pig, and rat colon (Hukuhara et al. 1958, Costa & Furness 1976, Furness et al. 2002) and human jejunum (Grider 1989a). Studies using partitioned organ baths have used guinea-pig ileum, in which transmission within the intermediate and recording chambers is fully blocked or very substantially reduced by hexamethonium, implying that the primary transmitter at interneuron-to-interneuron and interneuron-to-motor neuron synapses is ACh acting at nicotinic synapses (Tonini & Costa 1990, Johnson et al. 1996). This agrees with the immunohistochemical identification of the neurons: only one class of ascending interneuron occurs in the small intestine, and these contain choline acetyltransferase (Chapter 2). There is also a component of excitatory transmission that is muscarinic, probably at IPAN-to-interneuron synapses, and is blocked by hyoscine or atropine. Despite the observation that ascending interneurons have tachykinin immunoreactivity (Chapter 2),
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transmission between interneurons is not attenuated by tachykinin receptor blockers (Johnson et al. 1996). In contrast to other studies, Spencer et al. (2000) reported that hexamethonium (300 μM) in most cases caused only about 50% reduction of the contractions elicited oral to a physiological stimulus, mucosal stroking, in the guinea-pig ileum. The residual response was substantially reduced by the purine receptor blocker PPADS. Transmission from IPANs to ascending reflex pathways In the guinea-pig ileum, transmission at the first synapse, between IPANs and interneurons, appears to be due to a mixture of cholinergic (nicotinic) transmission and tachykinins acting at NK3 receptors, with a small component of excitatory muscarinic transmission (Johnson et al. 1996). In the rat colon, however, studies using partitioned baths clearly implicate CGRP in transmission at this first synapse (Grider 1994). Two stimuli that elicit enteric reflexes, muscle stretch and mechanical stimulation of the mucosa, caused CGRP release in the central but not in the oral or anal chambers. Moreover, two different antagonists of CGRP receptors reduced, but did not block, ascending excitatory reflexes (Grider 1994). It is pertinent that IPANs do not contain CGRP in the guinea-pig small intestine (Gibbins et al. 1985), whereas CGRP is contained in these neurons in mice (Furness et al. 2004b), humans (Timmermans et al. 1992), pig (Scheuermann et al. 1987a), sheep (Chiocchetti et al. 2004) and rat (unpublished). Thus CGRP is a probably a co-transmitter released from IPANs in most species other than guinea-pig. Descending inhibitory reflexes In the guinea-pig small intestine or colon, the overall transmission in descending inhibitory pathways in a non-partitioned bath is not blocked by hexamethonium when the distance that the reflex travels is less than about 10 mm (Costa & Furness 1976, Smith & Furness 1988, Johnson et al. 1996). However, at distances greater than 30 mm in the small intestine the reflex was almost blocked (Smith & Furness 1988) and in the colon it was blocked at distances greater than about 10 mm (Costa & Furness 1976). Although most IPANs make synaptic connections close to their cell bodies, some IPANs project long axons in an anal (but not an oral) direction. In the guinea-pig small intestine, many extend for up to 30 mm (Brookes et al. 1995). Consistent with this anatomical arrangement, there is a part of the reflex pathway, over distances of about 20–30 mm from the stimulus, in which transmission is not prevented by block of synaptic transmission in low Ca2+, high Mg2+ solution (Johnson et al. 1996). So it is possible that nicotinic antagonist-resistant transmission occurs in monosynaptic reflex pathways, from IPANs
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direct to motor neurons. IPANs with long descending axons also occur in the guinea-pig colon (Bian et al. 2004). However, at distances greater than about 15 mm transmission from the IPANs appears to be nicotinic (Bian et al. 2004). In other species, dog, human, and rat, where the investigations possibly involved longer pathways, nicotinic receptor antagonists also blocked the descending inhibitory reflexes (Hukuhara et al. 1958, Grider 1989a, Kanada et al. 1993, Furness et al. 2002). In the rat colon, transmission at the first synapse, between IPANs and interneurons or motor neurons, is prevented by nicotinic receptor antagonists (Bian et al. 2003). The nature of the transmitter at the first synapse in the pathway, between IPANs and descending interneurons, appears to differ between species and between regions of the intestine. In rat and guinea-pig colon, transmission appears to be predominantly nicotinic (Bian et al. 2003, 2004). However, transmission at this first synapse in the guinea-pig small intestine is largely resistant to nicotinic block (Johnson et al. 1996), and transmission from IPANs to motor neurons in monosynaptic pathways was unaffected by hexamethonium (Bian et al. 2000). When mucosal compression was used to elicit descending reflexes in the guinea-pig small intestine, the NK3 receptor antagonist SR142801 reduced the reflex response, but it did not do so when the stimulus was distension (Johnson et al. 1998a). NK1 receptor antagonism had no effect in either case. This suggests that transmission from the submucosal IPANs, which are activated by mucosal distortion, is partly through tachykinins acting at NK3 receptors, but transmission from myenteric IPANs is through another transmitter. Similar to ascending excitatory reflexes that are discussed above, CGRP appears to be a co-transmitter at the IPAN-to-interneuron synapse in the rat colon (Grider 1994). Because CGRP is contained in the IPANs it is likely to be a co-transmitter from these neurons in most species, including humans (see above). Transmission between interneurons in descending pathways is difficult to evaluate, even in a partitioned bath, because the intermediate chamber contains synapses between the long descending axons of IPANs and interneurons, as well as synapses between interneurons. The reduction in transmission through the intermediate chamber that is caused by NK3 receptor antagonism (Johnson et al. 1996) is probably due to a block of IPAN to interneuron synapses because IPANs, but not descending interneurons, contain tachykinins. Another component of transmission to descending interneurons is through a slowly acting transmitter that is not a tachykinin acting through NK1 or NK3 receptors (Thornton & Bornstein 2002), but is possibly ATP acting through P2Y receptors (Bornstein et al. unpublished). The classes of descending interneuron are those immunoreactive for ChAT plus somatostatin (SOM), those immunoreactive for ChAT plus NOS, VIP and other substances, and those immunoreactive for ChAT plus 5-HT
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(Chapter 2). The ChAT/5-HT neurons do not make connections with the inhibitory muscle motor neurons (Young & Furness 1995), and pharmacological studies, which show a lack of effect of receptor antagonists, suggest that 5-HT is not a transmitter in local descending inhibitory motility reflexes (Yuan et al. 1994). Electron microscope studies show that the ChAT/SOM neurons receive very few inputs from IPANs, but that over 80% of their inputs are from other ChAT/SOM neurons. On the basis of these observations, it has been suggested that ChAT/SOM descending interneurons are a conduit for the passage of migrating myoelectric complexes along the guineapig intestine (Chapter 2). This theory alone does not explain the slow rate of progress of the MMC (Chapter 5). Somatostatin neurons can also be activated by locally applied mucosal distortion stimuli (Bornstein et al. 1991a) or by distension (Thornton & Bornstein 2002), and are therefore likely to have some local role. Nevertheless, the ChAT/NOS neurons are the ones most likely to be directly involved in local descending inhibitory reflexes (Chapter 3). The ChAT/NOS interneurons form descending chains (Chapter 2). A component of fast excitatory transmission is blocked by PPADS, an antagonist of purine (ATP) receptors (see above), so it is possible that ATP contributes to fast EPSPs in descending interneurons or the inhibitory motor neurons. In fact, purine receptor antagonists reduce transmission from interneurons to the final motor neurons, suggesting that one of the transmitters of descending pathways is ATP, acting at P2X receptors (Bian et al. 2000). Transmission from the long axons of IPANs to the inhibitory motor neurons appears to involve a tachykinin component. IPANs contain tachykinins and immunoreactivity for NK1 receptors is prominent on the NOSimmunoreactive inhibitory motor neurons (Portbury et al. 1996b). Moreover, the NK1 receptor antagonist SR140333 reduces descending inhibitory reflexes (Johnson et al. 1998a). In these experiments, inhibitory responses in the muscle anal to distension or compression of the mucosa were recorded in a three-chambered organ bath. When SR140333 was applied to the recording chamber, reflex responses were reduced by 40–50%. Tachykininmediated slow EPSPs, blocked by SR140333, have been recorded from the inhibitory motor neurons (Alex et al. 2002, Thornton & Bornstein 2002). This is consistent with data from analysis of responses to reflex activation in the guinea-pig small intestine, using receptor internalization as an indication of their activation. NK1 receptors are rapidly internalized following binding of agonists and it was found that activation of reflex pathways, by movement of the mucosal villi, caused receptor internalization (Southwell et al. 1998). These experiments implied that tachykinins were released synaptically during the reflexes, acted on the NK1 receptors of the inhibitory motor neurons, and caused receptor internalization.
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NO, which is probably released from descending interneurons containing NOS, does not appear to be a primary transmitter between neurons. In fact, it appears to act as a retrograde transmitter, being released from the cell bodies of the interneurons and acting back on the endings of the terminals of IPANs to reduce transmitter release (Yuan et al. 1995). Descending excitatory reflexes It is generally accepted that excitation is conducted down the intestine by reinforcement from a moving bolus, as first clearly enunciated by Bayliss and Starling (1899). However, once activated, the excitatory pathways can propagate excitation for several centimeters in an anal direction without this reinforcement (Chapter 5). Descending excitatory reflexes, elicited by distension, are reduced by about 75% by the 5-HT3 receptor antagonist granisetron (Monro et al. 2002), and descending excitation evoked by electrical stimulation of intramural nerves is also reduced by a blocker of 5-HT3 receptors (Jin et al. 1989), implying that the 5-HT containing neurons are involved in this component of propulsive reflexes (Fig. 4.7). Descending excitatory reflexes
Fig. 4.7 Transmitters of ascending and descending reflex pathways supplying the muscle.
Transmitters are indicated adjacent to the synapses where they act. Information flows from the IPANs to motor neurons directly, or through one or more interneurons. Primary transmitters and their receptors are indicated (e.g. ACh: Nic = primary transmitter, acetylcholine, acting at nicotinic receptors). Where roles are minor or uncertain, the information is in parentheses.
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are also sensitive to block of ATP receptors, suggesting that ATP may also be a transmitter in the pathway (Spencer et al. 1999, Monro et al. 2002). Comparison of the pharmacology of peristaltic reflexes recorded using a modified Trendelenburg method (Fig. 3.2B) suggests that there is divergence in the pathways for peristalsis and the descending excitatory reflex (Monro et al. 2002). This is consistent with ACh/NOS interneurons having a primary role in the peristaltic reflex and ACh/5-HT neurons being important for descending excitation. Transmission from entero-endocrine cells to IPANs Chemicals in the lumen or mechanical stimulation of the lining of the intestine do not directly excite the responsive endings of IPANs that innervate the mucosa. Instead, they trigger the release of 5-HT (and possibly other mediators) from entero-endocrine cells. The apical surfaces of the endocrine cells are in contact with the luminal contents and their basal surfaces are adjacent to the connective tissue space beneath the mucosal epithelium. These enteroendocrine cells are often called enterochromaffin cells because they give a positive histochemical reaction with the chromaffin staining technique. The cells were recognized by histologists in the 19th century and were found to contain 5-HT by Erspamer and Asero (1952). They are very numerous throughout the small and large intestine (Solcia et al. 1981). Addition of 5-HT to the fluid bathing the lumen of the intestine, or its local application to the luminal surface, elicits peristaltic reflexes (Bülbring & Lin 1958, Hukuhara et al. 1960). These experiments were conducted in vitro, where the mucosal barrier is not maintained, and so 5-HT is able to cross the epithelium and act on the nerve endings in the mucosa. In addition, 5-HT release occurs when intraluminal pressure, a stimulus for enteric reflexes, is increased (Bülbring & Crema 1959). Later experiments showed that 5-HT activates the intramucosal endings of IPANs that have cell bodies in either the myenteric or submucosal ganglia (Kirchgessner et al. 1992, Bertrand et al. 1997, Pan & Gershon 2000). The mucosal endings are almost certainly the sites from which 5-HT initiates peristaltic and secretomotor reflexes. Pharmacological experiments have cemented the hypothesis that 5-HT released from entero-endocrine cells is an initiator of enteric reflexes, although the receptor types on the responsive endings of the IPANs are different in different animal species and different intestinal regions. Using the dog jejunum, in which the stimulus and recording segments were separately perfused, it was found that block of 5-HT3 receptors in the stimulus region substantially reduced the ascending excitatory and descending inhibitory reflexes (Neya et al. 1993). Consistent with this, the excitation of IPAN terminals in the mucosa of the guinea-pig small intestine is blocked by 5-HT3 receptor an-
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tagonists (Bertrand et al. 2000), and descending reflexes elicited by distension in the rat small intestine are also reduced by these antagonists (Kanada et al. 1993). In addition, the facilitation of peristalsis caused by intraluminal 5-HT in a Trendelenburg-type preparation of guinea-pig small intestine was substantially reduced by 5-HT3 receptor antagonists, but not by antagonism of other receptors, including 5-HT1 and 5-HT4 receptors (Tuladhar et al. 1997). However, in the guinea-pig colon, 5-HT3 and 5-HT4 receptor antagonists, applied to the stimulus compartment of a partitioned organ bath, both reduced reflex responses, and their effects were additive (Foxx Orenstein et al. 1996). In similarly prepared segments of human jejunum and rat colon, responses were reduced by a 5-HT4 antagonist, but not a 5-HT3 antagonist (Foxx Orenstein et al. 1996, Grider et al. 1996). In another study, 5-HT3 and 5-HT4 receptor antagonists both reduced descending inhibition in the rat colon (Bian et al. 2003). Consistent with the involvement of 5-HT4 receptors, agonists for these receptors enhance peristaltic reflexes in the guinea-pig and rat colon (Foxx Orenstein et al. 1998, Kadowaki et al. 2002). The contribution of another receptor type, the 5-HT1P receptor, needs to be considered. This receptor has been defined by its pharmacological properties, but has not yet been cloned. Nevertheless, the pharmacology is suggestive of it having a role in the activation of IPANs by 5-HT released from entero-endocrine cells (Gershon 2004). In addition to being involved in the initiation of peristaltic reflexes, 5-HT is an initiator of secretomotor reflexes (Cassuto et al. 1982b, Sidhu & Cooke 1995, Cooke et al. 1997b). 5-HT applied intraluminally in rats induced secretion that was inhibited by the nicotinic blocker hexamethonium (Cassuto et al. 1982b). Stroking the mucosa also causes 5-HT release and activates secretomotor reflexes in preparations of human jejunum, consisting of mucosa plus submucosa, but lacking the myenteric plexus (Kellum et al. 1999). Pharmacological analysis showed that the reflex depended on the activation of 5-HT4 receptors. 5-HT receptors as therapeutic targets are considered in Chapter 7. Roles of interstitial cells of Cajal in neuromuscular transmission Interstitial cells of Cajal (ICC; see Chapter 5) are functionally interposed between the terminals of motor neurons and the muscle (Ward & Sanders 2001), a suggestion that was originally made by Cajal on purely morphological grounds (Cajal 1911). This view was held by many microscopists whose work has been comprehensively reviewed by Thuneberg (1982). Huizinga et al. (1990) reported that electrical stimulation of inhibitory motor neurons elicited inhibitory junction potentials of reduced amplitude when the ICC of
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the myenteric plexus (ICCMY) and deep muscular plexus (ICCDMP) were both dissected away. The absence of ICC was confirmed histologically. Similar results were obtained by comparison of inhibitory and excitatory transmission in the stomach of normal mice and W/WV mutants that lack ICC in the muscle (Burns et al. 1996, Ward et al. 2000). In W/WV mice, the component of inhibitory transmission that is mediated via NO was lost. Interestingly, cholinergic transmission could be partly restored in the mutant mice by blocking ACh metabolism, by inhibiting ACh esterase (Ward et al. 2000), implying that under these circumstances ACh could diffuse to and act directly on the muscle. In a study in which ICCMY and ICCDMP were both disrupted by treating neonatal rats with antibodies to Kit, excitatory and inhibitory transmission to the circular muscle of the ileum were impaired, and neural transmission to colonic circular muscle was abolished (Torihashi et al. 1995). Consistent with the ICC receiving direct innervation from enteric motor neurons, they have receptors for the inhibitory transmitters NO (Shuttleworth et al. 1993, Young et al. 1993), VIP (Epperson et al. 2000), and ATP (Burnstock & Lavin 2002) and for the excitatory transmitters ACh (Epperson et al. 2000) and tachykinins (Portbury et al. 1996b). Transmitters of secretomotor and vasodilator neurons Secretomotor transmission to the mucosa has both cholinergic and noncholinergic components (Cooke & Reddix 1994). The cholinergic component is blocked by muscarinic receptor antagonists (Keast et al. 1985c) and the primary transmitter of the non-cholinergic secretomotor effect is probably VIP (Jodal & Lundgren 1989, Cooke & Reddix 1994, Reddix et al. 1994). When secretomotor reflexes in the cat small intestine were provoked by cholera toxin, VIP was released and the release was completely blocked by tetrodotoxin (Cassuto et al. 1981b). Moreover, secretomotor reflex responses that were evoked by intraluminal application of cholera toxin in the rat jejunum were abolished by a VIP receptor antagonist (Mourad & Nassar 2000). Rotavirus-induced secretion is also blocked by VIP receptor antagonism (Kordasti et al. 2004). In humans, the copious secretion caused by cholera toxin is not reduced by atropine, but it is mimicked by VIP or by VIP-secreting tumors (Kane et al. 1983). Thus, if an enteric reflex is involved in cholera toxin’s action in humans as in other mammals, the final secretomotor neuron is likely to be a VIP neuron. If a final cholinergic neuron also participates it would seem that its effect is overshadowed by that of VIP. Distension-evoked secretomotor reflexes in the rat colon were also substantially reduced when the actions of VIP were curtailed by desensitization of VIP receptors by prolonged exposure to the peptide, by inactivation of VIP using anti-VIP antibodies, and by a VIP receptor antagonist (Schulzke et al. 1995). About 80–90% of the
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secretory response was attributed to VIP. In experiments on guinea-pig colon, it was found that a VIP receptor antagonist reduced the response to electrical stimulation of secretomotor neurons (Reddix et al. 1994). Pye Smith and co-workers in 1874 (Pye Smith et al. 1874) reported that magnesium sulfate applied to the mucosa of the feline small intestine evoked a copious secretion, through what the authors assumed to be an enteric reflex, but that this secretion was not blocked by atropine. The receptor type that is activated by VIP in human and rat intestine is the VPAC1 receptor (Banks et al. 2005). Experiments in which single neurons were stimulated, and the resulting changes in the diameter of submucosal blood vessels were measured, provide direct evidence for the presence of both cholinergic and non-cholinergic vasodilator neurons (Neild et al. 1990, Vanner & Surprenant 1991, 1996, Kotecha & Neild 1995). In a series of experiments in which stimulating electrodes were placed on individual ganglia and both vasodilation and hyperpolarization of arteriolar smooth muscle was recorded, dilation was blocked by muscarinic antagonists in 48% of cases, was partly blocked in 31%, and was unaffected in 21% (Kotecha & Neild 1995). The observations suggest that the stimuli activated individual or a small number of vasodilator neurons. Thus some vasodilator neurons are purely cholinergic and some are noncholinergic. A component of non-cholinergic vasodilation was not accompanied by membrane hyperpolarization and was mimicked by VIP (Kotecha & Neild 1995). Long vasodilator reflexes that pass via the myenteric plexus appear to impinge primarily on the cholinergic secretomotor/vasodilator neurons (Reed & Vanner 2003). Immunohistochemical studies, combined with denervation, confirm that there are intrinsic cholinergic and non-cholinergic neurons innervating submucosal arterioles (Li et al. 1998). The cholinergic neurons are immunoreactive for ChAT and calretinin and the non-cholinergic neurons are immunoreactive for VIP. It is thus probable that a primary transmitter for non-cholinergic vasodilator transmission is VIP (Eklund et al. 1980, Furness & Costa 1987, Jodal & Lundgren 1989, Blank et al. 1990, Kotecha & Neild 1995, Vanner & Surprenant 1996, Li et al. 1998). There is a cholinergic component of transmission from secretomotor neurons in human (Morris & Turnberg 1980), but as the human mucosa is richly innervated by VIP-immunoreactive axons (Ferri et al. 1982a), it seems likely that there is also a VIP component. VIP has potent secretory effects in the human intestine (Krejs 1982), and the production of VIP by endocrine tumors is the acknowledged cause of the hypersecretion that occurs in the Verner-Morrison (watery diarrhea, hypokalemia, achlorhydria: WDHA) syndrome (Modlin et al. 1980). The relative contribution of cholinergic and non-cholinergic secretomotor neurons might differ between species. For example, secretomotor responses to distension in dogs are antagonized by atropine (Caren et al. 1974).
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Synapses in secretomotor and vasodilator pathways Secretomotor reflexes involve direct IPAN-to-secretomotor synaptic connections (Chapter 3) as well as connections through interneurons. In both cases the primary transmission is cholinergic at nicotinic synapses. The IPANs are cholinergic and when they are activated by physiological stimuli they cause fast EPSPs in other submucosal neurons that are blocked by nicotinic receptor antagonists (Pan & Gershon 2000), and both secretomotor and vasodilator reflexes are inhibited by nicotinic receptor block with hexamethonium (Caren et al. 1974, Cassuto et al. 1982b, Reed & Vanner 2003). Transmitters of motor neurons innervating gastrin cells Gastrin-releasing peptide (GRP, mammalian bombesin) is a transmitter from motor neurons to gastrin-containing endocrine cells of the antral mucosa. Nerve fibers that innervate the cells are immunoreactive for GRP (Holst et al. 1987, Miller et al. 1989, Sjövall et al. 1990), and GRP, or its amphibian equivalent bombesin, are powerful stimulants of gastrin release. Gastrin release that is caused by stimulation of the vagus is abolished by desensitization of receptors for GRP (Holst et al. 1987). Partly digested protein in the gastric lumen initiates an enteric reflex that results in the release of gastrin from the endocrine cells, and this release is reduced by atropine and by a GRP receptor antagonist (Schubert et al. 1992). Together, atropine plus the GRP receptor antagonist abolish the reflex release of gastrin. Analysis of chemical codes indicates that the neurons that innervate the gastric mucosa can be subdivided into two groups, cholinergic (ChAT-immunoreactive) and NOS-immunoreactive populations (Pfannkuche et al. 1998). However, the NOS neurons innervate only the base of the mucosa, adjacent to the muscularis mucosae (Furness et al. 1994). Therefore, the neurons innervating the gastrin cells are likely to be cholinergic, that is, they release both ACh and GRP as co-transmitters, consistent with the pharmacological analysis of Schubert et al. (1992). Summary and conclusions A feature of the enteric nervous system is that its neurons utilize multiple transmitters. Each enteric neuron has a primary transmitter, and may also have secondary transmitters that act on the same post-synaptic cells as the primary transmitter, and modulatory transmitters that act presynaptically. The primary transmitters are conserved between mammalian species, and probably between all vertebrate species. The primary transmitter of excitatory motor neurons to gastrointestinal smooth muscle is acetylcholine (ACh), which acts through muscarinic
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receptors on the muscle and interstitial cells of Cajal. Tachykinins are secondary transmitters of excitatory muscle motor neurons. They act at NK1 and NK2 receptors. ACh, GABA, and opioid peptides are modulatory transmitters of these excitatory motor neurons. Each of these modulators inhibits transmitter release. The primary transmitter of inhibitory motor neurons to gastrointestinal smooth muscle is nitric oxide, and secondary transmitters are VIP and PACAP, which both act at the same (VPAC) receptors, ATP, and possibly carbon monoxide. Transmission to the muscle is mainly indirect, via interstitial cells of Cajal (ICC). The primary transmitters of secretomotor neurons that control fluid secretion in the small and large intestines are VIP (with PACAP) and ACh. These transmitters also act at connections between secretomotor/vasodilator neurons and intrinsic arterioles in the gastrointestinal tract. Secretomotor neurons that cause acid release in the stomach have ACh as their primary transmitter. Motor neurons to the gastrin cells of the stomach utilize as the primary transmitter the peptide gastrin-releasing peptide (GRP) and as a secondary transmitter ACh. GRP is also a transmitter to pepsinogen-secreting cells of the gastric corpus (see Chapter 6). The major transmitter for fast excitatory transmission at synapses between enteric neurons is ACh, which impinges on post-synaptic nicotinic receptors. Other fast transmitters are ATP (at P2X receptors) and 5-HT (at 5-HT3 receptors). Slow excitatory transmission to enteric neurons is mediated through tachykinins (at NK1 and NK3 receptors), CGRP, ATP (at P2Y receptors), 5-HT, and by other transmitters that are yet to be identified. The transmitters of slow inhibitory transmission to enteric neurons include norepinephrine, somatostatin and possibly 5-HT. Presynaptic receptors at which the release of transmitter is modulated also occur at synapses on enteric neurons; these include muscrinic and 5-HT1A receptors through which transmission is reduced and 5-HT4 receptors, through which it is enhanced. The activation of the mucosal endings of IPANs is indirect. The primary transmitter, which is released from entero-endocrine cells, is 5-HT, and the receptor types on the axons of IPANs are 5-HT3, 5-HT4, and 5-HT1P receptors.
5: Neural control of motility
The purpose of this chapter is to draw together information that can be used to explain the motility patterns that are observed in vivo in the context of what is known of the organization of enteric nerve circuits and the physiology of their constituent neurons, which are analyzed in Chapters 1 to 4. The chapter also includes an analysis of the physiological roles of the sympathetic innervation of the gastrointestinal tract in inhibiting gastrointestinal motility. It is first necessary to describe the rhythmic oscillations in excitability of the muscle on which enteric neurons act. Rhythmic activity of gastrointestinal muscle The muscle of the gastrointestinal tract exhibits rhythmic, regular contractions. The contractions are observed in isolated tissue, and are therefore not triggered by circulating hormones, and are not prevented by atropine, which suggests that they are not due to the activity of excitatory neurons (Magnus 1904a). They are also unaffected by block of nicotinic receptors (Thomas & Kuntz 1926). However, as is pointed out later, atropine and nicotinic receptor block substantially reduces contraction amplitudes in the small intestine, because the intensity, but not the presence, of rhythmic activity is controlled by enteric neurons. Many years after the studies of Magnus and Thomas, the persistence of the rhythmic activity in the presence of tetrodotoxin confirmed the independence of the oscillation from neural activity (Liu et al. 1969). The rhythmic contractions of the muscle are the consequence of rhythmic changes in its electrical activity, first reported by Alvarez and Mahoney (1922), who showed the correspondence between the depolarizing electrical potentials and contractions in the stomach (Fig. 5.1). The depolarizing potentials are commonly called slow waves, although other terms have also been used. In later studies, the shapes of the slow waves were deduced by integrating extracellular records (Bozler 1945). Bozler also found that the waves were present even when there was no contractile activity and that the magnitudes of the waves were directly related to the strength of the 132
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Fig. 5.1 Contractile and electrical activity of the distal stomach. A, B: The correspondence between contractions of the antrum (A) and electrical slow waves recorded simultaneously with surface electrodes (B) by Alvarez and Mahoney (1922). This was the first demonstration of the close temporal relationship between contractile and electrical activity in the stomach. C: Intracellular recording of gastric slow waves. This record shows long-lasting depolarizations with superimposed action potentials. Reproduced from El Sharkawy et al. (1978).
contractions. However, it was not until 1961 that slow waves were recorded with intracellular microelectrodes (Bortoff 1961, Bortoff & Weg 1965). These and subsequent studies revealed prolonged depolarizing potentials (Fig. 5.1). Action potentials were often superimposed on these depolarizations. Slow waves, and the corresponding rhythmic contractions, occur at about 3 events per min in the human stomach, and 11–12 per min in the duodenum, with a decline in frequency to about 8 per min in the distal ileum. In the dog ileum, the slow wave-associated rhythmic contractions occur at about 10–14 per min (Bayliss & Starling 1899, Alvarez & Mahoney 1922, Ehrlein et al. 1987) (see Fig. 3.1). It is interesting that rhythmic contractions at this frequency were recorded from the intestine of a living dog by Legros and Onimus in 1869; a record from their work is reproduced in Furness and Costa (1987). In smaller animals, greater frequencies are encountered; the slow waves and slow wave-entrained contractions occur at 30 per min in the rat small intestine (Ruckebusch & Fioramonti 1975, Scott & Summers 1976) and at 25 per min in the guinea-pig (Galligan et al. 1985). The mechanism of generation of the slow waves was intensely investigated from the late 1970s, but the problem was not resolved until the roles of the
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interstitial cells of Cajal (ICC) in their generation was discovered in the early 1990s, although generation of the slow waves by ICC had been suggested earlier, based on careful analysis of their structures and location (Thuneberg 1982). It had been realized that the slow waves were a property of the whole tissue. Unlike isolated cardiac muscle cells that generate cardiac action potentials, single smooth muscle cells from the gastrointestinal tract do not have an intrinsic ability to generate rhythmic electrical slow waves (Farrugia 1999). Moreover, slow waves can be recorded from the full thickness of the isolated circular muscle of the small or large intestine but, in many species, if the inner part is removed the remaining muscle does not exhibit slow waves or phasic contractions (Durdle et al. 1983, Hara et al. 1986, Smith et al. 1987). Suzuki et al. (1986) prepared strips of muscle from the cat small intestine with and without ICC, whose presence was determined by methylene blue staining. Slow waves were recorded from preparations with ICC present, but not from preparations where they were absent. Subsequent to these investigations, recordings made from isolated ICC showed that, unlike intestinal smooth muscle cells, they were capable of generating spontaneous depolarizing events (Langton et al. 1989, Tokutomi et al. 1995, Thomsen et al. 1998). The suspicion that ICC were the generators of slow waves was greatly strengthened when it was discovered that intestinal muscle from mutant mice that lack subpopulations of ICC did not generate slow waves (Ward et al. 1994, Huizinga et al. 1995). These mice were deficient in the gene for c-kit, a protein that is present in ICC and that appears to be essential for their development. An alternative approach to inactivating ICC is to incubate them with neutralizing antibodies to c-kit. When this was done, preparations of the murine stomach lost their ability to generate slow waves (Ördög et al. 1999). The previous discussion covers one of the roles of ICC, the generation of slow waves in the muscle. These cells, or subgroups of them, are also intermediaries in transmission from enteric neurons to the muscle (Chapter 4), they probably contribute to co-ordination of the electrical activity of adjacent muscle regions (Horiguchi et al. 2001), and they may have a role in detecting mechanical forces in the muscle (Fox et al. 2002). Structure and properties of interstitial cells of Cajal Interstitial cells of Cajal (ICC) were identified histologically by Cajal and several other authors by the use of methylene blue and silver staining in the 1890s (Cajal 1892, 1893, La Villa 1898, Dogiel 1899). Dogiel (1899) seems to be the first to call them cells of Cajal. ICC have small cell bodies and several elongated processes (Fig. 5.2). Cajal (1911) surmised that they are neuron-like cells that are interposed between nerve fibers and the muscle, because, like neurons, they are preferentially stained by methylene blue and
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Fig. 5.2 Interstitial cells of Cajal. A: Drawing by La Villa (1898) of interstitial cells in the deep
muscular plexus of the small intestine of the guinea-pig that had been stained with methylene blue. B: Drawing by Taxi (1965), also of intestitial cells in the deep muscular plexus of the guinea-pig small intestine. C: Interstitial cells at the level of the myenteric plexus of the rabbit small intestine are depicted in a preparation stained with a Bielschowsky-Gros silver method (reproduced from Richardson 1958). D: Interstitial cells in the deep muscular plexus of the guinea-pig small intestine, stained with Champy’s zinc iodide-osmium method. Calibration: 10 μm.
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silver stains. They are also stained by another method that stains neurons, the Champy zinc iodide-osmium method (Taxi 1965, Stach 1972). An excellent monograph that reviews the early light microscope descriptions of ICC has been published by Thuneberg (1982). In it he points out that similar cells appear to have been described by several authors prior to Cajal (1892, 1893). However, it was Cajal who recognized the special nature of these cells. The interstitial cells have been subclassified by the regions in which they are found, with different authors adopting slightly different terminology. Three groups of ICC appear to be the most important for regulating gastrointestinal motility – those that are found at the level of the myenteric plexus (ICCMY), those within the circular muscle layer (intramuscular ICC, ICCIM), and those near the inner surface of the circular muscle (ICCDMP in the small intestine and ICCSM in the colon; DMP standing for deep muscular plexus, see Chapter 1, and SM for submucosa, which they abut). As soon as electron microscope methods were used to examine ICC it was apparent that they are not neurons, in particular they do not contain transmitter vesicles and the cytoplasmic components of their cell bodies do not resemble those of neuron cell bodies (Richardson 1958, Taxi 1959). Subsequent to these studies there have been numerous investigations of the ultrastructure of ICC, which have provided slightly variant descriptions (Komuro 1999). The uncertainty may have been contributed to by the difficulty of unequivocally identifying the ICC, for example in distinguishing them from fibroblasts that have some similarity of shape. However, with the advent of improved techniques to locate the cells and of cell markers such as c-kit, a consistency in descriptions of the ICC has emerged. ICCDMP and ICCSM exhibit large numbers of cysternae beneath the surface membrane, an abundant endoplasmic reticulum associated with the cysternae, frequent mitochondria and numerous intermediate filaments (Rumessen et al. 1982, Berezin et al. 1988, Komuro et al. 1999). These cells form gap junctions with each other and with smooth muscle cells; the presence of these junctions provides structural evidence of the electrical coupling between these cells that is so apparent from physiological investigations. ICCDMP and ICCSM are also closely approached by nerve fiber varicosities containing synaptic vesicles, and they exhibit receptors for neurotransmitters (Chapter 4). ICCMY also contain numerous mitochondria and smooth endoplasmic reticulum, but caveoli are not a consistent feature (Horiguchi & Komuro 1998, Komuro 1999). These ICC also connect to smooth muscle and to each other through gap junctions. However, they are less densely innervated than ICCDMP and ICCSM. ICCIM contain numerous mitochondria, intermediate filaments and sub-surface caveoli. They form gap junctions with ICC and smooth muscle cells and have close approaches from nerve fibers (Komuro 1999).
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The relationships between electrical events in ICC and in smooth muscle cells have been studied in the guinea-pig stomach by simultaneous intracellular recording from ICC and the muscle cells (Dickens et al. 1999, Hirst & Edwards 2001, Hirst et al. 2002, Cousins et al. 2003). Cell identity was confirmed by intracellular injection of a marker, neurobiotin. Intracellular recordings from gastric ICCMY in acutely isolated tissue with smooth muscle cells also present reveal pacemaker potentials with an initial depolarization lasting about 2 s, followed immediately by a plateau potential lasting about 8 s (Dickens et al. 1999, Hirst & Edwards 2001) (Fig. 5.3). Slow waves occur in the smooth muscle a fraction of a second after the pacemaker potentials. The slow waves in the muscle have two components – an electrotonically conducted depolarization that is superimposed by an active (regenerative) potential. In the mouse small intestine, it is also the ICCMY that are responsible for driving slow wave activity in the muscle; in the c-kit mutants, in which slow waves are abolished, ICCMY are depleted, but ICCDMP are still present. However, in other species, ICC at the inner surface of the small intestine appear to be the generators of slow waves (Hara et al. 1986). In the dog colon, ICCSM appear to be the major source of slow waves (Smith et al. 1987). As has already been discussed in Chapter 4, ICC are principal targets of excitatory and inhibitory motor neurons that control the motility of the gastrointestinal tract.
Fig. 5.3 Simultaneous intracellular recordings from an ICCMY and a smooth muscle cell of the circular muscle layer of guinea-pig gastric antrum. The onset of the pacemaker potential precedes the onset of depolarization in the circular muscle layer. Each pacemaker potential consists of two components – an initial component followed by a plateau component. Furthermore, each slow wave consists of two components – an initial passive component and a secondary regenerative component which is superimposed. Reproduced from Hirst and Ward (2003).
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Relationship between slow wave activity and neural control The slow wave rhythm dominates the pattern of movement of the body and antrum of the stomach (Chapter 3), whereas in the fundus, which acts to regulate gastric volume, the underlying slow waves have less importance. Electrical recordings from the intestine, in vivo, reveal the continuous generation of slow waves (Fig. 5.4). However, the slow waves do not necessarily initiate contractions, and the intensities of the contractions are strongly modulated by enteric neurons. Each small region of the intestine generates slow waves at its own intrinsic frequency and, because the successive regions are electrically coupled to each other, the occurrence of slow waves in one short region influences the occurrence of slow waves in the adjacent region, with regions of higher frequency tending to drive those of lower frequency (Diamant & Bortoff 1969). Because there is a gradient of frequency along the small intestine from the duodenum to the terminal ileum, there is a conduction of slow waves from oral to anal. Thus, when slow waves are raised above threshold by nerve activity in the small intestine they tend to push the contents in an anal direction. When records are taken along the intestine it is found that more proximal regions drive more distal areas at the intrinsic frequency of the proximal region, until the disparity in driving frequency and intrinsic frequency is too great for the distal region to follow (Fig. 5.4). At this point there is a region of irregular activity, distal to which regular activity at a lower frequency is established
Fig. 5.4 Slow waves of electrical activity in the muscle of the cat small intestine, recorded in vivo. Eight records taken with electrodes at 2 cm intervals in the cat duodenum are shown, the upper trace being from the most oral electrode. The slow waves progress from one site to the next, as shown by the dashed lines. At some points (fourth record) there is waxing and waning of the waves and some of them fail to propagate. When this happens, there is a decrease in the frequency of the slow waves, as shown by the changes in separation of the dashed lines. Reproduced from Diamant and Bortoff (1969).
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(Diamant & Bortoff 1969). The entrainment of slow waves in regions of the intestine, separated by regions of irregular activity, leads to the formation of frequency plateaus, between which frequency drops (Diamant & Bortoff 1969). Although the frequency is apparently the same within the frequency plateau, the slow waves do progress in the anal direction within this region. Slow waves last 2–5 s, vary in amplitude up to about 20 mV, occur at frequencies of 3 to 40 per min, depending on region and species, and travel at speeds of about 1–15 cm/s (Table 5.1). Slow waves only cause physiologically significant contractions in the small intestine when they are of sufficient amplitude to generate action potentials (Daniel et al. 1959, Bass et al. 1961, Hara et al. 1986). In general, slow waves without action potentials cause very small changes in tension, and the contractions of the muscle are graded according the number of action potentials (Fig. 5.5). The small intestine requires the activity of enteric excitatory motor neurons for the slow waves to be brought to threshold for significant contraction. Thus block of reflexes that activate the excitatory neurons with nicotinic receptor blockers such as hexamethonium, or block of transmission from the excitatory neurons to the muscle with muscarinic receptor antagonists, such as atropine or scopolamine, essentially paralyzes the small intestine (Quigley et al. 1934, Youmans et al. 1943, Reinke et al. 1967) although, as pointed out earlier, small oscillating contractions are still seen. It is worth remembering at this point that gastric slow waves do not require neural activity to reach threshold to generate propulsive contractions (Chapter 3 and below), although their amplitudes are clearly influenced by neural activity.
Table 5.1 Properties of slow waves
Species
Region
Frequency (cycles/min)
Apparent speed of propagation from oral to anal (cm/s)
Human
Stomach Duodenum Terminal ileum Transverse colon Stomach Duodenum Terminal ileum Mid-colon Duodenum Jejunum Terminal ileum Colon Small intestine Duodenum
3 11–13 8–10 2–6 & 6–13 5 18–19 10–12 5–8 & 18 18 18–19 13 4.5–6 25 36–40
12
Dog
Cat
Guinea-pig Rat
14–17 0.5–3
4 3.5–5 0.5
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Fig. 5.5 A, B: Spontaneous electrical and contractile activity recorded from a cell located in the inner circular muscle layer of a whole-thickness preparation of the human jejunum. Part of the record in A is shown on a greater timescale in B. The records show that the amplitudes of contractions (upper) are related to the numbers of action potentials that are triggered by the slow waves (lower trace) and that slow waves that do not generate action potentials cause very small or no contractions. Reproduced from Hara et al. (1986).
Gastric motility Motility is discussed for the stomachs of monogastric species such as human and dog. The stomach has a reservoir function; it increases volume as it fills, and even relaxes prior to food arriving. It also has a function to mix the food with gastric juices and to push the liquefied products of gastric digestion into the duodenum. These functions are often considered to be related to regions, the fundus (proximal stomach) being primarily associated with the gastric reservoir function and the corpus and antrum (distal stomach) being associated with gastric mixing (Kelly 1981). This is a useful convention, but when the stomach fills the body (or corpus, the two terms are interchangeable) both increases in volume and generates contractile waves. Gastric relaxation that is necessary for this volume accommodation is mediated through enteric inhibitory muscle motor neurons (see Table 2.1). Mixing is associated with gastric peristaltic contractions that push the contents against the pyloric sphincter, which lets a small amount of liquid enter the duodenum with each contraction, while solid material is retained in the stomach. A basis for the distinct regional difference is that slow waves do not occur in the fundus, but occur in the oral part of the corpus from where they are conducted to the distal antrum (Kelly et al. 1969, Hinder & Kelly 1977, El Sharkawy et al. 1978). Gastric reservoir function (principally the proximal stomach) Cannon (1898) observed by cine-radiography that the volume of the fundic region of the stomach increased when the stomach was filled and reduced when it was less full. Changes in the gastric corpus are less pronounced, and
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the antral volume is unchanged. Moreover, the pressure in the stomach does not increase or increases very little as it is filled (Kelling 1903), implying that the muscle of the proximal stomach relaxes to accommodate the meal. That this is the case was shown by Cannon and Lieb (1911) who demonstrated that relaxation occurred even before the food arrived, a phenomenon called receptive relaxation by these authors (Fig. 5.6). The relaxation occurs when the pharynx or esophagus is distended and can be observed even when the esophagus is severed and no food reaches the stomach (Abrahamsson & Jansson 1969). The reflex is prevented if the vagus nerves are cut. Relaxation of the proximal stomach also occurs if the gastric volume is increased, for example by distension by inflation of an intragastric balloon. As balloon volume is increased from 1 to 300 ml in a dog, intragastric pressure rises only slightly, and remains constant when the balloon volume is further increased to 800 ml (Kelly 1981). This accommodation reflex is substantially reduced after vagotomy in cat, dog, and human (Abrahamsson & Jansson 1973, Wilbur & Kelly 1973). A vagally mediated gastro-gastric reflex relaxation can also be elicited if distension is confined to the antrum (Abrahamsson 1973). In addition, there is a small residual component of accommodation that is due to an intrinsic reflex (Andrews & Bingham 1990). The existence of an intrinsic accommodation reflex is confirmed by studies of the isolated stomach in vitro (Hennig et al. 1997). The final inhibitory neurons in these pathways are not adrenergic (Jansson 1969b); they are intrinsic (enteric) inhibitory motor neurons that utilize co-transmitters, including NO, ATP, and VIP (Chapter 4).
Fig. 5.6 Recording of the receptive relaxation of the stomach made in an unanesthetized cat.
The cat was eating minced meat and made swallows at the times indicated by the arrows. It takes 10–11 s for the food to traverse the esophagus and yet gastric relaxation occurred within about 2 s of the first swallow. The relaxation persisted as long as the cat was swallowing food. The asterisks mark artifacts caused by the cat’s movements. Reproduced from Cannon and Lieb (1911).
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The main transmitter mediating relaxation of the gastric fundus is NO (Desai et al. 1991, Hennig et al. 1997, Tack et al. 2002). As the volume in the stomach reduces, the fundus contracts. This also appears to be a vagally mediated effect (Wilbur & Kelly 1973). Thus the fundus adjusts its volume both by relaxation and contraction. Through this mechanism, the fundus is thought to exert slight pressure on the contents, so that the gastric reservoir always pushes the contents towards the distal stomach where the work of digestion occurs. Gastric peristalsis and mixing (the distal stomach: corpus and antrum) It was pointed out in Chapter 3 that gastric peristalsis, which occurs in the body and antrum, is not prevented when the myenteric plexus is cut through or nicotine is given in a dose that blocks peristalsis in the intestine. Moreover, the frequency of peristalsis corresponds to the frequency of gastric slow waves which trigger action potentials and contractions (Fig. 5.1). Contractions are often undetectable when gastric slow waves fail to elicit action potentials (Kelly et al. 1969). Although more refined analysis in vitro shows that Ca2+ entry during the plateau phase of the gastric slow wave can contribute to contraction (Sanders & Publicover 1989), the presence of action potentials is nevertheless a good indicator that the slow waves have reached threshold to cause physiologically significant contractions. Thus, gastric peristalsis is generated by the slow waves and, unlike peristalsis in the small intestine and colon, it does not necessarily require augmentation by the actions of excitatory neurons to be observed. Nevertheless, there is good evidence that neurons modulate the strength of contractions caused by gastric slow waves, and that excitatory nerve activity is in fact necessary for full amplitude contractions (Andrews et al. 1980). Peristaltic contractions, and the slow waves that underlie them, commence in the oral part of the gastric corpus and progress towards the pyloric canal (Cannon 1898, Kelly et al. 1969) (Fig. 5.7). In humans they occur at 3/min (Code et al. 1952) and travel at about 0.3 cm/s in the proximal antrum and 0.6 cm/s in the distal antrum (Hinder & Kelly 1977). The frequency of waves is similar in different species, in dog about 4–5 per min, in cat 4–6 per min and in guinea-pig about 3–4 per min. The speed of propagation of gastric slow waves and peristalsis (the same thing) is slower than slow wave speeds in the intestine, but faster than nerve-mediated peristaltic waves or phase III of the MMC in the small intestine (Table 5.2). Unlike the peristaltic waves in the intestine, gastric slow waves do not occlude the lumen (Fig. 5.7). They compress the contents, and as they progress they push some of the fluid over the surface of the more solid contents and aspirates of gastric juice, containing small particles and dissolved components of the food, enter the duodenum
Dog duodenum (Bass et al. 1961) Dog jejunum (Schemann & Ehrlein 1986)* Rat small intestine (Ruckebusch & Fioramonti 1975) Human duodenum (Andrews et al. 2002)*
14–18 5.0 ± 1.9
Rabbit small intestine (Lind et al. 1991)
Human (Kellow et al. 1986)
0.01 (0.6 cm/min) 1
Dog (Szurszewski 1969)
Human (Kellow et al. 1986)
0.02–0.07 (1.3–4.3 cm/min) 0.02–0.3 (1.2–1.9 cm/min) (dog)
Dog (Szurszewski 1969) Dog (Schemann & Ehrlein 1986)
0.05–0.1 (3.5–6.2 cm/min)
Cat (Cannon 1902)
0.08–0.18 (4.5–11 cm/min)
Dog (Ehrlein et al. 1987)
0.07–0.12 (4–8 cm/min) 0.03–0.08 cm/sec (2–5 cm/min)
Enteric nervous system
Enteric nervous system
Enteric nervous system
Enteric nervous system
ICC coupled to smooth muscle
ICC coupled to smooth muscle
Generator
*Contractions were recorded. They occurred at 11–12 per min and I have assumed that these are the mechanical manifestations of slow wave activity.
Peristaltic rush/giant migrating complex
Phase III of the MMC (distal half of small intestine)
Phase III of the MMC (proximal half of small intestine)
Intestinal peristalsis
1.7–2.8
1–2
Cat small intestine (Bortoff et al. 1984)
Intestinal slow waves
Human (Hinder & Kelly 1977)
0.3 (corpus) 0.6 (antrum) 3–5
Gastric peristaltic waves
Species/Reference
Speed of conduction (cm/s)
Propagated event
tine, slow waves travel about 10–50 times faster than propagated events that are driven by the enteric nerve circuits
Table 5.2 The relative speeds of conduction of contractile events (and their underlying electrical counterparts) in the stomach and small intestine. In the small intes-
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Fig. 5.7 The movements of the cat stomach, recorded by X-ray cinematography in vivo. A: X-
ray image during gastric digestion. Barium sulfate, to provide contrast, was added to a meal of meat. Contractions, marked 1–4, can be seen. These begin in the gastric body and move to the antrum, but they do not occlude the lumen. The asterisk indicates an aspirate of gastric juice entering the duodenum. The white dots are images of small metal balls sewn to the stomach to indicate its outline. Reproduced from Gianturco (1934). B: Drawing of the X-ray image of the stomach of an unanesthetized cat, during digestion. Arrows indicate the peristaltic contractions of the body and antrum. Reproduced from Cannon (1911).
(Fig. 5.7). At any instance in time, 3–4 peristaltic waves can be observed in the stomach. When the stomach is empty there is little evidence of gastric peristalsis (e.g. Fig. 5.8), but once it contains food the peristaltic waves are manifest and continue to be generated at more or less the same frequency until the stomach is empty (Cannon 1911). In the empty stomach, only about 25% of slow waves exhibit action potentials and cause contractions in the distal stomach, but when the stomach is distended practically all slow waves have action potentials and evoke contractions (Kelly et al. 1969). Because the proximal
Fig. 5.8 Pressure waves recorded from the human stomach. The records show the stomach when peristaltic activity is weak, moderate and strong. Note that the frequency of contractions does not change; it remains at the frequency of the slow waves. Reproduced from Code et al. (1952).
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stomach relaxes during filling, records of the volume of the whole stomach show enhanced contractions superimposed on an overall increase in gastric volume. The vagus nerves and reflex centers in the brainstem dominate control of gastric motility The augmentation of the gastric contractions when the stomach is artificially distended with fluid is almost entirely through vago-vagal reflexes (Andrews et al. 1980). When the antrum, or the whole stomach, is extrinsically denervated, antral peristaltic contractions are smaller and emptying times of solids are prolonged (Moore et al. 1946, Mroz & Kelly 1977, Andrews et al. 1980). The peristaltic waves become less intense, and less effective in pushing liquid past the pylorus (McSwiney 1931, Wilbur & Kelly 1973), although with time after vagotomy there is some restoration of contraction amplitude (Andrews & Bingham 1990). The strengths of the antral contractions are sequentially reduced when the vagal branches entering the antrum are successively cut, from proximal to distal (Daniel & Sarna 1976). Vagotomy was used in many thousands of patients to treat peptic ulcer before drug therapies were developed, and in most of these patients the pyloric sphincter was slit to relieve the gastric stasis that ensued (Seymour & Andersen 1999). In the earliest operations, performed in 1943 and 1944, the pyloric sphincter was not slit and the patients suffered from gastric retention (Dragstedt & Schafer 1945). As detailed above, the reservoir function of the stomach is also controlled through the vagus. Intracellular microelectrode recordings from individual gastric neurons indicate that the majority, at least two-thirds, of gastric neurons receive direct cholinergic excitatory synaptic inputs from the vagus (Schemann & Grundy 1992). These experiments were done by stimulating a vagal branch connected to an isolated region of gastric corpus, and it is possible that not all inputs to each neuron were retained or effectively stimulated, so the data might underestimate the numbers of neurons receiving excitatory synapses from vagal axons. Morphological studies also indicate that the majority of gastric neurons receive vagal input, and even suggest that the vagal inputs outnumber those that arise from intragastric neurons (Lawrentjew 1931, Filogamo & Gabella 1970, Holst et al. 1997). Some of these vagal fibers are presumed to directly innervate the inhibitory muscle motor neurons, which are about onethird of the neurons in the corpus, and the excitatory muscle motor neurons, which are about 40% of the neurons (Grundy & Schemann 2002). Nevertheless, a number of studies indicate that there is intrinsic activity of excitatory cholinergic neurons, even in the completely isolated stomach. Intracellular microelectrodes have demonstrated the spontaneous occurrence
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of fast EPSPs in some enteric neurons in the isolated stomach (Schemann & Wood 1989b), and other investigations have demonstrated an excitatory tone that is reduced by tetrodotoxin, or by antagonists of muscarinic or nicotinic receptors (Beani et al. 1971, Meulemans et al. 1993, Hennig et al. 1997). Hennig’s experiments show that the amplitudes, but not the frequencies of occurrence, of contractile waves are reduced when transmission from excitatory neurons to the muscle is prevented by tetrodotoxin. The effectiveness of the excitatory neurons is enhanced when the stomach is distended (Hennig et al. 1997), presumably because their rates of firing are increased. Although release of excitatory transmitter, in the absence of vagal input, enhances gastric slow waves, there is little evidence for an intrinsic reflex that is organized like that in the small intestine and is necessary for gastric peristalsis. After acute vagotomy, gastric distension causes very much weaker phasic contractions than are seen in the vagally innervated stomach (Andrews et al. 1980). The residual responses to distension are reduced by hexamethonium, indicating that there is a component of the enhancement of gastric peristaltic waves that is due to intrinsic reflexes. The conduction of gastric peristaltic waves is not affected by cutting through the myenteric plexus (Cannon 1912), which is unlike the effect on the peristaltic reflex in the small intestine and colon whose conduction is prevented by this procedure (Chapter 3). Furthermore, if the muscarinic receptor agonist carbachol is applied to the isolated stomach in which all nerve-mediated events have been prevented by tetrodotoxin, gastric peristaltic waves are restored (Hennig et al. 1997). This suggests that neuronal circuits are not required to co-ordinate peristaltic movement. Although reflex responses can be evoked in the stomach by local distension (Yuan et al. 1997), these responses are not as robust or clearly organized to promote propulsion in the stomach as are reflexes that can be elicited in the intestine. Distension of either the fundus or antrum caused inhibition in the corpus, and distension of the corpus caused inhibition in the fundus but no response in the antrum. Distension of the corpus did not affect the slow waves in the antrum. Only after pharmacological block of the inhibition was an ascending excitatory reflex (from the antrum to corpus) revealed, and this response was small, compared to excitation that could be evoked by electrical stimulation. There was no evidence for a descending excitatory reflex that might augment gastric peristalsis. IPANs, the types of neurons through which reflexes in the intestine are initiated, appear to be absent or very rare in the stomach. IPANs have AH electrophysiological characteristics and have Dogiel type II morphology (Chapter 2). AH neurons are completely absent from populations of neurons that have been sampled by electrophysiological recording in the corpus (Schemann & Wood 1989a) and are quite rare in the antrum (Tack & Wood 1992).
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Consistent with the paucity of AH neurons, previous morphological studies had shown that Dogiel type II neurons are not found in the gastric corpus and are rare in the antrum (Lawrentjew 1931). A small number of calbindin neurons in the antrum have a multipolar morphology, similar to IPANs of the small intestine (Reiche et al. 1999). These amount to about 0.5% of all neurons. These data fit with the lack of evidence from studies of the stomach for peristaltic reflexes of the type observed in the small intestine and colon. It is concluded that gastric peristalsis is a consequence of contractions that are induced in the muscle by slow waves that are themselves generated by the pacemaker activity of ICC. The slow waves are normally close to the threshold for generating contractions in the muscle, and some of them do so even when the stomach is empty. Food in the stomach evokes a vago-vagal reflex that activates enteric cholinergic excitatory motor neurons whose effect is to augment the slow waves and increase the amplitudes of peristaltic waves. A background activity of the motor neurons, which may be enhanced by activation of intrinsic reflexes when the stomach is distended, could add to the vagal excitatory effect. Another factor that is most likely involved in the normally behaving animal or person is a cephalic phase of augmentation of gastric activity, via the vagus, that occurs before or at the time of ingestion of food. Patterns of small intestine motility and their intrinsic neural control The migrating myoelectric complex In humans and in other mammals, notably carnivores, that take food intermittently, the activity of the small intestine has one pattern of activity in the unfed (interdigestive or fasted) state, and one that occurs following a meal. The fasted state is characterized by the periodic occurrence of a region of intense contractile activity, phase III of the migrating myoelectric complex (MMC), that travels slowly from the gastroduodenal junction to the end of the ileum. In contrast, the MMC occurs at all times in ruminants and other herbivores, which relates to the more or less continuous supply of ingestible material from the stomach(s) to the small intestine in these species (Ruckebusch & Bueno 1976, Ruckebusch & Pairet 1985). The first description of the MMC seems to be that of Boldyreff (Boldyreff 1905). However, its description by Szurszewski (1969), who refers to earlier studies of similar phenomena, led to a resurgence of interest. Szurszewski seems to have been the first to realize that the MMC traverses the full length of the small intestine and that it is a recurring event. The periodic activity that occurs during the fasted state was later divided into four phases – I, a quiescent phase, II, an
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Fig. 5.9 Manometric recordings of the different phases of the human migrating motor complex (MMC) as it progresses from the duodenum (D1, 2) to the jejunum (J1, 2, 3). The dashed lines show propagation of the leading edge of two phase III complexes. The four phases of the complex (I, II, III, and IV) are indicated. Reproduced from Soffer (1998).
irregular phase, III, the phase corresponding to the migrating electric complex described by Szurszewski, and IV, a brief cycle of irregular activity at the end of phase III (Fig. 5.9). Together, these four phases are considered to constitute the MMC (Carlson et al. 1972), although phase III alone is often meant by some authors when they refer to the MMC. Phase III is also called the activity front. In the discussion of the MMC, I have concentrated on data from dog and humans, species in which there are distinct fed and interdigestive patterns of activity. The MMC is not only a cycle of changes in contractile activity: in co-ordination with the changes in motility are periodic increases in gall bladder contraction, pancreato-biliary secretion and intestinal secretion (Read et al. 1977, Vantrappen et al. 1979, Keane et al. 1980, Sarr et al. 1980, Traynor et al. 1984). Phase III of the MMC consists of a slowly moving band of quickly moving contractions, that is, the rapidly conducted contractions sweep over and create the slower moving complex (Szurszewski 1969, Ehrlein et al. 1987, Andrews et al. 2002). Phase III of the MMC progresses in an anal direction because the rapidly conducted contractions, which are generated by the slow waves reaching a threshold to cause contraction, commence at points that are successively displaced more anally (Schemann & Ehrlein 1986). The contractions are sufficiently intense that they occlude the lumen of the intestine. To demonstrate the occlusive power of phase III of the MMC, liquid contrast medium has been injected into the lumen of the small intestine of unanesthetized dogs, in advance of the activity front. The contrast medium was completely
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Fig. 5.10 The migrating myoelectric complex (MMC) recorded with closely spaced electrodes
in the dog jejunum. The MMC progresses slowly along the intestine. It consists of superimposed fast-moving contractions (dashed lines), generated by the slow waves. The contractile waves are initiated at progressively more anal positions as the complex progresses anally. Reproduced from Ehrlein et al. (1987).
propelled by the phase III activity (Ehrlein et al. 1987). The same authors used closely spaced transducers to record the contractions that were evoked by individual slow waves at successive sites in the jejunum (Fig. 5.10). It can be seen that as the phase III activity slowly progresses along the intestine, the contractions rapidly traverse the activity front. These rapidly sweeping occlusive contractions, each commencing slightly more anal than the one before, create a slower moving event that is very effective in propelling the material in the lumen of the small intestine. This effect has been likened to a person sweeping a floor: as the sweeper moves steadily forward, the broom makes numerous quicker sweeps of the same region. This analogy has led some commentators to refer to the MMC as the intestinal housekeeper (Hasler 2003). The progression of the band of strong phase III motor activity that characterizes the MMC is dependent on the enteric nervous system. Neither vagotomy, nor sympathetic denervation, nor complete extrinsic denervation of the small intestine blocks the initiation or progress of the MMC (Marik & Code 1975, Buéno et al. 1979, Marlett & Code 1979, Aeberhard et al. 1980). The MMC even occurs when the jejunum and ileum are autotransplanted and the nerves are stripped from the arterial supply to the transplant (Sarr et al. 1989). However, it is blocked by systemic atropine or hexamethonium, implying that the activity of enteric muscle motor neurons and enteric interneurons are essential to its occurrence (Ormsbee et al. 1979, El Sharkawy et al. 1982). Sarna (1981) devised an ingenious experiment which confirmed the essential role of the enteric nervous system for the progression of the MMC. Cannulae were inserted into arteries of mesenteric arcades that supplied 3–5 cm long segments of canine small intestine. Drugs
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could be injected into the cannulae while the dogs were unanesthetized and the MMC was being recorded through implanted electrodes. The local infusion of tetrodotoxin just before the expected occurrence of phase III of the MMC abolished the phase III activity at the site of injection and for some distance more anal. However, a new migrating complex occurred more distally. Injection of atropine or hexamethonium into the cannulae also blocked the MMC. With hindsight, it is apparent that phase III of the MMC was recorded in the extrinsically denervated dog small intestine by Bayliss and Starling (1899), who described migrating contractile events that traveled at 6 cm/min. In the more recent work of Szurszewski (1969), also in dog, the speed of propagation of the MMC was given as 3.5–6.2 cm/min (average 5.3) in the proximal small intestine and 1.2–1.9 cm/min (average 1.5) in the distal small intestine. The speed of the MMC also changes along the intestine in humans, from 4.3 cm/min in the proximal jejunum to 1.3 cm/min in the upper ileum and 0.6 cm/min in the distal ileum (Kellow et al. 1986). It progesses at a similar speed (4 cm/min) in the ileum of a small mammal, the guinea-pig (Galligan et al. 1985), and at 2.5 to 10 cm/min in the rabbit (Ruckebusch & Pairet 1985). Migrating events, similar to the MMC and traveling at 6 cm/min, have also been recorded in the isolated small intestine (Perkins 1971). In humans, the four phases of the MMC run in a cycle that lasts 84–112 min (Hasler 2003). Phase I occupies 40–60% of the cycle, phase II, 20–30%, phase III about 5–10%, and phase IV is a brief transition period of irregular activity at the end of phase III. Although the contractile activity associated with the MMC and its local propagation is dependent on the enteric nervous system, the mechanism of initiation of the MMC is still unclear. The MMC commences in the human gastric antrum about one-third of the time, but it is often first observed in the duodenum (Kellow et al. 1986). Moreover, after intestinal transection phase III can commence distal to the transection, often out of phase with the intact, more proximal small intestine (Sarr & Kelly 1981). In humans, initiation at the gastro-duodenal junction appears to depend on hormones, notably motilin, which is the product of endocrine cells of the upper small intestine. Increases in the concentration of plasma motilin occur just before phase III activity in humans, but only when the MMC commences in the gastric antrum (Bormans et al. 1987). Motilin infusion induces a phase III-like event in humans (Vantrappen et al. 1979), although it initiates only those events that begin in the gastric antrum, and then often after considerable delay (Luiking et al. 2003). In dogs, the MMC is abolished for a period of several hours following the infusion of anti-motilin antiserum (Lee et al. 1983). Motilin induces phase III of the MMC in dogs, but does not do so in pigs (Buéno et al. 1982) or rabbits (Guerrero Lindner et al. 1996).
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The function of phase III of the MMC is to remove the epithelial cells that are continuously shed from the intestinal mucosa, the secretions that occur even in the interdigestive period, and undigested food from the small intestine (Szurszewski 1969, Hasler 2003). An index of the physiological need for such an event is the accumulation of bacteria that occurs in the small bowel within a few hours of impaired propulsion, along with associated malabsorption, mucosal injury, and development of adynamic ileus (Vantrappen et al. 1977, Summers 2003). During digestion, the stomach pushes liquid and very small particles into the duodenum. However, larger non-digestible particles (more than 1 mm in diameter) can be left. These are emptied into the duodenum by the gastric phase III contractions (Mroz & Kelly 1977). Activity during phase II of the MMC and in the fed state Feeding in human and carnivores converts the pattern of activity to the fed state, in which the phases of the MMC are replaced by irregular activity that in humans lasts for 1–2 h, before the MMC is once again observed. The patterns of movement of the small intestine during phase II of the MMC and in the fed state are similar. In both conditions, about 15% of contractile events are stationary (contractions that are confined to lengths of 3 cm or less), 30% of contractile events are conducted for 3–9 cm and the remaining events are conducted for greater distances (Dusdieker & Summers 1980). The following physiological basis for the similarity is suggested. After phase III activity of the MMC, there is a period of 15–30 min during which intestinal movement is essentially absent (phase I). This is sufficient time for sloughed enterocytes, mucus secretion, and fluid from the pancreato-biliary system and stomach to accumulate in the lumen. This material provokes enteric reflexes that cause local contractions and propagated contractions that run for short distances. The initiation of reflexes by food in the intestine is similar, thus the patterns of movement are similar. The stationary contractions occur irregularly and push the contents both orally and anally (Ehrlein et al. 1987). When the contracted area relaxes back to its original diameter, the contents return to the formerly contracted region. Thus there is a to and fro movement of the contents. Such mixing movements that segment the contents were originally described by Cannon, who recorded their occurrence in cats which had been fed a contrast medium and were observed by X-ray (Cannon 1902). Cannon’s description suggests a greater regularity of segmental contractions than was later described. The undoubted purpose of segmentation is to mix the chyme with the digestive juices and to expose it to the absorptive surface of the mucosa. Clusters of contractions that are effectively non-propulsive are also observed (Ehrlein et al. 1987). Although they tend to run anally for a few centimeters, when
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they stop the chyme generally runs back to where it was. Thus the clustered contractions contribute to mixing the contents. Peristalsis in vivo Migrating clusters of contraction or of slow waves with associated spike bursts occur both in the fed state and as part of phase II of the MMC (Dusdieker & Summers 1980, Ehrlein et al. 1987). These involve a similar phenomenon to phase III of the MMC, in that the migrating clusters consist of rapidly moving slow wave-associated contractions that form a slowly moving complex. In the dog these clusters of contractile events run for variable distance, from 3 to over 40 cm, before dying out. They are propulsive and resemble peristaltic events that have been described both in vivo and in vitro. A relaxation sometimes precedes these contraction clusters (Fig. 5.11). The migrating clusters move more quickly than phase III of the MMC. They have speeds of 7.5 cm/min in the jejunum and 4 cm/min in the ileum (Ehrlein et al. 1987) (slow waves move at about 200 cm/min in the dog small intestine). Similar slowly moving waves of circular muscle contraction were described by Cannon (1902, 1911, 1912) who observed them by X-ray in conscious cats. Cannon described these as peristaltic waves that proceeded in an oral to anal direction for distances of 4 or 5 cm and up to 7–8 cm. They traveled at
Fig. 5.11 Clustered (peristaltic), propagated contractions during phase II of the migrating
motor complex in intact dogs, recorded with closely spaced electrodes in the ileum (A) and jejunum (B). The cluster moves slowly along the intestine, but within the clusters individual contractions progress quickly (dashed lines). The individual contractions, which are generated by slow waves, are conducted for 5–7 cm in the ileum and 3–6 cm in the jejunum. A stationary cluster of contractions is underlined. Peristaltic complexes are sometimes preceded by relaxation (arrows). Similar peristaltic events are seen in the fed state. Reproduced from Ehrlein et al. (1987).
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1–2 cm/min. Incidentally, Cannon reported that movements of the intestine, which were probably contractile events evoked by slow waves (amplified by neural activity) traveled at 2–5 cm/s, i.e., at the same speed that slow waves travel. Slow wave speeds of 14–20 cm/s are recorded in the canine duodenum in vivo (Bass et al. 1961). It should be emphasized that the migrating peristaltic clusters are superimposed on the resting baseline pressure of 2–8 mmHg (Sherrington 1915, Abbott et al. 1943, Fink 1959). Thus, unlike peristaltic contractions that are studied in vitro, they are unlikely to be initiated simply by the intraluminal pressure. It is presumed that they are elicited by a mixture of the stimulation of IPANs by movements of the mucosa, by the intraluminal chemical environment and by the slight tension that occurs in the wall at the low intraluminal pressure that naturally occurs in the intestine (see Chapter 3). In order to study peristalsis in vivo many authors have provoked peristaltic events by distension, mechanical stimulation of the mucosa, or by chemical stimuli (Bayliss & Starling 1899, Bolzer 1949, Hukuhara et al. 1958, Hukuhara & Miyake 1959, Thomas & Baldwin 1971). Essentially the same phenomena to the spontaneous peristaltic events are seen, contractions that run from oral to anal for variable distances at speeds around 2 cm/min, that are sometimes preceded by detectable relaxation of the circular muscle. The peristaltic events that are deliberately provoked, like those that occur spontaneously, consist of clusters of rhythmic contractions, although the clusters are sometimes obscured because the individual contractions are fused (Fig. 3.1) or the length that is recorded from is too great to resolve the contractions. Nakayama (1962) used electrophysiological methods to record slow wave activity in anesthetized dogs during peristaltic reflexes that were provoked by a cotton wool bolus inserted into the lumen. He found that the slow waves that occurred during the peristaltic contraction cluster produced a greater number of action potentials, but that slow wave frequency of occurrence was not changed (Fig. 5.12).
Fig. 5.12 Electrical correlate of the ascending excitatory component of the peristaltic reflex of
the dog. The electrode recorded slow waves that had only a few superimposed action potentials, until a bolus was inserted immediately anal to the recording electrode (A). There was a prompt increase in the number of action potentials superimposed on each slow wave (this can be compared to the increased intensity of contractions during peristalsis in Fig. 5.11). At B, the bolus had been propelled 3 cm from the recording site and beyond this time the action potentials returned to the pattern seen before the bolus was introduced. Reproduced from Nakayama (1962).
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If the baseline pressure within the intestinal lumen is elevated above its normal resting level, to a pressure of 10–12 mmHg, clusters of contractions are still observed, despite the fact that the threshold for eliciting the peristaltic reflex is exceeded (Bogeski et al. 2003). Of course, clustering also occurs at normal pressure, but it might be that the intestine is close to the threshold for initiating peristalsis, and it meanders below and above the threshold. What is the mechanism that determines that peristaltic clustered contractions are limited to a brief period of 30–60 s at any one point in the intestine? This possibly occurs because the excitation is generated by the activity of groups of IPANs that are linked through excitatory synapses and that exhibit prolonged after-hyperpolarizing potentials (AHPs) after they fire action potentials (Chapter 2). In Chapter 3 it has been estimated that several thousand IPANs are activated when a peristaltic reflex is evoked. The activity in these neurons is augmented and co-ordinated by their reciprocal synapses. After IPANs have fired a small number of action potentials a compound late AHP develops that can persist for 2–30 s (Chapter 2 and Fig. 2.9). This AHP can block the invasion of action potentials into the cell body and their passage from the input processes to the output processes of IPANs (Chapter 2). It is suggested that before a peristaltic event the IPANs have an insufficient level of activity to provide the synaptic drive necessary to generate action potentials in the excitatory motor neurons. However, as IPANs are recruited during the quiescent period they act on each other through reciprocal synapses and there is eventually sufficient activity within the group of IPANs to activate the reflex. At this time the IPANs may fire at sufficient rates to generate AHPs that now restrict their output to interneurons and motor neurons and the peristaltic event subsides. At the same time, the propulsion of the contents may reduce the signals from distension that activate IPANs, contributing to the diminution of reflex activity. We have investigated the effect of blocking the IK channels whose opening is responsible for the late AHP. This dramatically changed the pattern of motility (Fig. 5.13). Whereas before the channels were blocked the intestine exhibited regular clustered peristaltic events, in the presence of the channel blocker there was an almost continuous series of contractions. These occupied only short distances (3–4 cm) and were not propulsive. This resembles a conversion of propulsive to mixing movements (Cannon 1902, Ehrlein et al. 1987) in the intestine. There is evidence that inhibitory motor neurons, which release NO as one of their principal transmitters (Chapter 4), may be continuously active, thus providing an inhibitory tone in the intestine. This could in part explain the lack of contractions elicited by slow waves between peristaltic events. The theory is supported by the large increase in contractile activity that occurs when NO synthesis is blocked (Bogeski et al. 2003; Ferens et al. 2005).
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Fig. 5.13 The effect of blocking the AHP of the intrinsic primary afferent neurons by the IK channel blocker TRAM34, given as an intravenous bolus, on the pattern of motility in the rat jejunum. Regular clusters of propulsive (peristaltic) contractions were converted to frequent non-propulsive (mixing) contractions. Furness and Ferens, unpublished.
Transit times, relation to contractile events Transit times through the human small intestine, which is 6–7 meters long, are 2–4 h (Read et al. 1986, Von Der Ohe & Camilleri 1992), which represents a speed of about 3–5 cm/min. In the normal rat, the transit time in the small intestine is about 100 min (Galligan & Burks 1982, Luck et al. 1993), corresponding to a speed of about 1 cm/min. This is slightly slower than the speeds of peristaltic complexes of 2–8 cm/min, but is considerably slower than individual propulsive waves that move at rates of around 1 cm/s (Table 5.2). Relationship between movements of the circular and longitudinal layers of the external muscle coat The major work of segmentation, of peristalsis, and of phase III of the MMC involves the circular muscle contracting to occlude the lumen. When the circular muscle contracts and shortens, it thickens. Were that thickening to be predominantly in the longitudinal direction of the intestine, the gut would lengthen and the occlusion of the lumen would be reduced. Thus it is not surprising that the longitudinal muscle contracts at the same time as the circular, as this restricts lengthening of the intestine and accentuates luminal occlusion. The simultaneous contraction of the two muscle layers was reported during propulsive contractions of the canine intestine in vivo (Bayliss & Starling 1899) (see Fig. 3.1) and has also been noted in the esophagus (Roman 1982, Mittal et al. 2005) and in other regions and species (Smith & Robertson 1998). Simultaneous electrical records from the two muscle layers also indicate that slow waves occur in synchrony (Suzuki et al. 1986). When
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a peristaltic reflex is induced by an imposed increase in intraluminal pressure, it is observed that the contraction of the longitudinal muscle begins slightly before that of the circular (Trendelenburg 1917). This has been called the preparatory contraction of the longitudinal muscle. In response to suggestions that there may be reciprocal movements of the longitudinal and circular layers (that is, the longitudinal muscle relaxes and the intestine lengthens when the circular muscle contracts), the relationship between their contractions has been re-examined recently, and the literature reviewed (Smith & Robertson 1998). These authors confirmed that the two layers contract together during propulsive reflexes of the intestine. Imaging methods to reveal Ca2+ transients in the muscle also show that the two layers are excited at the same time when motility reflexes are initiated (Stevens et al. 2000). There are many instances in which recordings from the intestine show that the longitudinal muscle layer elongates at the same time that the circular muscle contracts (Sarna 1993, Grider 2003). My interpretation of these observations is that when the contraction of the circular muscle is sufficiently strong it can overcome the longitudinal muscle contraction and force the longitudinal layer to lengthen. Nevertheless, the literature does indicate that force generated by longitudinal muscle restricts elongation when the circular muscle contracts. The peristaltic rush or giant migrating contraction Some authors have described very fast-moving rings of contraction in the small intestine (Meltzer & Auer 1907, Alvarez & Mahoney 1924, Sarna 1987). The rings are about 2–3 cm long and travel at about 1 cm/s. Once initiated at a point in the small intestine, they generally continue undiminished the full length of the organ. In fact, they can traverse the entire small intestine and continue into the large intestine. They occur infrequently and unpredictably in fasted dogs (Sarna 1987). These events are more frequently seen as a reaction of the intestine to toxic conditions, in which the intestine is working to expel the toxin. Peristaltic rushes can be induced by pathogens such as V. cholerae, nematode infestation or E. coli toxin (Mathias et al. 1982, Cowles & Sarna 1990, Lind et al. 1991). The reaction of the intestine to bacterial pathogens is accompanied by copious watery fluid secretion. Irritant substances, such as castor oil (ricinoleic acid) also cause peristaltic rushes (Mathias et al. 1978). It is feasible that purgatives such as castor oil exert their main effect by eliciting these expulsive reflexes. If the intestine is transacted and re-anastomosed, the peristaltic rush ends at the anastomosis (Otterson & Sarna 1994). This observation, and the speed of propagation of the peristaltic rush, suggests that it is mediated by a motor program of the enteric nervous system.
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Emesis and retropulsion in the small intestine Vomiting can be initiated from or through the central nervous system by a variety of circumstances, including unpleasant odor, stress, vestibular stimulation in motion sickness, and somatic pain. It can also be initiated by toxic compounds in the gut, such as hypertonic NaCl, CuSO4, products of putrefaction, and plant alkaloids. Nausea and vomiting are elicited through pathways that converge on the area postrema (Hatcher 1924, Borison & Wang 1953). The area postrema has been lesioned in human patients with intractable vomiting, which stopped the vomiting episodes and raised the threshold dose of apomorphine that triggered vomiting (Lindstrom & Brizzee 1962). When nausea is initiated from the gastrointestinal tract, the principal afferent pathway is the vagus nerve (Wang & Borison 1951, Andrews et al. 1990). Vomiting involves co-ordinated activities of somatic muscles: the glottis is closed and thoracic, diaphragmatic, and abdominal muscles contract to produce a high intra-abdominal pressure which forces the gastric contents towards and into the mouth. At the same time, or soon after, strong contractions occur in the small intestine and are conducted towards the stomach, carrying with them the contents of the proximal ileum, jejunum and duodenum (Alvarez 1925, 1940, Ehrlein 1981). The events cause large increases in intraluminal pressure and travel rapidly, at 2–6 cm/s, in an oral direction (Stewart et al. 1977, Ehrlein 1981, Lang et al. 1986, 1999). The retropulsive contractions are blocked by atropine, indicating that they are caused by the activation of enteric cholinergic motor neurons (Stewart et al. 1977, Lang et al. 1986). Sub-diaphragmatic vagotomy prevented the intestinal retropulsion in response to a centrally acting emetic, apomorphine, but did not prevent the retching or vomiting associated with contractions of the body wall. Thus the vagus is the efferent nerve through which the reverse peristalsis of vomiting is triggered. Whether a patterned output from vagal efferent pathways coordinates the retropulsion, or whether the vagus sets off a motor program in the enteric nervous system is unknown. Motility of the colon The essential role of the enteric nervous system in the colon is illustrated by the unfortunate conditions of Hirschsprung’s and Chagas’ diseases. In the former the enteric ganglia are congenitally absent and in the latter a trypanosome infection destroys the ganglia. In both cases the muscle can contract in response to electrical stimulation or muscarinic agonists, but co-ordinated propulsion is lost and the colon is incapable of normal function (Truelove 1966). The colon fills almost continually from the small intestine, it removes water and electrolyte from its contents as it converts the liquid material arriving from
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the ileum into solid or semi-solid feces and it empties the contents of the left colon and rectum intermittently. Its movements reflect these functions. The muscle of the colon, like that of the small intestine, exhibits slow wave activity, and, like the small intestine, the slow waves cause contractions when they are of sufficient amplitude (Christensen et al. 1969). Mixing movements in the cecum, right colon and approximately the first half of the transverse colon expose the contents to the absorptive epithelium. These consist of local contractions (often called haustral contractions) and contractions that push the contents, often in a retrograde direction towards the ileo-cecal valve, as well as in an anal direction for short distances (Cannon 1902, Hertz & Newton 1913, Ritchie 1971, Christensen 1989). This mixing retains fecal material in the right colon for periods of up to 48 h (Christensen 1994). In addition to the mixing movements, very occasional movements that propel part of the contents in an anal direction occur. These have been called colonic peristalsis (Mann & Hardcastle 1970) or colonic mass movements (Christensen 1989). These occur about 10 times per day, and are associated with a desire to defecate (Hasler 2003). The mass movements are probably the consequence of peristaltic reflexes that are readily evoked by distension or mucosal irritation in the colon (Bayliss & Starling 1900b, Raiford & Mulinos 1934, Hukuhara & Miyake 1959, Frigo & Lecchini 1970). The mass movements are reduced in amplitude by hexamethonium and atropine, which indicates their dependence on neural activity (Tomaru et al. 1993). The human colon, unlike the small intestine, does not have an MMC (Hasler 2003). The defecation reflex is triggered by neurons whose axons reach the colon through the pelvic nerves, which are believed to initiate activity in the enteric nervous system of the colon and rectum, including causing a relaxation of the internal anal sphincter that empties the distal part of the colon (Truelove 1966). When propulsive movements are recorded from the colon in vivo or in vitro, the propelled bolus is preceded by a relaxation (Bayliss & Starling 1900b, Frigo & Lecchini 1970) that is much more prominent than it is in the small intestine, which suggests that the descending inhibitory component of the peristaltic reflex (Chapter 3) is more important in the colon than it is in the small intestine. The importance of the enteric inhibitory innervation of the colon and rectum is also suggested by the obstructive contraction of the distal part of the colon that occurs in Hirschsprung’s disease, in which the contractions of more proximal regions cannot push the contents past the distal, constricted, region from which enteric innervation is lost. The recto-anal reflex A useful test of the integrity of the rectal and anal innervation is to distend the sigmoid colon or to irritate its mucosa. The response is a relaxation of
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the internal anal sphincter, which occurs only if the enteric reflex circuitry is present (Christensen 1994; and see section below on anal sphincter). Neural control of the esophagus In most species, the external muscle of the esophagus is composed of striated muscle fibers over almost all its length. In the human, approximately the first third is striated, the middle third is mixed and the last part has a smooth muscle wall. The controlling motor innervation of the striated muscle comes directly from vagal fibers that do not synapse in enteric ganglia. Consistent with this, section of the vagal fibers causes paralysis of the striated muscle esophagus, which does not recover its function (Ingelfinger 1958). Moreover, the influence of distension caused by the food that is swallowed on the regulation of peristalsis in the striated muscle part is through vago-vagal pathways (Roman 1982, Lu & Bieger 1998). Motor program that control the speed and strength of contraction of the striated muscle are generated by circuits in the brainstem (Bieger 1993, Jean 2001). Thus, although the myenteric plexus is prominent in the striated muscle esophagus, it does not have a key role in generation or control of peristalsis. Nevertheless, myenteric neurons do supply an innervation to about a third of the endplates and thus, unlike motor endplates elsewhere, individual endplates in the esophagus receive dual innervation, one axon being from a vagal motor neuron and the other originating from a cell body in the myenteric plexus (Neuhuber et al. 1994, Kuramoto et al. 1996, Wörl et al. 1997, Wu et al. 2003a). In the cat, monkey, and rat the vagal endings are immunoreactive for CGRP, and the endings of myenteric origin have NOS immunoreactivity, implying that transmission from these neurons is nitrergic. In the pig, the vagal endings have PACAP immunoreactivity and the intrinsic endings are NOS immunoreactive as in other species (Wu et al. 2003a). Double staining using antibodies to CGRP and NOS indicates that both vagal and enteric nerve endings are closely apposed at the endplates, such that they may interact presynaptically (Wörl et al. 1997). Presynaptic inhibition of vagal excitatory transmission has been shown to occur by experiments in which enteric NOS neurons were stimulated indirectly (Izumi et al. 2003). The stimulation reduced vagally induced muscle contraction, and inhibition of vagal transmission to the esophageal muscle was abolished by blocking NOS activity with l-NAME. Thus the enteric nervous system seems to have a role in modulating peristalsis in the upper esophagus. The enteric innervation may have a greater role in young animals, because all motor endplates receive an enteric innervation at days 4–10 postnatal, after which there is partial withdrawal of innervation, and in adult about 35% of the endplates receive an enteric innervation in addition to the vagal innervation that all striated muscle cells receive (Breuer et al. 2004).
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Other functions of esophageal enteric neurons in the striated muscle part of the esophagus still require investigation, but could include control of the muscularis mucosae and of mucus secretion. To different extents, the external coat of the distal part of the esophagus in some species (e.g. human, possum, cat, but not dog) is composed of smooth muscle. The lower esophageal sphincter in all species is composed of smooth muscle. The smooth muscle is innervated by motor neurons with cell bodies in the myenteric ganglia. In contrast to the striated muscle part of the esophagus, the smooth muscle esophagus is not paralyzed by vagotomy, and in fact continues to be able to exhibit propulsive peristaltic contractions in response to a bolus of food introduced into its lumen (Cannon 1907, Tieffenbach & Roman 1972). Moreover, peristaltic reflexes can be elicited by distension of a segment of the smooth muscle esophagus that is removed from an animal, such as the possum (Christensen & Lund 1969). Nevertheless, the central nervous system, through the vagus, does appear to exert command over the distal esophagus. Under normal circumstances, the smooth muscle part of the esophagus receives swallowed food from the more proximal esophagus and there is a well-co-ordinated transition of propulsion between the two parts. This co-ordination depends in part on central command centers directing the peristalsis in the distal (smooth muscle) esophagus. In one series of experiments it was shown that deviation of the swallowed food bolus at the cervical level did not prevent progression of the peristaltic wave into the distal esophagus (Janssens et al. 1976), implying that once the motor program was set up it continued without the need for distension of the smooth muscle part of the esophagus. This concept is supported by the observation that peristaltic activity of the distal esophagus can be elicited by distension of the pharynx or upper esophagus, or even by electrical stimulation of the vagal afferent nerves supplying this region, and occurs when the proximal esophagus is prevented from contraction by blocking transmission from its motor innervation with curare (Roman 1982). Gall bladder motility The contraction of the gall bladder in response to a meal is dependent on release of the duodenal hormone cholecystokinin (CCK). After either a meal or CCK infusion, the gall bladder slowly contracts and empties bile into the duodenum (Shaffer et al. 1980, Lawson et al. 1983). The degree of contraction is closely correlated with CCK concentration in the plasma (Wiener et al. 1981) and is blocked by CCK receptor antagonists (Hildebrand et al. 1990). There is good evidence that the responses have a substantial indirect component, through the actions of CCK on neurons in the gall bladder, and may have a lesser direct component through CCK action on muscle of the gall
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bladder. The contraction of the gall bladder in response to intravenous CCK or to intraduodenal infusion of nutrient is blocked or substantially reduced by atropine or hexamethonium, suggesting that the physiological effect of CCK is dependent on nicotinic excitatory transmission to the cholinergic motor neurons of the gall bladder (Fisher et al. 1985, Hanyu et al. 1990, Nelson et al. 1996). Intracellular microelectrode recordings from enteric neurons in the gall bladder show that CCK does not have a post-synaptic action on the gall bladder enteric neurons, but potently increases the amplitudes of fast EPSPs (Mawe 1990, Bauer et al. 1991). The threshold for facilitation of the amplitude of the fast EPSP was 10 pM in guinea-pig, and a maximum effect occurred at 1 nM (Mawe 1990). In humans, plasma levels of CCK rise to 5–10 pM after a meal (Masclee et al. 1990). A major component of the fast excitatory input to gall bladder neurons comes from the vagus, and transmission at the vagal endings is facilitated by CCK (Mawe et al. 1994). However, in humans, responses of the gall bladder to intraduodenal fat are not reduced by vagotomy (Masclee et al. 1990), although there is about 20% reduction in the response to a meal (Fisher et al. 1985). Moreover, in transplant recipients of liver plus gall bladder, in which all extrinsic innervation of the gall bladder is severed, the contraction of the gall bladder following a fatty meal is normal, as is the release of CCK (Vezina et al. 1994). It is possible that adaptation of enteric circuits occurs after the transplant; nevertheless, the data suggest that intrinsic excitatory connections between enteric neurons within the human gall bladder (where CCK has its effect) are active after extrinsic denervation, and that there are possibly intrinsic gall bladder reflex pathways. The existence of neuronal activity derived from intrinsic circuits is supported by the effects of CCK on the isolated guinea-pig gall bladder, where CCK-induced contractions are blocked by tetrodotoxin or atropine (Brotschi et al. 1990), and CCK’s actions are to facilitate transmission by a presynaptic action (Mawe 1990). There is also evidence for intrinsic secretomotor reflexes in the gall bladder (Chapter 6). Sphincters The purposes of the sphincters are to restrict and regulate the movement of the contents of the gastrointestinal tract between adjacent regions. The lower esophageal sphincter is normally tonically contracted, which restricts reflux of gastric contents into the esophagus. It relaxes during swallowing, prior to belching, and during vomiting. The pyloric sphincter (pyloric canal) is almost closed during digestion. It maintains a narrow lumen through which small aspirates of gastric fluid are pushed during digestion (Fig. 5.7). It largely prevents bile being refluxed into the stomach. The ileo-cecal sphincter (and ileo-cecal valve) restricts the bacteria-laden contents of the cecum gaining
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access to the terminal ileum, and regulates ileal emptying. The internal anal sphincter, which is part of the digestive tract, and the external anal, striated muscle, sphincters maintain anal continence. Finally, the sphincter of Oddi regulates passage of contents between the pancreato-biliary system and the duodenum. Each of the sphincters of the digestive tract consists of smooth muscle which is in continuity with the circular muscle coat of the immediately proximal region. The myenteric plexus is continuous across the sphincters, connecting the organ that lies proximal to that which is distal to the sphincter, except, of course, for the internal anal sphincter. The enteric motor neuron supply to sphincter muscle matches that of the non-sphincter muscle; the sphincters are innervated by excitatory motor neurons whose principal transmitter is acetylcholine, but which also contain tachykinins, and by inhibitory motor neurons whose principal transmitter is nitric oxide, but which also have PACAP, VIP, and ATP as co-transmitters (see Chapter 4). The lower esophageal sphincter This sphincter is usually held closed by contraction of the sphincter muscle, and is relaxed when food enters the stomach from the esophagus or when there is release of gas from the stomach into the esophagus, or during vomiting. The resting tension (tone) of the lower esophageal sphincter muscle is primarily caused by maintained activity of enteric cholinergic motor neurons that supply the muscle. Their cell bodies are in the myenteric plexus close to the sphincter and in the distal esophagus (Brookes et al. 1996). The tone that they generate is halved by vagal cooling, and the remaining tone is reduced to a low level, about 15% of the original tone, by blocking transmission to the neurons with hexamethonium (Reynolds et al. 1984). Atropine reduced the tone to the same level that was observed after hexamethonium. Thus the activity in the final motor neurons is generated by excitatory input from the vagus, augmented by local excitation through the enteric nervous system. Consistent with this study in the cat, human lower esophageal sphincter pressure is reduced to about 30% by an effective dose of atropine (Dodds et al. 1981). Nevertheless, there does seem to be a contribution to tension in the sphincter which is intrinsic to the muscle (Roman 1982). Lower esophageal sphincter relaxation is caused by swallowing and by food in the esophagus. Swallowing-induced relaxation is prevented by vagal cooling which, along with other evidence, indicates that it is a vago-vagal reflex (Reynolds et al. 1984). With the vagus nerve blocked by cooling, or when the lower esophagus is extrinsically denervated, esophageal distension still causes relaxation of the sphincter, indicating that there is a local (enteric) descending inhibitory reflex (Mann et al. 1968, Reynolds et al. 1984). The final motor neurons
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are enteric inhibitory neurons, and the primary transmitter that causes muscle relaxation is NO (Tottrup et al. 1992, Boeckxstaens 2005). Distension of the stomach causes the lower esophageal sphincter to increase its contractile force, presumably as a protection against reflux of gastric contents into the distal esophagus (Franzi et al. 1990). When the distension is made slowly, transient relaxations of the sphincter are superimposed on the increased sphincter tone (Franzi et al. 1990). The transient relaxations are initiated from the cardiac part of the stomach, close to the sphincter. The physiological function of these transient relaxations is probably in belching, to allow gas to be released from the stomach. Both the reflex increase in sphincter pressure when intragastric pressure is increased (Diamant & Akin 1972) and the transient relaxations (Martin et al. 1986) are mediated through vago-vagal reflex pathways. Pyloric sphincter The pyloric sphincter remains almost closed most of the time. Even strong contractions of the antrum, which lead to high pressures oral to the sphincter, do not cause it to relax (Fig. 5.7). So, although the sphincter has an inhibitory innervation from enteric neurons (Vogalis & Sanders 1990), increased pressure oral to the sphincter does not elicit a descending inhibitory reflex. In this respect it is unlike the lower esophageal sphincter and the small intestine distal to the sphincter. Thus the powerful contractions that occur in the antrum push the gastric contents against the sphincter and only small aspirates get past with each contraction (Cannon 1911). Because the sphincter is virtually closed to the passage of solids, but the antral peristaltic waves do not occlude the lumen (Fig. 5.7), the solids are pushed backwards towards the gastric body, almost bouncing off the sphincter with each gastric contractile wave (Kelly 1980). Removal or lesion of the sphincter destroys this function, and the stomach empties at an unacceptably fast rate, commonly resulting in the dumping syndrome (Humphrey et al. 1972). Acidification of the proximal duodenum increases the baseline pressure and contractile activity of the pyloric sphincter (Allescher et al. 1989). The reflex is abolished by hexamethonium and is substantially reduced by atropine, suggesting that it is an intramural ascending excitatory reflex. Ileo-cecal junction This region has different anatomy in different species, although in all it acts to restrict the reflux of ceco-colic contents and the bacterial load they bear into the terminal ileum (Bogers & Van Marck 1993). In humans, the end of the ileum protrudes slightly into the cecum where it forms a papilla. This results
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in the circular muscle of the cecal wall forming a cuff around the circular muscle of the last part of the ileum, which has a sphincter-like appearance. The mucosal folds of the papilla have the appearance of a valve. So reflux may be limited both by a smooth muscle sphincter and the mucosal folds. The idea of a sphincter function is supported by studies in human subjects, in which a zone of elevated intraluminal pressure was located at the ileo-cecal junction and distension of the cecum increased pressure at the junction (Dinning et al. 1999). A high pressure zone is also present in the distal ileum and ileo-cecal junction in other mammals (Quigley et al. 1985, Malbert 2005). An inhibitory innervation, in which transmission is through NO, has been demonstrated for the sphincter in dog (Boeckxstaens et al. 1991). These neurons may be involved in reflex relaxation of the sphincter when a pressure wave in the ileum arrives (Kelley & De Weese 1969, Dinning et al. 1999). Increasing the colonic intraluminal pressure causes the sphincter zone to contract, which appears to be similar to ascending excitatory reflexes observed in the lower esophageal sphincter, in this case presumably restricting reflux of cecal or colonic contents into the distal ileum (Quigley et al. 1985). Internal anal sphincter The internal anal sphincter is a thickening of the circular muscle of the rectum; there is no further smooth muscle beyond this point, and the myenteric plexus finishes at the sphincter. The sphincter muscle is usually contracted, which provides closure of the last part of the anal canal (Rattan 2005). A reflex relaxation of the sphincter, the recto-anal reflex (Fig. 5.14), is evoked when feces enters the normally empty rectum during defecation, or when the rectum is artificially distended or its mucosa is irritated (Gowers 1877, Schuster et al. 1965). This reflex is dependent on the integrity of the enteric plexus; when the neurons are missing in congenital aganglionosis (Hirschsprung’s disease) there is no relaxation in response to rectal distension, but the muscle is still capable of contracting, for example when exposed to agonist drugs. As is the case for the other sphincters, the relaxation is mediated through the enteric inhibitory motor neurons and VIP and NO contribute to the transmission (Biancani et al. 1985, Rattan et al. 1992). Sphincter of Oddi (choledeco-duodenal sphincter) The sphincter of Oddi controls the movement of bile and pancreatic secretion into the duodenum (Woods et al. 2005). It is formed by the circular muscle layer of the wall of the common bile duct as it enters the duodenum, and part of it is within the duodenal wall. Its structural organization differs considerably between species. There are also differences in its physiology: in some species
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Fig. 5.14 The recto-anal reflex. This is a reflex that is mediated through the enteric nervous system, and can be initiated by mucosal irritation or distension of the rectum. In this experiment, in a human volunteer, the rectum was distended with different volumes of air, at the dots. This resulted in graded relaxation of the internal anal sphincter (lower trace). Reproduced from Schuster et al. (1965).
it seems simply to contract and relax to allow the passage of the bile, whereas in others there is evidence of peristaltic movements that push the bile into the duodenum (Mawe et al. 2003). Like the other sphincters of the gastrointestinal tract, the sphincter of Oddi is innervated by excitatory and inhibitory enteric motor neurons. A meal or infusion of CCK relaxes the sphincter in cat and human (Toouli et al. 1982, Behar & Biancani 1987, Mawe et al. 2003). The relaxation appears to be caused indirectly, by activation of the inhibitory motor neurons. In the anesthetized cat, relaxation of the sphincter in response to CCK was blocked when tetrodotoxin was infused into its arterial supply (Behar & Biancani 1987). In some other species (including dog, rabbit, and opossum), CCK promotes peristalsis-like propulsion in the extraduodenal part of the sphincter, and this can be inhibited by atropine or tetrodotoxin, suggesting that activation of enteric neurons contributes to the enhancement of propulsive contractions (Cox et al. 1990). Muscle of the mucosa The mucosa has two types of muscle, a sheet-like muscle at its interface with the submucosa, the muscularis mucosae, and strands of muscle which extend between the glands in the stomach and colon, and which run within the cores of the villi in the small intestine. Each of these is composed of smooth muscle. Although there has not been extensive investigation of these muscles, they do appear to receive both excitatory and inhibitory innervation from enteric neurons. The muscularis mucosae exhibits spontaneous contractile activity, which in one study has been shown to be due to spontaneous action potential generation in the muscle (Morgan et al. 1985).
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The villous smooth muscle, by its rhythmic contractions, is supposed to agitate the luminal contents to disrupt the unstirred water layer that might restrict diffusion of nutrients to the intestinal epithelium (King & Robinson 1945). Transmural stimulation, to activate nerve fibers in the muscularis mucosae, cause both contractions and relaxations. The contractions are inhibited by atropine, and are therefore likely to result from stimulation of enteric cholinergic motor neurons (Gallacher et al. 1973). Relaxations in response to nerve stimulation are blocked by apamin (Angel et al. 1983, 1984), suggesting that they are mediated through enteric inhibitory neurons (see Chapter 4). Hambleton (1914) reported that exposure of the intestinal lumen of the dog to nutrients and various chemical solutions caused rhythmic shortening and elongation of the villi which were blocked by nicotine or atropine. He concluded that they were due to a local neural reflex. Stimulation of villous motility by vagal stimulation, blocked by atropine, has also been reported (Womack et al. 1988). The muscularis mucosae in the stomach may also have a mixing role. In elegant experiments, the pressures in the lumens of gastric glands have been measured and found to oscillate with a rhythm of 4–5 oscillations per min (Synnerstad et al. 1998). Addition of VIP reduced the basal pressure and significantly reduced the amplitudes of oscillations. The muscle strands in the mucosa were closely apposed by VIP-immunoreactive nerve fibers. VIP is one of the constituents of inhibitory neurons supplying gut muscle, the other prominent constituents being NO and ATP (Chapters 2, 4). Thus the muscularis between the glands may receive an enteric inhibitory innervation, as shown for the bulk of the muscularis mucosae. The authors cautiously suggest that the contractions of the muscle and the oscillating pressure it causes may assist in emptying the contents of the gland into the gastric lumen. Mechanism of sympathetic inhibition of motility in nonsphincter regions The histochemical observations that noradrenergic fibers ramify extensively in myenteric ganglia but are rare in the musculature of non-sphincter parts of the intestine suggest that the sympathetic inhibition of gastrointestinal movement is caused by the noradrenergic axons interacting with enteric neurons. It is thus interesting that Lister (1858) had concluded that the sympathetic mesenteric nerves caused inhibition by acting on the myenteric ganglia. He wrote: “the inhibitory influence does not operate directly on the muscular tissue, but on the nervous apparatus by which its contractions are, under ordinary circumstances, elicited.” Lister had found that the propagation of contraction away from a point of local irritation of the intestine which, he had concluded from other experiments, depended on the intramural ganglionated
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plexuses, was prevented by stimulation of the thoracic source of the splanchnic nerves. However, he found that he could still provoke a contraction of the inhibited intestine by local stimulation. A neural site of action is implied by the observations that exposure of the intestine to norepinephrine and sympathetic nerve stimulation both reduce the amount of acetylcholine released from intrinsic cholinergic neurons (Paton & Vizi 1969, Vizi & Knoll 1971). Stimulation of mesenteric nerves at 10 Hz reduced the resting output of acetylcholine from rabbit jejunum by 45% (Vizi & Knoll 1971). Elegant evidence that noradrenergic neurons inhibit movement through an action on enteric cholinergic neurons comes from experiments in vivo (Kewenter 1965, Jansson & Martinson 1966, Hulten & Jodal 1969, Jansson 1969b, Jansson & Lisander 1969, Jansson et al. 1969). Kewenter (1965) showed that nerve-mediated excitation of the intestine, but not direct excitation by acetylcholine, was strongly inhibited by sympathetic nerve activity. If noradrenergic nerves supplying the ileum were tonically active, only weak excitation occurred when the vagus nerves were stimulated. If noradrenergic transmission was blocked by ergotamine or guanethidine, the effect of vagal stimulation was markedly enhanced. In contrast, contractions in response to acetylcholine were not influenced by the level of activity of noradrenergic nerves. Jansson and his colleagues demonstrated a similar result for the stomach. The activation of noradrenergic nerves during an intestino-gastric inhibitory reflex did not affect the basic myogenic tone of the stomach, but it inhibited contractions in response to stimulation of vagal cholinergic nerves (Jansson & Martinson 1966). In contrast, the gastric contraction caused by exogenous acetylcholine was unaffected by the reflex firing of noradrenergic nerves to the stomach. Direct stimulation of the noradrenergic nerves at up to 10 Hz produced prompt inhibition of movement when vagal fibers were active but gave only weak inhibition after the animals were treated with atropine. It should be pointed out that the stomach of the cat is still capable of substantial relaxation to appropriate stimuli after the administration of atropine (Jansson 1969a). In addition, stimulation of noradrenergic nerves supplying the stomach, in which there was an elevated activity of enteric cholinergic neurons, markedly inhibited the cholinergic contractions even at low frequencies of stimulation (Jansson & Lisander 1969). After the cats had been given atropine, similar stimulation of noradrenergic nerves failed to relax the gastric muscle. Moreover, the activation of noradrenergic nerves by stimulation of the hypothalamus or of somatic pressor afferents inhibits vagal excitation of the cat stomach, but does not affect contractions caused by exogenous acetylcholine (Jansson 1969b, Jansson et al. 1969). A ganglionic site of action of inhibitory noradrenergic fibers also applies to the colon (Hulten & Jodal 1969). Low frequencies of stimulation of noradrenergic neurons suppressed contractions
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caused by pelvic or vagus nerve stimulation or by spontaneous activity of enteric excitatory neurons, but neither the myogenic contractions of the colon nor the contractions caused by exogenous acetylcholine were antagonized. In contrast to the results of Jansson and his colleagues, Reed and Sanders (1971) found that stimulation of splanchnic nerves equally reduced the excitatory effects of vagal stimulation and close intra-arterial injection of acetylcholine in the cat stomach, suggesting an action at the muscle in their experiments. Reed and Sanders stimulated the distal ends of the severed splanchnic nerves in cats under ether anesthesia at a frequency of 10 Hz, although the experiments of Jansson and Lisander (1969) suggest that stimuli at frequencies of 2 or 4 Hz act through the ganglia, but those at 8 or 16 Hz also act directly on the muscle. The effects on motility of activation of noradrenergic neurons through central reflex pathways can be mimicked by direct stimulation at 2–8 Hz (Jansson & Martinson 1966). The direct action on the muscle could be a result of release from the few noradrenergic fibers which run in the musculature, or overflow of norepinephrine from fibers supplying the myenteric plexus, or a combination of these effects. When cholinergic actions are abolished by muscarinic blocking agents (hyoscine or atropine), stimulation of paravascular nerves with frequencies as low as 1–2 Hz still inhibits intestinal movements in vitro (Gillespie 1960, Burnstock et al. 1966, Campbell 1966) and Gershon (1967) found that a norepinephrine inhibition of the guinea-pig stomach occurred with stimulus frequencies of 5 Hz even though excitation of intramural ganglia was blocked by hexamethonium and the action of acetylcholine on the muscle was blocked by hyoscine. This implies that norepinephrine released from nerves within the musculature is more effective in vitro, probably because it diffuses from the terminals in the myenteric plexus to the muscle, whereas in vivo it would be swept up into the circulation. This inhibition is probably due to norepinephrine released both from the myenteric plexus and from the nerves within the muscular layers of the guinea-pig stomach. Investigations utilizing intracellular microelectrodes indicate that noradrenergic nerve fibers supplying the myenteric ganglia cause a presynaptic inhibition of cholinergic transmission to S neurons (Hirst & McKirdy 1974b). Exogenous norepinephrine caused a similar depression of transmission (Nishi & North 1973b). Stimulation of noradrenergic nerves at frequencies up to 50 Hz caused no changes in membrane properties of myenteric neurons, even though some myenteric neurons can be hyperpolarized by agonists acting at α-receptors (Morita & North 1981a). In contrast, noradrenergic neurons supplying submucous ganglia have direct inhibitory effects on neuronal excitability by hyperpolarizing the neurons (Chapter 6). Inhibition of acetylcholine release from enteric cholinergic neurons is mediated mainly through α-receptors of the α2 subtype (Drew 1978). Direct
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actions on the muscle can be through either α- or β-receptors. Whether one or both of these receptor types is present on the muscle differs between regions of the gastrointestinal tract and between species. Sympathetic innervation of the sphincters Histochemical studies reveal a noradrenergic innervation of gastrointestinal sphincters that is often markedly more dense than in adjacent non-sphincter regions (Costa & Gabella 1971, Gillespie & Maxwell 1971, Mori et al. 1971, Furness & Costa 1973b, Howard & Garrett 1973). With few exceptions, those investigators who were able to record mechanical responses from sphincter muscle following stimulation of the sympathetic supply reported a constrictor effect which was mimicked by epinephrine and norepinephrine. The lower esophageal sphincter is contracted by epinephrine or norepinephrine and by sympathetic nerve stimulation in cat, rabbit, dog, monkey and human (Brucke & Stern 1938, Botha 1962). The sphincter of Oddi is contracted by stimulation of the splanchnic nerves (Doyon 1894, Westphal 1923), but a reflex relaxation to distension of the gall bladder mediated by a pathway through the celiac ganglion has been reported in the cat (Wyatt 1967). There are both α-excitatory and β-inhibitory receptors for catecholamines in the muscle and exogenous norepinephrine can cause relaxation or contraction, depending on the conditions (Persson 1971). There is general agreement that in a wide variety of mammals the ileo-colic sphincter contracts to sympathetic nerve stimulation and to epinephrine and norepinephrine (Malbert 2005). In dog and guinea-pig, stimulation of sympathetic nerves causes contraction of the distal ileum (Smets 1936, Munro 1953), so that this final part of the small intestine may function with the ileo-colic sphincter to inhibit transport between the ileum and the colon. The internal anal sphincter is contracted by stimulation of the sympathetic nerves in all mammals with the probable exception of the rabbit, in which it relaxes in response to sympathetic nerve stimulation and to epinephrine (Langley & Anderson 1895, Langley 1901). The human internal anal sphincter contracts when exposed to epinephrine or norepinephrine (Friedman 1968). In all the sphincters, contractions elicited by epinephrine and norepinephrine are blocked by α-adrenoceptor-blocking drugs and are sometimes reversed to relaxations blocked by β-adrenoceptor-blocking drugs. The βadrenoceptor agonist isoprenaline usually relaxes the sphincters. The sphincter muscle therefore has α-excitatory and β-inhibitory receptors, but it seems that it is the α-receptors which are activated physiologically by norepinephrine released by the sympathetic neurons. The excitatory effect of noradrenergic nerves at the sphincters is a direct effect of norepinephrine on receptors in the muscle.
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Physiological effects of noradrenergic neurons on motility in undisturbed animals There is little or no tonic activity of pathways from the central nervous system to activate the noradrenergic nerves that cause inhibition of non-sphincter regions of the gastrointestinal tract, but there is some evidence which suggests that noradrenergic nerves normally exert a tonic constrictor action on some sphincter muscle. The lack of tonic, centrally directed, activity of the motility inhibiting sympathetic neurons contrasts to the readily observable tonic activity of intestinal secretomotor inhibitory pathways (Chapter 6) and of sympathetic vasoconstrictor neurons to gastrointestinal blood vessels (e.g. Kremer & Wright 1932). In most cases where a tonic central drive to noradrenergic nerves that affect motility has been proposed, the experiments have involved exposure of the viscera or surgical interference at the time of observation. Laparotomy, exposure, or handling the intestine causes a profound and long-lasting inhibition of gastrointestinal tone and even cutting the surface skin of an anesthetized animal causes a significant suppression of motility (Cannon & Murphy 1906, Olivecrona 1927, Garry 1934, Jansson & Lisander 1969). Cannon (1906) avoided this difficulty by using an X-ray method to examine movements of the stomach and intestines of cats before and after sympathectomy. His experiments show that there is no tonic sympathetic inhibition of gastric movement or tonic closure of the pyloric sphincter in rested cats. Once the effects of surgical interference has subsided, there is no evidence of continued noradrenergic inhibition of gastric motility (Jansson & Lisander 1969), although McCrea (1926) had described experiments in which X-ray examination before and after section of sympathetic nerves did indicate some tonic activity of these fibers in the dog. Cannon’s (1906) experiments indicate that there may be tonic inhibition of small intestine motility induced by certain substances when present in the contents, because the passage of protein (lean beef) was accelerated by severing the splanchnic nerves, although the passage of carbohydrate (mashed potato) was almost unaffected. Removal of the inferior mesenteric ganglia of the cat causes little appreciable change in the activity of the colon and rectum, which were examined by X-ray (M’Fadden et al. 1935). In contrast, removal of the celiac ganglia can have profound effects, suggesting that peripheral entero-gastric reflexes can be active (see below). In fasted dogs, all phases of the intestinal migrating motor complex (Chapter 5) are present after removal of the celiaco-mesenteric plexus (Marlett & Code 1979). There was increased myoelectric activity during phase II, and the duration of this phase increased. The speed of propagation of the contractile complex, phase III, along the intestine was increased. In guinea-pigs, destruction of noradrenergic neurons with 6-hydroxydopamine also increased the
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duration of phase II, but had little effect on other features of the periodic motor activity of the small intestine (Galligan et al. 1986). Even with X-ray techniques, it is difficult to determine whether the sympathetic nervous system contributes to the maintenance of sphincter tone. However, there is no evidence that tonic activity of noradrenergic nerves is a normal requirement of sphincter function. For example, in laboratory animals with their entire sympathetic chains excised, there are no apparent changes in the digestive process and no long-lasting changes in sphincter activity (Cannon et al. 1929, Moore 1930). Division of sympathetic nerves does not usually cause any significant change in the tone of the lower esophageal sphincter (Ochsner & Debakey 1940, Ingelfinger 1958). In the case of the ileo-cecal sphincter, activity in sympathetic neurons appears to contribute to tone in dog and rat, but not in cat (Elliott 1904, Hinrichsen & Ivy 1931). Langley and Anderson (1895) found that section of the lumbar splanchnic nerves reduced the tone of the internal anal sphincter in the cat and that there was an even greater reduction if the inferior mesenteric ganglia were removed. Reflex activities of sympathetic neurons that affect motility Sympathetic pathways to the non-sphincter parts of the gastrointestinal tract are involved in two types of reflex affecting motility. The first are reflexes between gut regions in which a distal region regulates activity in a more proximal region in order to adjust the rate of transit and composition of the material that the proximal region passes on to the distal region. The second type is generalized protective reflexes in response to adverse conditions in the abdominal cavity or even beyond the abdomen. The first type of reflex is an entero-enteric regulatory reflex, the second is a generalized sympathetic inhibitory reflex. Sympathetic entero-enteric regulatory reflexes There are many different entero-enteric regulatory reflexes through which one part of the digestive tract influences another. Some are mediated through hormones, some through both hormones and neurons and some through neurons alone (or at least predominately). In this section, only entero-enteric reflexes that involve activation of sympathetic neurons are discussed. These reflexes are mediated through prevertebral ganglia and do not require connection with the central nervous system (Fig. 3.7), whereas more generalized sympathetic inhibitory reflexes pass through the CNS. It is certain that the two pathways can be both activated at the same time, and failure to recognize this leads to some experiments in vivo being quite difficult to interpret. Reflexes that arise from intestinal distension and, through noradrenergic nerves,
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contract sphincter muscle are included here as intestino-intestinal inhibitory reflexes because they oppose the normal movement of the digesta. General features of sympathetic entero-enteric regulatory reflexes Kuntz and his collaborators were the first to demonstrate that some intestinointestinal reflexes bypass the central nervous system (Fig. 5.15). They investigated the reflex in cats in which the prevertebral ganglia were decentralized and the intestine was transected between the points of stimulation and recording (Kuntz 1940, Kuntz & Van Buskirk 1941, Kuntz & Saccomanno 1944). Distension of the distal ileum or of the colon caused inhibition of the proximal ileum and application of nicotine to the celiac ganglia (thus blocking excitatory transmission) prevented the reflex (Kuntz & Van Buskirk 1941). Following criticism that the decentralization might have been incomplete, Kuntz and Saccomanno (1944) reported experiments that leave no doubt that the reflex was entirely peripheral. The authors performed one series of experiments on animals in which the entire spinal cord distal to the lower cervical region was removed (in some cases the vagi were also divided) and a second series in which the inferior mesenteric ganglia were decentralized by bilateral removal of the lumbar sympathetic chains and section of the celiac roots and hypogastric nerves a week or more before the experiment. At the time of the experiment, the colon was transected between the point of distension in the distal half and the recording point in the proximal half. In both series, the intestino-intestinal inhibitory reflex could be successfully elicited. A similar reflex was also demonstrated in the small intestine. Semba (1954a) initiated a similar reflex inhibition of the small intestine by the application of hypertonic solution (5% saline) to an adjacent, but unconnected, segment. In his experiments, the reflex was unaffected by adrenalectomy, vagotomy, preganglionic sympathectomy or destruction of the spinal cord below T1, but was eliminated when nicotine was applied to the celiaco-mesenteric plexus. Reflex inhibition of bile flow that is observed when the jejunum is distended is mediated through an entero-biliary pathway passing through the celiac ganglia (Kuntz & Van Buskirk 1941). Kuntz (1938) reported on experiments in which he severed different sources of axons that run to the celiac ganglia in the cat and had allowed time for their nerve endings to degenerate. When he cut the splanchnic nerves and removed the lumbar sympathetic chains, in some regions of the ganglia there was disappearance of the axon endings innervating most neurons, whereas in other regions there were groups of neurons with innervation still intact. Additional lesion of the vagus nerves caused no greater loss of synaptic inputs. Furthermore, if the nerves running from the celiac ganglia to the stomach and intestines were severed, degenerating fibers were found
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Fig. 5.15 The experiments that Kuntz (1940) used to demonstrate the existence of intestinointestinal reflexes that bypass the central nervous system. In one series of experiments, Kuntz lesioned the nerve connections between the central nervous system and the inferior mesenteric ganglion, effectively eliminating inputs to the ganglion that arise from the lumbar colonic and intermesenteric nerves (1 in diagram). When the nerve endings had degenerated, there was a loss of all innervation of some neurons. When he cut the colonic nerves (2), he found nerve fibers in the peripheral stumps (3) that did not degenerate. He deduced that these were axons of enteric neurons. In separate experiments, Kuntz distended a distal segment of colon that was connected to the proximal segment only through the inferior mesenteric ganglion (panel A). When Kuntz distended the distal segment between the arrow and x, the regular contractile waves in the proximal segment were interrupted (panel B). Adapted from Kuntz (1940).
on the central (celiac ganglion) side of the lesions. Kuntz concluded that all celiac ganglion neurons are multiply innervated, that a large proportion receive innervation from pre-ganglionic sympathetic neurons and that some receive innervation from fibers arising in the enteric plexuses (that is, from
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the stomach and intestine). In a final sentence, Kuntz wrote: “In view of the numbers of synaptic connections in the celiac ganglia which persist following degeneration of the splanchnic and vagus nerve fibers, these ganglia must be regarded as reflex centers of considerable functional significance.” Since that time, many structural studies have confirmed that neurons in the gut make synaptic connections in prevertebral ganglia (Kuntz 1940, Ungváry & Léránth 1970, Costa & Furness 1983, Dalsgaard et al. 1983, Macrae et al. 1986, Lindh et al. 1988, Gibbins et al. 2003). The intestinofugal neurons contain peptides, for example VIP, that can be used to trace their connections (Macrae et al. 1986, Lindh et al. 1988). Ross (1958) estimated that 30% of the axons in nerves of the mesentery of the small intestine arise from nerve cells in the gut wall. In experiments similar to those of Ross, Schofield (1960) also found surviving axons in the peripheral stumps and, in addition, found signs of retrograde degeneration in neurons of the myenteric plexus, but not in neurons of the submucosa. Properties of the intestinofugal neurons Retrograde tracing experiments indicate that cell bodies of intestinofugal neurons in small mammals are in myenteric ganglia. Most of the neurons have Dogiel type I morphology, but some have Dogiel type II morphology (Kuramoto & Furness 1989, Messenger & Furness 1992, 1993, Ermilov et al. 2003). Activation of the intestinofugal neurons by distension or electrical stimulation causes fast EPSPs in the neurons with which they synapse in prevertebral ganglia (Crowcroft et al. 1971, Szurszewski & Weems 1976, Kreulen & Szurszewski 1979). The fast EPSPs are blocked by nicotinic receptor blockers, indicating that the intestinofugal neurons are cholinergic (Szurszewski & Miller 1994). This is supported by immunohistochemical data that show that the neurons are immunoreactive for choline acetyltransferase (Mann et al. 1995). Since the first study of synaptic events in the inferior mesenteric ganglia of the guinea-pig that arose from the activation of intestinofugal neurons in the colon (Crowcroft et al. 1971) there has been uncertainty whether the intestinofugal neurons are first-order (primary) afferent neurons, or whether they are second-order neurons. First-order neurons would respond directly to distending or other sensory stimuli, whereas second-order neurons would be activated indirectly via synaptic transmission from first-order neurons (Szurszewski & Miller 1994). The neurons receive nicotinic fast EPSPs from other neurons in the gut wall, so they have connections necessary for a second-order role (Sharkey et al. 1998). Crowcroft (Crowcroft et al. 1971) reported that fast EPSPs occurred in neurons of prevertebral ganglia both spontaneously and when the colon was distended. Application of an antagonist of nicotinic fast EPSPs to the colon
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markedly depressed the spontaneous synaptic activity of colonic origin in the inferior mesenteric ganglia. Essentially similar results were obtained by others who investigated responses arising from the gut in the inferior mesenteric ganglia of guinea-pig (Szurszewski & Weems 1976, Bywater 1993), the celiac ganglion of the guinea-pig (Stebbing & Bornstein 1993), and the superior mesenteric ganglion of the mouse (Miller & Szurszewski 1997). In some of these experiments, low Ca2+ and elevated Mg2+ were used to block all synaptic transmission in the gut wall; the ongoing activity in the intestinofugal neurons was almost completely abolished (Bywater 1993, Stebbing & Bornstein 1993, Sharkey et al. 1998). It should be noted that in the experiments that reported spontaneous activity of intestinofugal neurons, there was an undistended balloon, or distending rods in the lumen, even when distension was not being applied. The luminal inserts have a mechanical action on the mucosa, which does not need to be intense to evoke reflexes (Smith & Furness 1988). In contrast to the effects of synaptic block on the spontaneous or ongoing activity of intestinofugal neurons, responses to distension are reduced, but not abolished (Crowcroft et al. 1971, Parkman et al. 1993, Miller & Szurszewski 1997), or are relatively little affected (Szurszewski & Weems 1976, Stebbing & Bornstein 1993). Bywater (1993) noted that the initial part of the response to distension was not significantly reduced by synaptic block, but the response after 10–15 s of sustained distension was almost abolished. In experiments on the inputs from the colon to the superior mesenteric ganglion in the mouse, Miller and Szurszewski (1997) obtained similar results; block of nicotinic transmission in the colon reduced spontaneous activity of intestinofugal neurons, and the sustained responses to distension were reduced to a greater extent than were the initial responses to distension. Thus it appears that sustained sensory stimulation activates the intestinofugal neurons indirectly via synaptic transmission from first-order (primary) afferent neurons and that phasic distension activates intestinofugal neurons directly. The inputs to intestinofugal neurons might come directly or indirectly from IPANs (Chapter 2). Although it is possible that there are two classes of intestinofugal neurons, one activated indirectly and one directly, Szurszewski and Miller (1994) have made the interesting suggestion that the same neurons may be both directly and indirectly activated. Thus, under some circumstances they may serve as second-order neurons of the intestinofugal pathway, in others as primary afferent neurons, and at times they may be activated in both ways. Sharkey (1998) reported that all intestinofugal neurons have similar morphologies and that they all receive large amplitude fast EPSPs. This is consistent with the hypothesis that there is one type of neuron which can be both directly and indirectly activated.
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Sympathetic entero-gastric reflexes via the celiac ganglia Reflex gastric inhibition involving a peripheral pathway through the celiac plexus occurs when the jejunal or duodenal mucosa is exposed to acid or the jejunal mucosa is exposed to hypertonic solutions. In the dog, this reflex is not significantly modified by vagotomy, section of the splanchnic nerves or removal of the thoraco-lumbar sympathetic chains, but it is abolished by application of nicotine to the celiac plexus, removal of the plexus or division of the mesenteric nerves (Semba 1954b, Schapiro & Woodward 1959). The reflexes have been examined in detail in normal human volunteers and in patients with various lesions, vagotomy for gastric ulcer, lumbar sympathectomy for vascular disorders, and two patients with thoracic spinal cord destruction because of tumors (Schapiro & Woodward 1955). In all cases, reflex gastric inhibition was observed when 1 M HCl was placed in the duodenum, but not when isotonic NaCl was instilled. Moreover, the reflex was seen when acid was placed in the jejunum in two patients in which the intramural pathways were interrupted by gastric resection and gastro-jejunal anastomosis (presumably to manage duodenal ulcer). Thus, the enterogastric reflex circuit consists of an afferent pathway which arises from the upper small intestine and synapses with the cell bodies of efferent inhibitory (noradrenergic) neurons in the celiac plexus. Some authors have reported a partial reduction of the reflex after vagotomy, which could arise because of the reduction in cholinergic activity for the noradrenergic nerves to act against (see section on mechanism of sympathetic inhibition above). The enterogastric reflex seems to delay gastric emptying principally by its inhibition of antral motility and is not significantly aided by constriction of the pylorus. Cannon et al. (1929) found that the celiac ganglia could be removed from cats without disturbing their digestive tracts, indicating that the reflex is not essential in undisturbed animals, in which acid entering the duodenum is presumably adequately neutralized by bicarbonate secretion from the duodenum itself and from the pancreas. In the guinea-pig we have cut all nerves running between the celiac ganglia and the stomach and small intestine, and have been able to confirm the completeness of the operation histochemically (Macrae et al. 1986). In over 40 animals we have seen no evidence of deleterious effects. Generalized (spinal) sympathetic inhibitory reflexes Inhibition can be caused by a more widespread irritation of the gut, such as in peritonitis (Youmans 1949, 1968). Bayliss and Starling (1899) reported that handling of the intestine caused a generalized inhibition of the intestine that was prevented by sectioning the extrinsic nerves (splanchnic and vagus), but was not influenced by severing intramural nerve pathways. Soon after,
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it was reported that when animals were alarmed movements of the stomach were inhibited, an effect that was prevented by splanchnic nerve section (Auer 1910). The phenomenon was studied in more detail by Pearcy and Liere (1926) who found that irritation of the colon mucosa with a bristle brush, or what they describe as adequate distension, caused inhibition of the movements of the stomach and small intestine. These and similar reflexes caused by noxious stimulation of intra-abdominal structures, including the gut, are abolished by section of the splanchnic nerves, by section of dorsal root fibers, by destruction or anesthesia of the spinal cord or by the inhibitor of noradrenergic transmission guanethidine (Morin & Vial 1934, Chang & Hsu 1942, Youmans et al. 1942, Hukuhara et al. 1959, Johansson & Langston 1964, Jansson & Martinson 1966). Similar effects can be elicited by capsaicin, which is a stimulant of spinal afferent neurons, placed in the intestine (Mizutani et al. 1990). The reflex centers are in the spinal cord, because the reflex is not eliminated when ascending and descending pathways in the cord are interrupted (Chang & Hsu 1942). Thus the afferent limb of the reflex is spinal afferent neurons that make their central connections in the spinal cord, and the efferent limb is sympathetic pre-ganglionic and post-ganglionic neurons. Anatomical and physiological studies both indicate that at least one spinal interneuron is included in this pathway (Foerster et al. 1933, Franz et al. 1966, Pomeranz et al. 1968). Stimulation of medullary depressor areas or of hypothalamic sympathoinhibitory areas activates descending fiber tracts which suppress the spinal intestino-intestinal inhibitory reflex (Johansson et al. 1965, 1968). This means that brainstem or spinal cord stimulation can increase intestinal motility by reducing the inhibitory action of noradrenergic nerves, and it is important not to confuse such effects with the direct stimulation of excitatory pathways. The blockade of the spinal intestino-intestinal inhibitory reflex by stimulation of the medulla was often accompanied by a fall in blood pressure, but this was not a necessary condition to observe the effect. Transection of the cervical cord reduced intestinal motility and decreased the threshold pressure needed to elicit the reflex, suggesting that descending pathways exert a tonic restraint on the spinal intestino-intestinal inhibitory reflex under experimental conditions (Johansson et al. 1968). Generalized inhibition can also be elicited by painful stimuli applied to other abdominal or extra-abdominal organs; these include mechanical stimuli applied to the urinary tract, peritoneum, skin, kidney, uterus, gall bladder and testes (Alvarez 1948, Youmans 1968). When the handling of the intestine is extensive, or peritonitis is induced, inhibition of motility following the operative procedure can be long-lasting. This condition is known as postoperative or adynamic ileus. It has a sympathetic reflex component, but is also contributed to by other mechanisms, particularly in persistent ileus, which is characterized
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by intestinal inflammation and is not relieved by blocking sympathetic transmission (Bauer & Boeckxstaens 2004). In its acute phase, postoperative ileus is relieved by sympathectomy or splanchnic anesthesia (Cannon & Murphy 1907, Ochsner et al. 1928, Douglas & Mann 1941) or by inhibition of transmission from the post-ganglionic noradrenergic neurons effectively (Neely & Catchpole 1971, Petri et al. 1971, Dubois et al. 1974). Spinal anesthesia or sympathetic blockade is less effective when adynamic ileus results from severe and widespread peritonitis (David & Loring 1930, Bauer & Boeckxstaens 2004). In particular, prolonged ileus involves an inflammatory infiltration of the external musculature and myenteric plexus region and is not relieved by blocking noradrenergic transmission (Bauer & Boeckxstaens 2004). Functions of the sympathetic inhibition of motility: conclusions Noradrenergic nerves controlling gastrointestinal movement are quite dispensible in animals exposed to a laboratory environment (Cannon et al. 1929), but when the nerves do act they suppress digestive action, by inhibiting the firing of enteric muscle motor neurons, by contracting the sphincters and, to a slight extent, by relaxing the muscle of the non-sphincter regions of the gut. The generalized spinal entero-enteric inhibitory reflex evoked by noxious stimuli such as laparotomy, irritation of the peritoneum, handling of the viscera, or stimulating of pain fibers could logically be thought to be protective. However, why the rapid inhibition of the intestine should be protective is not clear. Alvarez (1948) suggests that it may be useful to an animal in the wild if the noises from digestion are quelled when an animal is alarmed and in proximity to a predator. The inhibition may have evolved as a preparation for evasive action, in which the digestive process is suppressed and a greater percentage of the cardiac output can be diverted to other regions, such as skeletal and cardiac muscle, where an increased blood supply might be demanded. Neurons are more sensitive than is smooth muscle to lack of an adequate blood supply, so it may be significant that noradrenergic nerves supply the nerve cells rather than the muscle. On the other hand, the peripheral reflexes that pass through prevertebral ganglia and bypass the central nervous system appear to regulate the progress of the contents of the gut. The peripheral entero-gastric reflex contributes to limiting the development of acidity or hypertonicity of the contents of the upper small intestine by delaying gastric emptying. Summary and conclusions Intrinsic reflexes of the enteric nervous system are essential to the generation of the patterns of motility that are observed in the small and large intestines.
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The major muscle movements in the small intestine are mixing, propulsive reflexes that travel for only small distances, the migrating myoelectric complex, peristaltic rushes, and retropulsion associated with vomiting. The same neurons are involved in each of these, but how the nervous system is programmed to produce these different outcomes is not known. In contrast to the intestine, peristalsis in the stomach is a consequence of spontaneous electrical events (slow waves) in the muscle that begin in the corpus and travel to the pyloric sphincter. The intensity of contraction is determined by the actions of the vagus nerves, which form connections with enteric neurons in the gastric wall. The proximal stomach relaxes to accommodate the arrival of food. This relaxation is also mediated through the vagus. Thus, the primary integrative centers for control of gastric motility are in the brainstem, whereas those for control of the small and large intestines are in the enteric plexuses. Reflexes can be generated in the gastric enteric circuits, but their importance, relative to control through the vagus, is not clear. In most mammals, the contractile tissue of the external wall of the esophagus is striated muscle, and in others, including humans, the proximal half or more is striated muscle. The striated muscle part of the esophagus is controlled by an integrative circuitry in the brainstem. Thus, although the myenteric ganglia are prominent in the striated muscle part of the esophagus, they are modifiers, not essential control centers, for esophageal peristalsis. The smooth muscle sphincters restrict and regulate the passage of the luminal contents between regions. In general, reflexes that are initiated proximal to the sphincters relax the sphincter muscle and facilitate the passage of the contents, whereas reflexes that are initiated distally restrict retrograde passage of contents into more proximal parts of the digestive tract. The progress of the contents in an oral to anal direction is restricted when sympathetic nerve activity increases. To achieve this, transmission from enteric excitatory reflexes to the muscle is inhibited and the sphincters are contracted. Under resting conditions the sympathetic pathways exert little influence on motility. They come into action when protective reflexes are activated.
6: Enteric neurons and the physiological control of fluid secretion and vasodilation
In this chapter, the control of fluid secretion in the intestines, stomach, gall bladder, and pancreas is discussed. In the small intestine and colon, the adjustment of movement of water across the mucosal epithelium is crucial to the organism, and the orchestration of electrolyte movement, primarily of Cl–, is mainly directed at the regulation of fluid movement between the tissue and the lumen. When water and electrolytes are secreted across the epithelium, they are drawn from the tissue (interstitial) fluid adjacent to the inner surface of the epithelium. The interstitial fluid is in turn derived from absorbed fluid and from the circulatory system. Thus the control of local blood flow is closely linked to control of water and electrolyte movement. In the stomach, the important electrolyte that is actively secreted is the hydrogen (H+). The purpose of this secretion is the acidification and consequent partial breakdown of solid material in food. Regulation of blood flow in the stomach, and probably in the intestine, is also important for tissue protection. In the gall bladder, secretion modifies the composition of the bile, and secretion from the pancreas neutralizes the fluid in the jejunum and delivers digestive enzymes. In each case there is control through the enteric nervous system. Water and electrolyte secretion in the small and large intestines It is not easy to make a true estimate of the amount of fluid that enters the small intestine. Davenport (1977) calculated that 5–10 liters of water, derived from food and drink, salivary secretions, gastric secretion, secretions of the pancreas and biliary system, and from the intestine itself, enter the lumen of the human intestine each day. To maintain the equilibrium of fluids in the body, electrolyte that is absorbed in the gastrointestinal tract must be matched by losses through perspiration, respiration, urination, and defecation. The critical role played by regulation within the intestine itself is exemplified by the life-threatening effects of bacterial toxins, such as cholera toxin, which act on enteric neurons to cause copious secretion of fluid across the intestinal epithelium, and the deleterious effect of deficiency of fluid secretion in cystic fibrosis. 180
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The osmolarity of the fluid in the small intestine is closely matched to that in the interstitial fluid, which is to be expected (Lindahl et al. 1997, Lundgren 2002, Field 2003), although there are local variations from isosmolarity, for example in the lamina propria of the villi. Water moves from the lumen in company with the absorption of nutrient molecules (Schultz et al. 1966, Hines et al. 1995, Wright & Loo 2000). The major process for the absorption of nutrients, such as the monosaccharide glucose, is co-transport with Na+, in this case through the Na+/glucose cotransporter SGLT1. The entry of these solute molecules draws water across the epithelium, which is one rationale for adding glucose to oral rehydrating fluids. About 180 water molecules must move to balance the movement of one solute molecule, the water concentration being about 55 molar, and the concentration of dissolved ions in the intestinal lumen being about 300 mM (Lindahl et al. 1997). Direct measurements suggest that the co-transport is of one glucose and 210–260 water molecules (Wright & Loo 2000). Another way to consider the relationship between dissolved nutrient molecules and the amount of associated fluid is that 100 g of carbohydrate, if broken down to monosaccharides, would require about 1.8 liters of water to be isotonic with blood and tissue fluids. Amino acid transport is also ion-coupled and in a similar way involves water movement in concert with absorption. The large fluxes of water from the lumen to the interstitium are partly balanced by intestinal secretion that is mediated by secretomotor reflexes. It has been estimated that SGLT1 absorbs 8–9 liters of water each day from the small intestine (Wright & Loo 2000). As implied in the previous paragraph, the most efficient way to shift water molecules across the epithelium is to move solute molecules, and this is in fact how the great majority of water moves. A small proportion may pass through specialized water channels, the aquaporins, and a small proportion uses a paracellular route to cross the epithelium (Wright & Loo 2000). The secretomotor neurons stimulate the epithelial cells to pump Cl– into the lumen, which takes with it counter ions, mostly Na+, and water (see Chapter 3). A major component of Cl– secretion is through the cystic fibrosis transmembrane conductance regulator (CFTR) channel. It is also likely that secretion of K+ ions, which is stimulated by muscarinic agonists (Joiner et al. 2003), is increased by the activity of cholinergic secretomotor neurons. This would also be accompanied by water secretion. Secretion in the small intestine is primarily from the crypts and the bases of the villi, and in the colon it is from the deep parts of the glands; conversely, absorptive processes are dominant in the more superficial parts of the mucosa (Welsh et al. 1982, Lundgren & Jodal 1997). These functions are not mutually exclusive, and in fact both functions can occur at all levels of the mucosa (Geibel 2005). Consistent with a predominance of secretory function in the
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crypts, there is a gradient of CFTR expression from the deeper parts to the more superficial mucosa (Strong et al. 1994). Reflex control of water and electrolyte secretion There are two types of stimuli that cause secretomotor reflexes – benign stimuli, such as nutrient molecules, and noxious stimuli, such as those provided by cholera and enterotoxins and by rotavirus (Lundgren 2002). A primary function of the reflex response to benign stimuli is to return water and electrolyte to the lumen and to maintain the osmolarity and pH of the microenvironment adjacent to the mucosal epithelium. The benign secretion in the small intestine is linked to regulation of whole body water and electrolyte status (Furness & Costa 1987, Vanner & MacNaughton 2004; and see below). Secretion may also serve a role in mixing the fluid close to the absorptive epithelium. The primary function of responses to noxious stimuli is to rid the intestine of the offending sources of noxious irritation. The danger of the secretomotor response to noxious stimuli is that the secretion itself can be debilitating, even causing death. Thus a protective reflex can lead to a pathological outcome. Responses to benign stimuli When the intestinal lumen contains an isotonic salt solution that includes 30 mM glucose, net fluid absorption is about 60% of that when mannitol is present instead of glucose (Sjövall et al. 1983b, 1984a). This difference is due to an enteric reflex that enhances water and electrolyte secretion when glucose is absorbed. Its presence is revealed by local intra-arterial injection of tetrodotoxin, which decreases the secretory flux that occurs when the lumen is irrigated by glucose-containing physiological salt solution (Sjövall et al. 1984a). Destruction of the myenteric plexus also abolished the reflex (See & Bass 1993). The final secretomotor neuron in the enteric pathway revealed by these experiments was not cholinergic as the reflex occurred in animals treated with atropine (0.5 mg/kg). There is convincing experimental evidence that the non-cholinergic transmitter is VIP, although there are also cholinergic secretomotor neurons (Chapter 4). Enteric reflexes also cause bicarbonate secretion in response to duodenal acidification, although other acid-sensitive mechanisms, including a neurally independent stimulation of prostaglandin production, also release bicarbonate (Flemström 1994). Bicarbonate ions pass through the same channels as Cl–, the channels being similarly permeable to both ions (Poulsen et al. 1994). The ratio of Cl– to HCO3– that passes through these channels is affected by local pH and ion availability. Thus when the lumen is acidic, the channels
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pass a greater proportion of HCO3–. This property of the channels assists in maintaining a neutral pH at the luminal surface. The secretomotor activity in the intestine needs to be regulated in relation to fluid and electrolyte balance in the whole body. This is achieved primarily through an ongoing brake on secretomotor reflexes that is mediated through sympathetic efferent pathways. The role of sympathetic nerves in adjusting secretion Secretomotor reflexes appear to be always active in the small intestine, and to be held in check by the activity of sympathetic neurons. Evidence for a sympathetic brake on secretion comes from the old observation that severing the sympathetic nerves leads to copious secretion, which has been termed paralytic secretion. Removal of the celiaco-mesenteric ganglia (Bernard 1859) and sectioning the mesenteric nerves (Moreau 1868) in dogs both increase secretion in the intestine. Wright et al. (1940) found that section of pre-ganglionic sympathetic nerves caused increased secretion from the entire small intestine in decerebrate cats. The observation that watery diarrhea follows section of sympathetic pathways has been confirmed by more recent work (Freedman et al. 1953, Marlett & Code 1979). The activity in the secretomotor neurons is presumably maintained by intrinsic mechanisms, because the paralytic secretion is not influenced by vagotomy (Wright et al. 1940, Freedman et al. 1953). Consistent with these observations, secretion of fluid into the lumen of the intestine is reduced by electrical stimulation of the sympathetic pathways (Wright et al. 1940, Sjövall 1984), and this reduction is prevented by block of transmission from the sympathetic noradrenergic neurons with the α-receptor antagonist phentolamine (Brunsson et al. 1979). The noradrenergic sympathetic neurons innervate the submucosal ganglia, but provide very few fibers to the mucosa itself (Furness & Costa 1974) (Chapter 3). Activation of sympathetic noradrenergic neurons causes large amplitude inhibitory postsynaptic potentials (IPSPs) in the cell bodies of secretomotor neurons (North & Surprenant 1985) and also decreases excitatory transmitter release in the submucosal ganglia by a presynaptic action (Shen et al. 1990, Shen & Surprenant 1990). The IPSPs were blocked by antagonists of α-adrenoceptors. Thus sympathetic inhibition is of the secretomotor neurons, not directly of the enterocytes. This confirms that the secretomotor neurons have a basal activity that is reduced by inputs from sympathetic neurons. The site of action of the sympathetic neurons was investigated in elegant experiments by Sjövall (1983a,b), who found that stimulation of the sympathetic nerve fibers was more effective when secretomotor reflexes were elicited by glucose in the lumen or by cholera toxin, than when reflexes were not active. In the glucose-perfused intestine, splanchnic nerve stimulation
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had an anti-secretory effect, whereas, if all glucose was replaced with mannitol, splanchnic nerve stimulation had no anti-secretory action. Moreover, norepinephrine infusion into the vasculature of the small intestinal segment reduced the secretion when there was glucose in the lumen, but it was not effective if intramural nerves were blocked by tetrodotoxin (Sjövall et al. 1983b). The action of applied norepinephrine was also eliminated by substitution of luminal glucose with mannitol (Sjövall et al. 1984a). In addition, the effect of sympathetic nerve stimulation was blocked by the norepinephrine α-receptor blocker phentolamine. Taken together, these results indicate that the enteric secretomotor reflex evoked by glucose is inhibited by sympathetic nerve activity. The physiological role of sympathetic inhibition of secretomotor reflexes is to conserve blood volume, and hence tissue fluid volume. This role was hinted at by experiments by Mailman and colleagues between 1967 and 1975 (Mailman et al. 1967, Mailman & Ingraham 1971, Mailman & Jordan 1975). The authors found that movement of water and electrolyte from body fluids into the intestinal lumen was reduced by hemorrhage or head-upward tilting, to mimic postural hypotension (Mailman et al. 1967, Mailman & Ingraham 1971). On the other hand, increasing blood volume by intravenous infusion of isotonic saline increased secretion into the intestine, and intravenous hypertonic solution reversed this effect (Mailman & Jordan 1975). The sympathetic reflexes involved in modifying secretion in line with whole body water and electrolyte homeostasis were directly investigated by Sjövall et al. (1982), who mimicked the hypotensive effect of blood loss by unloading carotid sinus baroreceptors with occlusion of the carotid artery in rats and cats. This caused a reduction in fluid secretion in the small intestine which was prevented when the sympathetic pathway to the intestine was disrupted by breaking nerves following the superior mesenteric artery or by cutting the splanchnic nerves, but not by vagus nerve section (Sjövall et al. 1982, Redfors et al. 1984). Hemorrhage of about 20% of blood volume in rats caused an inhibition of secretion that was prevented by the α-receptor blocker phentolamine or by chemical sympathectomy using 6-hydroxydopamine (Levens 1984). Data from conscious human volunteers also indicates that cardiovascular changes alter intestinal secretion (Sjövall et al. 1986). Manipulation that caused blood pooling in the lower limbs, estimated to be equivalent to hemorrhage of 600–800 ml, decreased the secretion of fluid, Na+, and Cl– into the jejunum, but did not affect glucose transport. Further evidence for the involvement of sympathetic control of secretomotor activity in cardiovascular regulation comes from experiments in which signaling from cardiac volume receptors was manipulated (Sjövall et al. 1984b). When reduced cardiac filling was mimicked, secretion into the jejunum (which was artificially increased by cholera toxin in these experiments)
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was lowered. The effect was substantially reduced by vagotomy, cutting sympathetic pathways to the gut and by blocking α-adrenoceptors (Sjövall et al. 1984b). Vagotomy is effective because it interrupts the cardiac afferent signals. Secretion into the duodenum is alkaline, as a protection against duodenal erosion by gastric acid. This alkaline secretion is also mediated through enteric reflexes, and like secretomotor reflexes in the more distal small intestine, it is influenced through blood volume preserving reflexes (Jönson et al. 1990). A 10% reduction in blood volume reduced secretion of bicarbonate by 44%. The effect was prevented by vagotomy, implying that the reflex is initiated through cardiac volume receptors, whose axons travel in the vagus nerve, and was also blocked by the α-adrenoceptor antagonist yohimbine, implying that the effect was mediated through sympathetic, noradrenergic inhibition of secretomotor reflexes (Jönson et al. 1990). The sympathetic reflex inhibition that has been discussed above is mediated via reflex pathways that travel through the central nervous system. There is also functional and structural evidence for peripheral reflexes that pass via prevertebral ganglia (Chapter 3). Wright et al. (1940) cut the splanchnic nerves on both sides as well as all other visible inputs on the central side of the celiac ganglia in cats, and they also removed parts of the sympathetic chains in the region. The duodenal secretion that followed the operation subsided after 2 days and the cat was re-examined after 10 days, at which stage removal of the decentralized celiac ganglia resulted in copious secretion from the ileum. It is thus pertinent that the cell bodies of the neurons that project to submucosal ganglia in guinea-pigs receive an input from myenteric neurons (Macrae et al. 1986), that is, there is morphological evidence for a peripheral reflex that would inhibit enteric secretomotor neurons (see Chapter 3). Intrinsic restraint on secretomotor neurons by the local hormone PYY Among the most potent endogenous inhibitors of secretion are the peptides NPY, PYY, and PP (Saria & Beubler 1985, Cox et al. 1988, Bilchik et al. 1993). PYY is contained in entero-endocrine cells, which are most numerous in the lining epithelium of the distal small intestine and colon (Ekblad & Sundler 2002), and it is released in the 1–2 hours following a meal and in response to intraluminal nutrients, especially lipid and bile salts (Onaga et al. 2002). Amongst these peptides, PYY acts preferentially on Y1 and Y2 receptors. However, it is more potent than the other peptides at Y2 receptors. In the isolated colonic mucosa of the mouse, a Y2 receptor antagonist increased secretion, indicating that endogenous ligand for the receptor was being continuously released (Hyland et al. 2003). Tetrodotoxin blocked the increase in secretion caused by the antagonist and reduced the antisecretory
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effect of the Y2 receptor agonist PYY(3–36). These data indicate that PYY probably exerts a major component of its antisecretory effect by inhibiting enteric secretomotor neurons. In tissue from a receptor Y2 knockout mouse, neither the Y2 receptor agonist nor the antagonist had any effect (Hyland et al. 2003). In isolated human and mouse colon mucosa, Y1 and Y2 receptor antagonists both increased secretion, indicating that these receptors are activated by locally released peptides in isolated tissues (Cox et al. 2001, Cox & Tough 2002, Hyland et al. 2003). A physiological restraint on secretion in vivo is suggested by experiments in which infusion of PYY, to achieve levels that are observed postprandially, increased fluid absorption in the canine small intestine (Bilchik et al. 1993). Secretory responses to toxins and irritants The body exerts a number of acute physiological responses to the presence of toxins and pathogens in the gastrointestinal tract. These include vomiting and diarrhea. Diarrhea has components of expulsive motility of the distal ileum and colon and of secretion of fluid into the lumen of the small intestine and colon. Fluid secretion when the intestinal lumen is confronted by toxins and irritants is a protective response. However, if the body is unable to manage the luminal threat, the copious secretion that ensues can endanger life. Secretory diarrheas claim millions of lives annually, primarily in tropical countries where pathogens are common and health-care measures are inadequate (Field 2003). In more affluent countries, secretory diarrhea is still common, particularly in infants and the elderly. Secretory diarrhea is also a problem in animal populations, including agricultural animals (Jones & Blikslager 2002). Toxin-mediated reflex secretion probably occurs at a low level in healthy individuals, who always have some presence of bacterial toxin in the lumen of the intestine. For the same reason, the intestinal mucosa exhibits a low level of inflammation at all times. Although it is clearly demonstrated experimentally that cholera toxin, heat-stable enterotoxin and rotavirus, and other pathogens and their toxic products cause secretomotor reflexes (Lundgren et al. 2000) (Chapter 3), the responses to these noxious agents involve many components (Cooke 1994, Jones & Blikslager 2002, Field 2003). These include the direct effects of toxins on enterocytes to increase secretory flux, stimulation of prostaglandin release from the affected epithelial cells, which have both direct effects to stimulate secretion by the enterocytes and indirect effects through stimulation of secretomotor neurons (Kimberg et al. 1971, Argenzio et al. 1996, Dekkers et al. 1997). 5-HT that is released by cholera toxin from enterochromaffin cells (Chapter 3) can trigger prostaglandin release (Beubler et al. 1989). Pathogens also stimulate cells of the mucosal defense system, including mast
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cells, lymphocytes, and macrophages, which release a variety of pro-secretory products, including histamine, leukotrienes, platelet-activating factor, prostaglandins, and interleukins (Field 2003). As might be expected of a noxious stimulus, bacterial toxin in the gut also activates extrinsic, capsaicin-sensitive afferent nerve endings (Rolfe & Levin 1997). The most urgent therapy for life-threatening secretory diarrheas is rehydration, usually by the oral route. However, oral rehydration therapy does not by itself prevent the diarrhea, and targeting the enteric nervous system, or its transmitter receptors, may be valuable as an adjunct therapy (Chapter 7). Can secretion and increased blood flow have a tissue-protective role? Mechanical stimulation of the mucosa, for example by a small brush, provokes hyperemia and secretion that are mediated through enteric reflexes (Biber et al. 1971, Sidhu & Cooke 1995, Vanner & Surprenant 1996). It is difficult to know the reason for these reactions to the stimulus, but it is possible that the abrasion of the delicate mucosa causes a tissue-protective inflammatory reaction, and by implication that mechanical abrasion can be a noxious stimulus. This idea is supported by the observation that part of the vasodilation is caused by release of histamine (presumably from mast cells) and of prostaglandins (Vanner et al. 1993). Secretion that is elicited by distension is also mediated in part by release of prostaglandins (Diener & Rummel 1990). In a model of chemical peritonitis induced by HCl, ethanol, bile, or bile salts applied to the serosal surface of the intestine, secretion (and presumably hyperemia) was also provoked through an activation of enteric secretomotor neurons, with involvement of both histamine and prostaglandin release (Brunsson et al. 1985, 1990). Both these reactions may have a role in increasing nutrient availability and removing noxious substances and the by-products of tissue damage. Tissue protection through increased mucosal blood flow has been clearly demonstrated in the stomach (see below), so it would not be surprising if the dilation caused by activity of enteric secretomotor/vasodilator neurons contributes to reducing intestinal mucosal damage. Gastric protection and gastric mucosal hyperemia Neurotransmitters released from the peripheral ends of extrinsic primary afferent neurons have significant effects in restricting the deleterious consequences of tissue damage in the stomach (Holzer 1998). Compromising the function of extrinsic gastric afferents by pretreating animals with capsaicin aggravates gastric mucosal damage caused by acid, ethanol, non-steroidal anti-
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inflammatory agents, and other chemicals (Holzer & Sametz 1986, Holzer et al. 1991, Uchida et al. 1993, Brzozowski et al. 1996). Capsaicin alone is not injurious. In fact, the short-term effect of capsaicin is protective, because it releases transmitter with a protective role from the nerve endings (Holzer 1998). CGRP released from the afferent nerve endings has a major role in the protection of the mucosa. Thus blocking CGRP receptors, or reducing its effectiveness by immunoneutralization, compromises gastric protection (Lambrecht et al. 1993, Merchant et al. 1995). Capsaicin applied to the spinal afferent pathways, but not capsaicin applied to the vagus nerves, 10–20 days before experiment, blocks the CGRP-dependent hyperemic response to acute application of capsaicin, indicating that gastric protection is dependent on the integrity of spinal afferent neurons (Li et al. 1991). Because the protection is reduced by the nerve conduction blocker tetrodotoxin applied selectively to the stomach, it probably involves axon reflexes (action potential invasion of collateral branches of activated nerve endings) (Holzer et al. 1991). Tachykinins are often co-localized in the gastric spinal afferent neurons, and tachykinins, released from the spinal primary afferent endings and acting through NK2 receptors apparently contribute to gastric protection (Stroff et al. 1996). The protective mechanism involves vasodilation and increased mucosal blood flow. CGRP and tachykinins cause vasodilation by releasing nitric oxide (NO) from vascular endothelial cells, and thus inhibition of NO synthase increases gastric damage (Whittle et al. 1990). Transmitter release from afferent endings in the colon also reduces the severity of damage consequent on inflammation, at least in the acute phase (Reinshagen et al. 1996). Balancing the supply and the secretion of water and electrolyte The presence of three classes of secretomotor neurons, two of which also provide vasodilator collaterals (Chapter 3), may provide a mechanism to balance secretion and vasodilation appropriate to the digestive state. As previously alluded to, for water and electrolyte equilibrium to be maintained, the amount of fluid lost or excreted should be matched by absorption from the alimentary tract. If more fluid is absorbed with nutrients or across the gastric mucosa, some can be passed back into the lumen under the control of secretomotor reflexes. Thus the source of secreted fluid in the small intestine is a mixture of serum electrolyte and locally absorbed electrolyte (Fig. 6.1). It is suggested that if sufficient fluid that is co-absorbed with nutrients is available, then the secretion will be dominated by activity of the cholinergic secretomotor neurons that do not have collaterals to the arterioles. On the other hand, if more fluid is required than can be supplied by absorbed electrolyte, there is activation of secretomotor/vasodilator neurons.
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Fig. 6.1 Schematic representation of the integration of secretomotor and vasomotor control in the small intestine. The circuitry provides for adjustment of the secretion of fluid and electrolyte into the intestinal lumen, according to the sources of the fluid and the needs of the whole body to maintain fluid homeostasis. The sources of fluid to supply the secretory flux are the fluid that is absorbed across the epithelium in company with nutrients and ions and fluid from the circulation. The existence of both secretomotor only (neuron 14) and secretomotor/vasodilator neurons (neurons 12, 13) allows the relative supply from these two sources to be adjusted. The enteric secretomotor reflexes are under tonic inhibitory control from the sympathetic neurons that innervate secretomotor neurons and the availability of fluid from the circulation is limited by activity of sympathetic vasoconstrictor neurons. If blood pressure or blood volume decreases (e.g. in hemorrhage) secretion and blood flow to the mucosa are both inhibited. The numbering of neurons corresponds to the listings in Fig. 2.1 and Table 2.1.
This intrinsic secretomotor/vasodilator system is governed by the central nervous system in two ways, through sympathetic inhibition of secretomotor/ vasodilator neurons, as explained above, and by sympathetic vasoconstrictor neurons that innervate arterioles in the gut wall (Fig. 6.1). Secretion of gastric acid Three natural stimulants have co-operative roles in causing acid secretion from the parietal cells: acetylcholine, released from the terminals of enteric neurons, gastrin, released from antral endocrine cells, and histamine, released from enterochromaffin-like cells (ECL cells) of the lamina propria close to parietal cells (Fig. 6.2). Receptors for each of these substances, which act synergistically,
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Fig. 6.2 Neural and hormonal control of gastric acid secretion. The acid-secreting parietal cell is stimulated by ACh released from enteric neurons and histamine from enterochromaffin-like (ECL) cells. The action of gastrin is primarily indirect, through stimulation of histamine release from ECL cells. Enteric neurons also innervate the G cells, and the adjacent somatostatin-secreting D cells. The primary transmitter of neurons innervating the G cells is gastrin-releasing peptide. For further details, see text.
are on the parietal cells in dog and human (Gillespie & Grossman 1964, Soll & Walsh 1979). There is ongoing (basal) release of acetylcholine, gastrin, and histamine and reduction in any one of these components tends to diminish acid secretion (Del Valle & Todisco 2003). In rats, and probably also in mice and rabbits, the major action of gastrin is indirect, through stimulation of histamine release from the ECL cells (Lindstrom et al. 2001), whereas in human it is likely that there are both direct and indirect effects of gastrin (Kulaksiz et al. 2000). In contrast to parietal and gastrin cells, the ECL cells do not seem to be under direct vagal control (Norlén et al. 2005). Histamine release in response to a meal, or in response to electrical stimulation of the vagus, was largely prevented by blocking gastrin receptors (Norlén et al. 2005). Thus the vagus acts to stimulate the release of gastrin, gastrin reaches the ECL cells through the circulation,
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gastrin increases histamine secretion from the ECL cells, and histamine acts synergistically with nerve-released ACh to stimulate the parietal cells. The vagus nerve increases the secretion of acid both via an excitatory pathway that impinges on the parietal cells (Olbe 1964, Knutson & Olbe 1974) and through the nerve-mediated stimulation of gastrin release (Nilsson et al. 1972, Farooq & Walsh 1975, Uvnäs Wallensten & Andersson 1977). In all cases where the vagus nerve is activated and gastric acid secretion is increased, the effect is diminished by hexamethonium, or other antagonists of ganglionic transmission. This presumably means that the final motor neurons both to the parietal cells and to the gastrin cells are in the gastric enteric ganglia, although some of the cell bodies could be in small ganglia that are found along the vagus nerve. Neural control of gastrin release Gastrin release in response to direct or reflex stimulation of the vagus is atropine-resistant in dogs (Smith et al. 1975, Dockray & Tracy 1980). In these experiments, antral pH was held constant at pH 6–7 to prevent changes in luminal acidity influencing gastrin release. In cats, gastrin release following electrical vagal stimulation is also unaffected by atropine (Uvnäs Wallensten & Andersson 1977), and in humans, inhibition of muscarinic receptors for acetylcholine does not diminish gastrin secretion induced by a meal (Walsh et al. 1971). Thus the vagal stimulation of gastrin release is not mediated through acetylcholine acting at muscarinic receptors. The non-cholinergic, nerve-mediated, stimulation of gastrin release is via a final neuron that utilizes gastrin-releasing peptide (GRP) as a neurotransmitter. GRP is a potent stimulant of gastrin release, by its direct action on gastrin cells (Basso et al. 1974, DuVal et al. 1981, Richelson et al. 1983). GRP occurs in nerve fibers but not in endocrine cells of the gastric mucosa (Dockray et al. 1979) and the GRPimmunoreactive nerve fibers are found close to the gastrin cells (Holst et al. 1987, Miller et al. 1989, Sjövall et al. 1990). Desensitization of GRP receptors abolishes the release of gastrin from the porcine stomach in response to vagus nerve stimulation (Holst et al. 1987) and a GRP receptor antagonist blocked the gastrin release caused when a vago-vagal reflex was induced by gastric distension (Weigert et al. 1997). Consistent with these observations, antisera to GRP added to the vascular perfusate of the rat stomach reduced the gastrin secretion caused by vagus nerve stimulation (either electrical or by DMPP) by 60% (Schubert et al. 1985). In rat there may be a cholinergic component of transmission to the gastrin cells, as atropine further reduced gastrin output; the atropine and anti-GRP effects were additive (Schubert et al. 1985).
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The cephalic component of control of gastric acid secretion Vagal stimulation of acid secretion can be induced by sham feeding, sight, or even the thought of food. This cephalic component of acid release is prevented by vagus nerve section (Pavlov 1902), and involves the vagal stimulation of enteric neurons that excite the parietal cells and of other neurons that cause release of gastrin (Feldman & Richardson 1986). If the vagal branches to the antrum are intact, increases in both serum gastrin and in acid output are observed, whereas if the antrum is denervated, plasma gastrin does not rise but there is still a significant, although diminished, increase in acid production (Tepperman et al. 1972). In humans, preventing gastrin involvement in the response by antrectomy reduces the acid secretory response to sham feeding by about 50% (Knutson & Olbe 1974). The role of gastrin in the response to sham feeding is further demonstrated by the diminution of the response when anti-gastrin antibodies are used to neutralize this hormone (Kovacs et al. 1997). Atropine abolishes acid secretion in response to sham feeding in dogs (Nilsson et al. 1972) and greatly reduces responses in humans (Feldman et al. 1979b). This suggests that the acetylcholine that acts on parietal cells has a potentiating (permissive) role in gastrin’s action and that when the action of acetylcholine on the parietal cells is blocked gastrin is very much less effective. This is confirmed by the observation, in humans, that if the acid-secreting portion of the stomach is vagally denervated, but the innervation of the antrum is intact, sham feeding causes a substantial rise in plasma gastrin, with no increase in acid output (Feldman et al. 1979a). These observations are consistent with observations on isolated parietal cells. When gastrin plus histamine, or muscarinic agonists (to mimic acetylcholine) plus histamine are given together, the release of acid is greater than the sum of responses of the two given separately (Soll 1981). Moreover, the effects of carbachol plus gastrin, if given in the presence of histamine, are more than additive. The gastric component of control of gastric acid secretion The gastric phase of acid secretion is evoked when food enters the stomach and can be elicited experimentally by gastric distension. Studies with antral and fundic pouches point to the existence of vago-vagal reflexes, initiated by gastric distension, that cause both gastrin and gastric acid release (Debas et al. 1974). Acid secretion in response to distension is almost completely abolished by vagotomy and by atropine (Debas et al. 1974, Grotzmyer et al. 1977, Schiller et al. 1980). The release of gastrin by distension is also primarily through a vago-vagal reflex (Weigert et al. 1997). Gastrin release caused by graded gastric distension is prevented by acute truncal vagotomy or chronic
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denervation using capsaicin (Weigert et al. 1997). Thus the final mechanisms for the gastric phase of acid secretion are similar to those for the cephalic phase, and both depend on the vagus. The gastric phase is also contributed to by the chemical nature of the food – peptic digests of protein, ethanol, and caffeine are all activators of the gastric phase (Del Valle & Todisco 2003). Cholinergic innervation of somatostatin cells in the antrum An inhibitory action of cholinergic nerves on gastrin release is suggested by investigation of responses to distension, sham feeding, or insulin hypoglycemia in humans (Farooq & Walsh 1975, Feldman et al. 1979b, Schiller et al. 1980). In each case, atropine causes an increase in the secretion of gastrin. These effects are likely to be mediated by the block of cholinergic inhibitory transmission to somatostatin cells in the antral mucosa, thus reducing the effectiveness of a somatostatin “brake” on release of gastrin (DuVal et al. 1981, Schubert et al. 1982, Schubert & Makhlouf 1982, Saffouri et al. 1984, Lloyd 1994). This conclusion is based on the following observations. Somatostatin is contained in gastric endocrine cells in intimate association with the gastrin cells (Larsson et al. 1979, Larsson 1984). Somatostatin inhibits gastrin release from the antral mucosa, and its site of action is within the antral mucosa (Harty et al. 1981). Moreover, if anti-somatostatin antibodies are added to the arterial inflow to the rat stomach, the concentration of gastrin in the venous outflow is increased (Short et al. 1985). Electrical stimulation of intramural nerves of the antrum reduced somatostatin release, and this reduction was blocked by atropine, in the presence of which electrical stimulation caused enhancement of release (Schubert et al. 1982). Release of somatostatin is inhibited by acetylcholine and this inhibition is blocked by atropine (Koop et al. 1982). Are there intrinsic gastric reflexes controlling acid secretion? The data appear to leave little room for the existence of an intrinsic reflex that elicits acid release, despite the fact that there are clearly internal connections within the gastric enteric nervous system (Schemann et al. 2001). In fact, the reliance on the vagus and a lack of capacity of intrinsic circuits to take over is indicated by the persistent reduction in acid secretion that follows vagotomy in humans (Dragstedt 1945, O’Leary et al. 1976) and experimental animals (Hirschowitz & Hutchison 1977). The only way in which extrinsic neural or hormone influences can be eliminated with certainty, so that intramural reflexes could be examined, is to use a preparation of isolated perfused stomach. Intraluminal protein increases gastrin and decreases somatostatin in the venous drainage from the isolated perfused rat stomach (Saffouri et al. 1984,
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Schubert et al. 1992). The responses were blocked by tetrodotoxin. Atropine reduced the increase in gastrin, and atropine plus a GRP receptor antagonist abolished the release of gastrin. Atropine also blocked the inhibition of somatostatin release. In fact, in the presence of atropine, intraluminal protein caused an increase in somatostatin release, supporting other data that indicate that there are cholinergic neurons that inhibit somatostatin release and non-cholinergic neurons that enhance release (see above). In contrast to the effects of a peptic digest of protein, gastric distension decreased gastrin secretion from the isolated stomach (Weigert et al. 1997). The effect was reduced by atropine and by tetrodotoxin to a similar extent. These data indicate that there are intraluminal reflexes that control gastrin and somatostatin release, but their physiological importance is difficult to judge. The intestinal component of control of gastric acid secretion There is also an intestinal component of the control of acid secretion. Both augmentation and reduction of gastric acid secretion can arise from duodenal stimulation. Augmentation is recognized as the intestinal phase of gastric secretion and was described in response to food in the duodenum by Leconte (1900). Distension of the duodenum can also initiate gastric acid secretion (Konturek et al. 1978). Instillation of concentrated fat, acid, or hyperosmolar solutions into the duodenum all reduce gastric acid secretion. However, the principal mechanism through which duodenal acidification is neutralized is by the stimulation of bicarbonate release by the hormone secretin, released from entero-endocrine cells in the duodenum that are stimulated in an acid environment (Johnson & Grossman 1969). Pepsinogen secretion The protease pepsin is found in the stomach following food ingestion, as Langley and Edkins described (Langley & Edkins 1886). They recognized that the secretion was of the pro-enzyme pepsinogen, which is cleaved in the gastric lumen to yield the active enzyme pepsin. Pepsinogen is released primarily from chief cells at the bases of the fundic glands, generally at the same time as gastric acid is secreted (Del Valle & Todisco 2003). Electrical stimulation of the central cut end of one vagus, the other vagus being intact, releases pepsin into the stomach (Harper et al. 1959) and distension of a vagally innervated gastric pouch causes pepsin release into the pouch (Grossman 1962). These observations indicate that there is nerve-mediated stimulation of pepsin release via the vagus and point to the existence of a vago-vagal reflex. Vagal stimulation increases acid output, mucosal blood flow, and contractile activity, as well as pepsin secretion. Concomitant stimulation of the splanchnic nerves
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reduced acid output, mucosal blood flow, and motility, but pepsin release was unaffected (Reed & Sanders 1971). Moreover, splanchnic stimulation alone did not increase pepsin output. The observations suggest an independence of the vagal pathways that stimulate pepsin release from those that cause acid production. The pepsin release in response to vagal stimulation is only partly blocked by a high dose of atropine; in the presence of atropine, vagal nerve stimulation increased release three-fold (Skak-Nielsen et al. 1988). GRP is a stimulant of pepsin release that acts directly on the pepsin-secreting chief cells (Fiorucci & McArthur 1990) and is contained in nerve fibers that surround these cells, and is thus a possible final transmitter to the chief cells. Gastric secretion of bicarbonate Bicarbonate that is secreted by the gastric mucosa and is trapped in the mucus at the surface of the gastric lining is an important part of the gastric defense against erosion. Some of the HCO3– arises as a by-product of the secretion of H+ by the parietal cells (and is delivered to the surface of the stomach by a vascular route), and part comes from the surface epithelial cells that contain the enzyme carbonic anhydrase, which enhances its formation. Because of its link to acid secretion, it is difficult to determine whether there is an independent control of HCO3– release by neurons. However, it is secreted into the gastric lumen when the vagus is stimulated (Forssell et al. 1985) or following sham feeding (Feldman 1985). Secretion can be induced by transmural stimulation and by muscarinic agonists in isolated sheets of antral mucosa, consisting of the mucosal plus submucosal layers (Suzuki et al. 1993). Nerve-mediated secretion was inhibited by atropine. Secretion into the gall bladder The gall bladder concentrates the bile salts by reabsorbing water and electrolyte primarily through coupled transport of Na+ and Cl– (Wood & Svanvik 1983). However, the bile remains approximately isotonic. At the same time, there is secretion of HCO3– into the gall bladder lumen, which makes the bile slightly alkaline. The gall bladder can also secrete water and electrolyte into its lumen, which may be observed as a decrease in net absorptive flux, or a reversal of absorption to secretion. The gall bladder mucosa is densely innervated by VIP-containing nerve fibers and VIP-immunoreactive nerve cells occur in the intrinsic gall bladder ganglia (Sundler et al. 1977, Keast et al. 1985b, De Giorgio et al. 1995). VIP reduces absorption in a dose-dependent manner, and can reverse absorption to secretion (Jansson et al. 1978, O’Grady et al. 1989). There is some evidence that intrinsic reflex pathways can control secretion through the VIP-containing secretomotor neurons. The
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secretion that is induced by experimental cholecystitis is not influenced by extrinsic denervation, but is strongly reduced by blocking action potential conduction in neurons by local intra-arterial tetrodotoxin, or by intraluminal application of local anesthetic (Jivegard et al. 1987). Hexamethonium also reduced secretion, implying that a reflex pathway with nicotinic synapses was involved. The secretion was not blocked by atropine, which indicates that non-cholinergic secretomotor neurons are activated (Jivegard et al. 1987). However, it was abolished by infusion of anti-VIP antiserum, but not by nonimmune serum infused into the celiac artery, which supplies the gall bladder (Nilsson et al. 1994). Vagal nerve stimulation also activates the VIP-containing secretomotor neurons of the gall bladder, and causes VIP release from the gall bladder (Bjorck et al. 1983, 1986). Similar to the small and large intestines, activation of sympathetic noradrenergic neurons innervating the gall bladder increases absorption/inhibits secretion (Björck et al. 1982, O’Grady et al. 1989). Pancreatic exocrine secretion Exocrine secretion from the pancreas into the duodenum has two components – secretion that is rich in digestive enzymes and an alkaline secretion. These are independently controlled, the enzyme secretion being stimulated by CCK and bicarbonate production by secretin, both hormones being released from the duodenum (Holst 1990, Owyang 1996). The two forms of secretion appear to be under independent neural control (Hickson 1970, Holst 1990). Both are elicited by vagus nerve stimulation, and hexamethonium blocks both forms of secretion, suggesting that both pathways have synapses in pancreatic ganglia. However, atropine blocks the enzyme secretion, while the alkaline secretion is not affected or only partly reduced. Another difference is that the physiological effect of secretin is largely direct, whereas CCK’s action is via vago-vagal reflexes and enteric neurons. Pancreatic alkaline secretion The hormone secretin is the major factor by which acid conditions in the duodenal lumen trigger bicarbonate secretion from the pancreas (Bayliss & Starling 1902), and there is good evidence that secretin acts directly on cells of the intralobular and small interlobular ducts to stimulate Cl– and HCO3– secretion (Owyang & Williams 2003). Secretin is released when the duodenal or jejunal lumen is acidified and causes secretion from the entirely extrinsically denervated pancreas (Bayliss & Starling 1902). Nevertheless, vagus nerve stimulation does cause secretion. In pigs, the secretion of bicarbonate in response to vagus nerve stimulation is atropine resistant (Holst et al. 1979).
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It is accompanied by release of VIP into the venous drainage of the pancreas, and is mimicked by VIP (Fahrenkrug et al. 1979), which is contained in nerve cells in the pancreatic ganglia and in nerve fibers that innervate the pancreatic acini (Holst et al. 1984). Thus there is evidence for both a direct hormonal and a non-cholinergic neuronal contribution to pancreatic bicarbonate secretion. In addition, in dogs, atropine reduces the bicarbonate release in response to secretin infusion, suggesting that secretin’s action is partly through stimulation of cholinergic neurons or there is a synergistic effect of cholinergic transmission and secretin (Singer et al. 1980). Although secretin was still effective in eliciting secretion from the transplanted or vagally denervated pancreas, the reduction when atropine was administered was not observed, suggesting that the action of ACh is indeed a synergism, and that the source of acetylcholine is nerves that are activated through an entero-pancreatic, vago-vagal reflex. Pancreatic enzyme secretion Pancreatic enzyme secretion has cephalic, gastric, and intestinal phases. Each appears to require mediation through the vagus, which points to an integration of the responses in the CNS, probably in the medulla oblongata. The cephalic phase can be investigated by sham feeding. In humans, the secretion of pancreatic enzyme in response to sham feeding is blocked by atropine, indicating that there is a vagal excitatory innervation of pancreatic ganglia (Anagnostides et al. 1984). The gastric phase of secretion appears to be minor; it can be elicited by gastric distension (Cargill & Wormsley 1979). The intestinal phase is elicited by products of the digestion of protein and fats that evoke CCK release from duodenal entero-endocrine cells. The CCK indirectly triggers the release of enzyme-rich secretion from the pancreas, as explained below. CCK also potentiates the stimulation of bicarbonate secretion by secretin. CCK or nutrient-stimulated pancreatic secretion is substantially reduced by atropine in both human and dog (Thomas 1964, Konturek et al. 1972, Adler et al. 1991, Soudah et al. 1992). In humans, the pancreatic enzyme secretion in response to a meal is almost completely blocked by atropine, but the plasma levels of CCK are unaffected (Adler et al. 1991). Moreover, also in humans, when CCK was infused intravenously at a rate that mimicked the levels seen after a meal, atropine reduced the secretion of enzyme from the pancreas by about 80% (Soudah et al. 1992). Responses elicited by supra-physiological concentrations of CCK were less sensitive to atropine. In vitro studies, using rat pancreas, showed that CCK caused a release of ACh that was blocked by tetrodotoxin or by inhibiting transmitter release by omitting Ca2+ from the bathing solution (Soudah et al. 1992). Moreover, the responses to low doses of CCK are less in patients after
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vagotomy than in normal subjects (Malagelada et al. 1974). Investigations in rat indicate that, at physiological concentrations, CCK initiates the reflex release of enzyme from the pancreas by stimulating the endings of vagal afferent neurons (Li & Owyang 1993, 1996). In these experiments, the effect of CCK at physiological levels was blocked by severing vagal afferent roots or by peri-vagal application of capsaicin, which destroys small-diameter afferent nerve fibers in the vagus. Nerve pathways connect the duodenum and pancreas directly, through the mesentery that join these organs (Kirchgessner & Gershon 1990). It is feasible that the direct pathways from the duodenum to the pancreas influence pancreatic secretion. Summary and conclusions Fluxes of fluid greater than the total blood volume of the body cross the epithelial surfaces of the gastrointestinal tract each day. Control of this fluid movement is of prime importance for the maintenance of whole-body fluid and electrolyte balance. The largest fluxes are across the epithelium of the small intestine, with significant fluid movement also occurring in the large intestine, stomach, pancreas, and gall bladder. Water flows between the lumens of digestive organs and body fluid compartments in response to movement of osmotically active molecules. The greatest absorption of water, 8–9 liters per day, accompanies inward flux of nutrient molecules and Na+, and the greatest secretion accompanies outward fluxes of Cl– and HCO3– in the small and large intestine, gall bladder, and pancreas. In each of these organs, fluid secretion is controlled by reflexes whose final motor neurons utilize VIP and ACh as neurotransmitters. In the small intestine and most of the colon the reflex circuits are intrinsic, in the enteric nervous system. They balance secretion with absorptive fluxes, and draw water from the absorbed fluid and from the circulation. The activity of the secretomotor reflexes is under a physiologically important control from inhibitory sympathetic nerve pathways that respond to changes in blood pressure and blood volume through central reflex centers. Fluid secretion is provoked by noxious stimuli, particularly by the intraluminal presence of certain viruses, bacteria and bacterial toxins. This secretion is due in large part to the stimulation of enteric secretomotor reflexes, although other mechanisms are also involved, such as the direct effects of toxins on the enterocytes, and release of cytokines. The physiological purpose is undoubtedly to rid the body of pathogens and their products. However, if the pathogens overwhelm the body’s ability to cope, the loss of fluid can become a serious threat to the organism. Secretion that is directly related to the chemical and enzymatic breakdown of ingested food, that is, HCl and pepsinogen secretion in the stomach, and secretion of pancreatic enzymes, is largely dependent on vago-vagal re-
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flexes. Enteric motor neurons are the final common pathway, but the roles of intrinsic reflexes are minor. Pancreatic secretion of bicarbonate, to neutralize the duodenal contents, is controlled by secretin, a hormone released from the duodenum, in synergy with activity of cholinergic and non-cholinergic neurons. Secretion into the gall bladder and bicarbonate secretion in the distal stomach are also nerve controlled.
7: Disorders of motility and secretion and therapeutic targets in the enteric nervous system
Changed patterns of motility and secretion in the gastrointestinal tract are a common occurrence, and range in severity from the inconvenience of a short-lived bout of diarrhea to chronic and debilitating diseases, referred to as enteric neuropathies. Because motility in the small and large intestine, and secretion of water and electrolytes, are under enteric nervous system control, it is logical that enteric neurons and the effectors that they control, especially smooth muscle and secretory epithelia, should be therapeutic targets. Current approaches have concentrated on receptors for neurotransmitters and hormones, and their degradative enzymes as targets. Targeting ion channels and second messenger systems that control the excitability of enteric neurons is discussed below. This has been considered before (Clerc et al. 2002), but has not been a focus of drug development. The possibility of using growth factors and neural stem cells to treat enteric neuropathies is also discussed. The enteric nervous system exhibits greater plasticity in the adult, particularly an ability to regenerate axons and restore synaptic connections, greater than is observed in the central nervous system (Giaroni et al. 1999). However, this plasticity is not being exploited therapeutically. Deviations from normal motility and secretion that involve the enteric nervous system include: congenital abnormalities of the enteric nervous system, such as Hirschsprung’s disease and hypertrophic pyloric stenosis; sporadic and acquired enteric disease, including enteric disorders associated with conditions affecting other body systems; and acute or chronic reactions of the gut to toxins, pathogens, or irritants (Table 7.1). A number of recent publications have discussed enteric nervous system disorders and their treatment (Camilleri 1990, Sharkey & Lomax 2001, Furness & Sanger 2002, Lundgren 2002, De Giorgio et al. 2004a,b, Spiller & Grundy 2004). It is not my intention to discuss the pathologies of these disorders, which have been well reviewed by others, except where it seems relevant to considerations of therapy.
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Table 7.1 Classification of enteric neuropathies
Congenital or developmental neuropathies Hirschsprung’s disease (colorectal aganglionosis) Intestinal aganglionosis (non-Hirschsprung’s) Hypertrophic pyloric stenosis Multiple endocrine neoplasia 2B Neuronal intestinal dysplasia (NID) Mitochondriopathies Sporadic and acquired disorders Intestinal pseudo-obstruction Slow-transit constipation Irritable bowel syndrome (IBS) Constipation-dominant IBS Diarrhea-dominant IBS Chronic constipation Chronic diarrhea Chronic diarrhea with substantial fluid loss Functional (non-ulcer) dyspepsia Auto-immune enteric neuritis Disorders secondary to other disease states Diabetic gastroparesis, diabetes-related motility disorders Enteric neuropathies associated with mental retardation or CNS disorders Iatrogenic or drug-induced disorders Laxative abuse colonic neuropathy Disorders initiated by anti-neoplastic drugs (e.g. vinca alkaloids, cisplatin) Postoperative ileus Opioid-induced dysfunction (e.g. when opioids used to treat chronic pain)
Therapeutic endpoints for motility disorders In the simplest of terms, disorders of intestinal motility include those in which progress of material along the intestine is too slow (ileus, pseudo-obstruction, constipation) and those in which it is too rapid (diarrhea). Irritable bowel syndrome (IBS) is an interesting disorder in that it can be constipation dominated (IBS-C) or diarrhea dominated (IBS-D), or may exhibit an alternation between the two conditions. Changes in motility are probably more subtle than often surmized when abnormalities of transit are considered, and may involve changed patterns of regulated motility, for example changes from activity that is predominantly propulsive to activity that is predominantly mixing (Chapter 5). Thus, a slow progress in the intestine could be due to inactivity of the enteric nervous system, such as appears to occur in ileus, or it might be a consequence of an inappropriate dominance of a mixing (nonpropulsive) pattern of motility or limited generation of propulsive activity. In general, the focus for drug treatments of various slow-transit disorders has been on compounds that have been rather crudely grouped as “prokinetics.” These include a variety of substituted benzamides, including one of the ear-
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liest introduced prokinetics, metoclopramide, and more recent derivatives (such as cisapride, prucalopride, and tegaserod), anticholinesterases, and muscle stimulants. Agonists at enteric 5-HT4 receptors The benzamide derivatives have a range of pharmacological actions, particularly on 5-HT receptors (Bockaert et al. 2004). Cisapride, for example, is a 5-HT3 receptor antagonist and 5-HT4 receptor partial agonist, whereas tegaserod, an aminoguanidine indole derivative, is a 5-HT2B receptor antagonist and 5-HT4 receptor partial agonist (Beattie et al. 2004). The effect at 5HT4 receptors has dominated theories of action of these compounds. When applied to the central compartment of a divided organ bath (see Chapter 4, Fig. 4.6), tegaserod caused polarized reflexes, contraction of the circular muscle on the oral side and relaxation on the anal side (Grider et al. 1998). The responses were prevented by a 5-HT4 receptor blocker. This is consistent with data that 5-HT4 receptors are located on the endings of IPANs in the mucosa (Chapter 4). Agonists of 5-HT4 receptors also increase the release of ACh by an action on 5-HT4 receptors of presynaptic nerve terminals of enteric neurons (Tonini et al. 2002). Thus, 5-HT4 agonists themselves initiate reflexes, or they lower the threshold at which physiological stimuli initiate enteric reflexes by acting on the endings of IPANs, and they increase the effectiveness of the reflexes by enhancing transmitter release (Gershon 2005). As might be expected of a compound that enhances peristaltic reflexes, tegaserod relieves constipation-dominant IBS (Prather et al. 2000, Müller-Lissner et al. 2001). Moreover, transit is slowed in 5-HT4 receptor knockout mice, which is consistent with 5-HT having a role, through 5-HT4 receptors, in initiating propulsive reflexes. Despite its usefulness, tegaserod shows only a small margin of therapeutic effectiveness above placebo (Crowell 2004). It is possible that other benzamide or aminoguanidine indole derivatives, or chemically unrelated 5-HT4 receptor agonists, will prove more effective. Would other excitants of IPANs be effective prokinetics? Receptor stimulants There have been few studies of other compounds that might stimulate the mucosal endings of IPANs, although this is a logical target given the positive effects of tegaserod. One stimulant is ATP, acting through P2X purine receptors (Bertrand & Bornstein 2002). ATP applied directly to the mucosal endings elicited action potentials that were recorded from cell bodies of IPANs in myenteric ganglia. The subunit composition of these P2X receptors has
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yet to be determined. It is feasible that agonists that are selective for the P2X receptors of the mucosal processes of IPANs will be useful prokinetics. The mucosal endings are exposed to many other possible stimulants, including products of entero-endocrine cells, such as serotonin, motilin, and cholecystokinin, and substances, such as proteases, prostaglandins, histamine, and a range of cytokines, released by cells of the mucosal defense system. Whether or not receptors for these occur on IPANs is worth exploring, as this might reveal therapeutically useful targets. Compounds targeting ion channels The numerous ion currents that control the excitability of IPANs have been studied in detail, and most are now identified (Chapter 2, Fig. 2.10). One of the most interesting from a control point of view is the current of the afterhyperpolarizing potential, IAHP . This current is active at rest, and is carried by intermediate conductance potassium (IK) channels. These channels are highly regulated in IPANs, and are closed by slow excitatory neurotransmitters, including tachykinins, and by various inflammatory mediators. Excitability of the neurons is enhanced by blockers of IK channels, such as clotrimazole or its derivatives, by agonists for receptors that are linked to IK closure, and by stimulants of the second messenger systems (PKA and PKC) that couple to IK channel closure (Chapter 2). Closure also changes the pattern of firing of action potentials, whose frequency is normally limited by the action potentialinitiated AHP (Chapter 2). It has been suggested that the AHP may assist in co-ordinating the firing of IPANs, and that, in the absence of the AHP, action potential firing is erratic (Thomas & Bornstein 2003). Thus, the change in action potential firing patterns caused by blocking the IAHP leads to an altered pattern of motility, in which co-ordinated propulsive reflexes become uncoordinated (Fig. 5.13). This tends to convert predominantly propulsive motility to predominantly mixing (or at least non-propulsive) motility. This would be an anti-diarrheal effect. Conversely, compounds that increase the opening probability of IK channels (a number of which have been found) could have an anti-constipatory effect by enhancing co-ordinated propulsive activity. Further in vivo experiments are required to assess whether agonists/antagonists at the IK channel may interfere with same or other K channels in both intestinal and extra-intestinal regions. IPANs are also excited by agonists at GABAA receptors, and this effect appears to be selective for IPANs (Cherubini & North 1984). Thus peripherally acting GABAA receptor stimulants would be predicted to have prokinetic actions. Further investigations of compounds that alter the excitability of IPANs, interneurons, and motor neurons may lead to useful therapies.
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Opioid receptors Morphine and related compounds, which are agonists at opioid receptors, have been used since ancient times for the treatment of diarrhea (Kreuger 1937). It is clear that the major sites of action of these drugs are on enteric neurons. Both opioid peptides and opioid receptors are present in the enteric nervous system. The peptides occur in motor neurons that supply the muscle, in interneurons, and in some secretomotor neurons (Chapter 2). Two opioid receptors, μ and κ receptors, are expressed on the endings of excitatory neurons that innervate the muscle (Tonini et al. 2002). The neurons contain the opioid peptides enkephalin and dynorphin, which act back on the endings to reduce transmitter release (Chapter 4). Thus when opioid agonists are applied ACh release from stimulated enteric neurons is reduced (Paton 1957, Schaumann 1957) and peristalsis is inhibited (Trendelenburg 1917). The release of tachykinins, which are co-transmitters with ACh (Chapter 2), is also reduced by opioid receptor agonists (Holzer 1984). Experimental evidence indicates that activation of opioid receptors reduces electrically evoked nonadrenergic, non-cholinergic inhibitory responses (Tonini et al. 1985) and the descending inhibitory reflex (Grider & Makhlouf 1987). In addition to an inhibition of propulsion along the intestine, opiate agonists reduce secretion of fluid into the lumen in the small and large intestine (Brown & Miller 1991). Antagonists of opioid receptors can increase release of ACh, indicating that endogenous opiates under some circumstances restrict transmitter release (Waterfield & Kosterlitz 1977). In human subjects, the antagonist naloxone, which enters the central nervous system, accelerates the movement of the contents (Kaufman et al. 1988), as does the peripherally restricted μreceptor antagonist alvimopan (Camilleri 2005). This points to the utility of opioid receptor antagonists as prokinetics, particularly in cases in which slow transit has been caused by opiates taken for pain relief. Peripherally restricted opioid receptor antagonist prokinetics could be used without compromising analgesic actions of opiates. Although a component of postoperative ileus may be caused by opiates given for pain relief (Bauer & Boeckxstaens 2004), there is possibly also a contribution by endogenous opiates that might be relieved by peripherally acting opioid prokinetics. Although opioid receptor agonists inhibit diarrhea, their profound motility-suppressing actions cause constipation, and the risk of this side effect restricts their use. In addition, their addictive properties make them generally unsuitable if they enter the central nervous system. An interesting approach is to treat diarrhea with an enkephalinase inhibitor, thus accentuating the effects of endogenous opioid peptides (Prado 2002).
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Therapies for secretory diarrheas Secretory diarrheas, that is diarrheas that involve substantial fluid loss, are consequences of enhanced transepithelial fluid movement, although changed patterns of motility may contribute (Jones & Blikslager 2002, Lundgren 2002). In many cases, they are a healthy reaction to a noxious presence in the lumen of the intestine, but they can lead to a deleterious outcome through excess fluid loss. The enteric nervous system controls fluid transport (Chapters 3, 6), and there is substantial evidence for the involvement of enteric neurons in the generation of some diarrheas, for example diarrhea triggered by cholera toxin, enterotoxins, and rotavirus (Chapter 6). Alongside their effects on enteric neurons, the toxins have direct secretomotor effects on the epithelium, and indirect effects through the stimulation of prostaglandin production (Field 2003). The possibility of targeting secretomotor reflex circuits for the treatment of secretory diarrheas has been reviewed recently (Lundgren 2002). Such treatment would almost always be in combination with oral rehydration therapy. The reflex circuits that are activated by noxious agents in the lumen were discussed earlier (Fig. 3.6). Activation begins when these agents trigger the release of 5-HT from enterochromaffin cells and prostaglandins from enterocytes. These act on the endings of IPANs that activate interneurons that in turn excite VIP-containing secretomotor neurons. Any part of this circuit might be potentially targeted, including 5-HT3 receptors on the endings of IPANs, neuro-neuronal synapses, and VPAC1 receptors on enterocytes. It might also be possible to use compounds that act on the enterocytes to prevent secretion, or on neurons to reduce their excitability or to reduce transmitter release. Drugs that antagonize 5-HT or VIP receptors have recently been tested (Kordasti et al. 2004). Rotavirus given as a single oral dose to mice caused diarrhea that lasted 3.3 days. The duration of diarrhea was reduced to 2.1 days by intraperitoneal injection of a VIP antagonist once each 10 hours for the 2 days (Kordasti et al. 2004). The antagonist was a peptide analog, and cannot be expected to be stable in vivo. However, the development of improved non-peptide antagonists of VPAC1 receptors might lead to effective anti-diarrheal compounds. The 5-HT3 receptor blocker granisetron, given once daily, reduced the duration of rotavirus-induced diarrhea by about 30% (Kordasti et al. 2004). The naturally occurring gut hormone PYY and the related neurotransmitter peptide NPY are effective anti-secretory agents with long-lasting effects (Saria & Beubler 1985, Cox et al. 1988, Brown et al. 1990). PYY reduces secretion (or enhances absorption) of water and electrolytes at physiological concentrations in vivo (Bilchik et al. 1993). It is possible that stable stimulants of PYY receptors could be effective in reducing hypersecretion. The
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human isolated mucosa is under tonic secretory suppression by endogenously released PYY or NPY, which is revealed by an increase in secretion that is caused by Y1 receptor agonists (Cox & Tough 2002). This indicates that PYY receptors, unlike many peptide receptors, are not desensitized by ongoing agonist exposure, and are therefore attractive therapeutic targets. Furthermore, the PYY receptor agonists are effective at nanomolar concentrations (Cox & Tough 2002). Enteric neuropathies involving neuronal loss or phenotypic changes The best-known gastrointestinal disorder with neuronal loss is Hirschsprung’s disease. Neurons fail to develop in the most distal parts of the bowel, and in variable proportions of the large intestine, and even in the small intestine. Hirschsprung’s disease is not a single entity, but can arise from a number of deficits in the expression of molecules that regulate the development of the enteric nervous system (Chapter 1). The only current “cure” is to remove the region in which enteric neurons have failed to develop. Other congenital disorders may involve more restricted neuron loss, even loss or phenotype change of particular subtypes of neuron. A peculiar neuropathological pattern with selected neuronal loss is observed in achalasia, a condition chararacterized by absent esophageal body peristalsis that is usually associated with failure of the lower sphincter to relax (De Giorgio et al. 2000). Achalasia is associated with loss of the inhibitory motor neurons, which is revealed by a deficiency of VIP or NOS immunoreactivity from fibers innervating the sphincter muscle (Aggestrup et al. 1983, Mearin et al. 1993, Wattchow & Costa 1996, De Giorgio et al. 1999). Although the symptoms of achalasia are prominent in the esophagus, the disorder can be associated with loss of enteric neurons in extra-esophageal segments, including in the small intestine. Loss of inhibitory innervation of the pyloric sphincter occurs in infantile hypertrophic pyloric stenosis (Wattchow et al. 1987, Vanderwinden et al. 1992). This may be a defect of enteric neuron maturation, because if the pylorus is slit to relieve the block the affected children remain free of the disorder after the cut pylorus has healed. Chronic intestinal pseudo-obstruction is also associated with enteric neuron degeneration (De Giorgio et al. 2000). In some patients with slow-transit constipation (STC), which is one of the most severe forms of idiopathic constipation, a substantial loss of the inhibitory innervation of the muscle, revealed by decreased numbers of VIP-immunoreactive nerve terminals, was reported (Koch et al. 1988), whereas in other patients the loss was in the enteric ganglia (Milner et al. 1990). However, no loss of VIP innervation was found in samples from adult female patients with STC, whereas there was a deficiency of tachykinin-immunoreactive fibers (Porter et
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al. 1998). A loss of tachykinin immunoreactivity from fibers innervating the muscle was confirmed in a study of children with STC (Stanton et al. 2003). In normal individuals, many of the excitatory fibers to the muscle contain tachykinins (TK) and ChAT, and some contain ChAT without TK in normal large intestine, whereas in colon from patients with STC, the numbers of TK-immunoreactive fibers innervating the muscle are reduced, the TK component of excitatory transmission to the muscle is diminished, but the cholinergic component of transmission is unaffected. Each of the examples just discussed involves a degenerative or developmental loss of enteric neurons, in some cases of specific neuron types, or a change in neuronal phenotype. These conditions may be treatable if therapies could be devised to reverse or replace the loss. One possibility is the use of growth factors that promote the proliferation and differentiation of enteric neurons, and/or guide the growth of enteric axons to their targets. Unlike neurons in the central nervous system, enteric neurons in adult animals re-grow and re-innervate their targets after they have been lesioned (Galligan et al. 1989a). Thus, signaling molecules for neuronal pathway guidance and target recognition in the enteric nervous system survive into adulthood. Many of the critical signaling molecules for maturation of enteric neurons are known (Gershon 2002, Newgreen & Young 2002a, Kapur et al. 2004, Young et al. 2004). Furthermore, the gut and peripheral nerves are both sources of neural crest-derived stem cells (Bixby et al. 2002, Joseph et al. 2004) that could probably be used to reconstitute a depleted enteric nervous system. Neural crest-derived stem cells harvested from rat intestine at embryonic day 14.5 can be sub-cloned in culture, and give rise to neurons and glia (Bixby et al. 2002), indicating their potential to be used in cell replacement therapy. Moreover, the enteric stem cells from rat gave rise to neurons and glia when transplanted into chick embryo hindlimb buds. Bone morphogenic protein 4 (BMP4), added in low concentration to cultures of gutderived neural crest-derived stem cells, promoted differentiation into neurons, even after the cells had been passaged through several generations. These observations suggest that there is a real possibility of using stem cells, combined with suitable growth and differentiation signaling molecules, to colonize the enteric system where neuronal deficiencies exist. Neuron replacement therapy will probably be enhanced when signaling molecules that direct the differentiation of specific types of enteric neuron are defined. Mitochondriopathies with intestinal manifestations Gastrointestinal disorders accompany a number of mitochondriopathies, the best documented being mitochondrial neurogastrointestinal encephalomyopathy (MNGIE) which is associated with food refusal, vomiting, diarrhea, and constipation in children in the first few weeks of life, and pseudo-obstruction
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at later ages, for example in teenage years (Mueller et al. 1999, Chitkara et al. 2003). MNGIE is due to a loss of function defect in thymidine phosphorylase, a nuclear DNA mutation, that leads to abnormal thymidine metabolism (Nishino et al. 1999). An insufficiency of thymidine to allow for mitochondrial DNA replication may explain the mitochondrial disorder. The multisystem mitochondriopathy MELAS, which is also associated with intestinal pseudo-obstruction, is caused by mutation of mitochondrial DNA (Chinnery et al. 2001). Gastrointestinal motility disorders in mitochondriopathies may have multiple causes, for example they could derive from deficiencies in neuron, interstitial cell of Cajal, or muscle function. However, as mitochondriopathies generally have neurological symptoms, a neuronal defect might be suspected. IPANs have greater mitochondrial numbers than do other enteric neurons, and the mitochondria have a critical role in sequestering Ca2+ from the cytoplasm in these neurons (Chapter 2). Thus disorders of oxidative phosphorylation may particularly target IPANs. IPANs express the KATP channel (Table 2.2), and when oxidative phosphorylation in these neurons is challenged they are hyperpolarized (Liu et al. 1999). This may contribute to pseudo-obstruction in mitochondriopathies with intestinal symptoms. It might be that these conditions could be improved with treatments that replenish energy stores of enteric neurons and drugs that enhance the excitability of IPANs. Irritable bowel syndrome and plasticity of enteric neurons Irritable bowel syndrome (IBS) is a chronic, fluctuating, and debilitating condition with an overall prevalence rate of about 10% in most industrialized countries that affects about 1% of the population at any one time, and for which there is no adequate therapy (Camilleri 2001, Crowell 2004). In addition to abdominal pain, which may occur as a result of the hyperactivity of the sensory nerve fibers supplying the gut, other predominant symptoms are due to altered bowel motility, which manifests as constipation, diarrhea, or fluctuations between the two (Kellow et al. 1988, Camilleri & Choi 1997, Drossman et al. 1999). It has large economic and social costs, including costs of health care and absenteeism (Talley et al. 1995). In the United States, it is estimated that the annual cost of IBS is $25 billion (Camilleri 2001). The incidence of IBS increases following inflammation such as bacterial enteritis and a high proportion of cases of IBS can be traced back to a period of intestinal inflammation (Gwee et al. 1996, Neal et al. 1997, Spiller 2003). Symptoms can persist for at least 6 years after an infective episode, even though the inflammation that precipitates IBS usually lasts only up to about 4 weeks (Neal et al. 1997, 2002). Moreover, 57% of patients with Crohn’s disease in remission report IBS-like symptoms (Simren et al. 2002). It has been estimated that an infective episode that lasts more than 3 weeks increases the
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probability of IBS developing by 11-fold (Spiller 2003). The implication is that the inflammatory process causes long-term changes in neuromuscular function, and specifically in the excitability of enteric neurons, that persist beyond the period of inflammation and result in the symptoms of IBS (Sharkey & Kroese 2001, Collins 2002, De Giorgio et al. 2004b, Spiller 2004). Evidence that supports a connection between inflammation and changed enteric neuron function is the invasion of the enteric nervous system by inflammatory cells in IBS, and the documented effects of inflammation on enteric neurons in animal models. Chadwick et al. (2002) examined mucosal biopsies from 77 IBS patients. In almost 90%, they found evidence of immune activation, including increased intra-epithelial lymphocytes, increased numbers of mucosal lymphocytes and increased mast cell numbers. The mean duration of symptoms before biopsies were taken was 2.5 years, so these patients were well beyond a triggering episode of acute infection or inflammation of the bowel. Evidence of increased mast cell numbers in the mucosa of IBS patients was confirmed in a study by Barbara et al. (2004). Electron microscopic evaluation showed a close spatial association between actively degranulating mast cells and nerve terminal supplying the mucosa of patients suffering from either constipation- or diarrhea-predominant IBS. Interestingly, degranulating mast cells in close vicinity (<5 μm) to nerve fibers were correlated with two clinical parameters, abdominal pain severity and frequency. More aggressive tissue sampling was undertaken by Törnblom (2002), who took full-thickness biopsies from the small intestines of 10 IBS patients. In each case, there were pathological changes in myenteric neurons, ganglia of 9 of 10 patients were infiltrated with CD3-positive T lymphocytes, but no lymphocytes were found in myenteric ganglia of non-IBS subjects. An intra-epithelial lymphocytosis also occurred in these patients. In six of the patients there was evidence of neuropathy, in which myenteric neurons were swollen, vacuoles were seen and some neuronal nuclei were pyknotic. The authors conclude that effects of inflammation on enteric neurons leads to the dysmotility that is characteristic of IBS. In animal models, post-inflammatory changes include invasion of myenteric ganglia by mast cells and lymphocytes (Rühl et al. 1995, Vallance et al. 1999, Sayani et al. 2004). In mice, as in human, transient inflammation results in persistent changes to neuromuscular function and in intestinal dysmotility (Barbara et al. 1997, Bercík et al. 2004). Also in mice, infection with T. spiralis is reported to reduce acetylcholine release from enteric neurons (Galeazzi et al. 2000). In addition, excitability of enteric neurons is increased following inflammation (Frieling et al. 1994a,b, Palmer et al. 1998, Linden et al. 2003, Liu et al. 2003). After nematode infection, neurons in the guinea-pig jejunum were depolarized relative to control, had increased input resistance and were more excitable (Palmer et al. 1998). Linden et al. (2003) found increased excitability of guinea-pig enteric neurons with post-action potential
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AHPs after TNBS (trinitrobenzene sulfonate)-induced inflammation of the colon. These neurons were assumed to be IPANs, but the inflammation could cause phenotypic changes that place this conclusion in some doubt. The properties of two particular ion currents (IAHP and Ih) in enteric neurons appear to be changed by inflammation, IAHP being depressed and Ih being enhanced. In this context, it is interesting that the expression of the channels of the IAHP (IK channels) is diminished in myenteric neurons of patients with inflammatory bowel disease (Arnold et al. 2003). These channels appear to be expressed in IPANs in human (Furness et al. 2004a), as they are in other species (Chapter 2). There is also immunohistochemical and immunoassay evidence of other changes in the neurochemistry of enteric neurons following inflammation, or in inflammatory bowel disease (Giaroni et al. 1999, Lomax et al. 2004). At present the drug therapies that have been applied to IBS have not been very effective, with the advantage over placebo only being in the range of 10–20% (Camilleri 2001, Crowell 2004). There has been little emphasis on the changes that occur in ion channel function and in neurochemistry in post-inflammatory IBS. It is possible that a greater understanding of the mechanisms of plastic changes in enteric neurons may reveal new and effective approaches to treatment. Summary and conclusions The identification of suitable target molecules in enteric neurons or the cells that they innervate and the search for therapeutically effective molecules that act on these targets to treat gastrointestinal disorders have produced limited outcomes. This has been partly due to the previous lack of detailed knowledge of structure and function of the enteric nervous system. However, understanding of the physiology and critical control points in the enteric nervous system has advanced rapidly in recent years, and continues to advance. In the past, the quest for targets and lead molecules has concentrated on neurotransmitters, hormones, and their receptors. However, much more is now known about ion channels and intracellular messengers that control excitability of enteric neurons, the effector cells that they innervate, and the entero-endocrine cells that activate enteric primary afferent neurons. The ion channels and intracellular messengers may provide new and viable targets. Moreover, although a number of enteric neuropathies involve degenerative and/or phenotypic changes in enteric neurons, targeting neuron replacement therapies or neuronal remodeling has been largely ignored. Impressive recent advances in the identification of molecules that are involved in regulating the differentiation, maturation, and guidance to targets of enteric neurons may provide ways to restitute neuropathic deficiencies that lead to disordered gastrointestinal function.
Epilogue: the future of enteric neurobiology
This book has focused on the juxtaposition of knowledge of the functional organization of the enteric nervous system and the physiological control of gastrointestinal motility and secretion. Some broad conclusions are reached. First, although the structure of the enteric nervous system appears similar from the upper esophagus to the terminal rectum, its degree of autonomy changes markedly. The contractile activities of the esophagus, especially its striated muscle part, and of the stomach, are largely controlled through centers in the medullary part of the brainstem that exert their effects via the vagus nerves. The vagus also has important roles in the direction of pancreatic and biliary functions. However, in the small and large intestines, the intrinsic control circuits operate with little vagal intervention. Nevertheless, the enteric control of these regions can be overridden by activity in sympathetic pathways. This is especially true of the control of fluid movement across the mucosa, in which enteric and sympathetic controls are closely integrated. The constituent neurons of the small intestine have been all identified, and the organization of the control circuits that they create is known in considerable detail. The physiology of the enteric nervous system is essentially the same in all mammals, although there are differences in details, such as the arrangement of ganglia and the chemical coding of enteric neurons. The chemical coding differences can lead to considerable confusion, and one of the important tasks in the future will be to determine the chemical identifiers of each of the functionally defined neurons in the human enteric nervous system. Some progress has already been made (Chapter 2), but a complete accounting for the enteric neuron types in humans that will be important for future investigations of enteric neuropathology has not been achieved. Despite the differences that are seen in the chemical coding of enteric neurons, the primary neurotransmitters of functionally equivalent neurons are conserved across mammalian species. Obtaining a more complete knowledge of the enteric nervous systems of the esophagus and stomach is an important challenge. It needs to be asked: what are the roles of the intrinsic connections between neurons in both the 211
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esophagus and stomach? Although intrinsic reflexes have been demonstrated in the stomach, it is not clear how important they are in governing its movements and in controlling acid secretion. Another theme of the book is to point out that the enteric nervous system has roles in controlling the gastrointestinal system under favorable circumstances and also when the gastrointestinal tract is under threat from noxious agents. The defensive role of the enteric nervous system is often overlooked, but it is actually unsurprising that the enteric nervous system plays a major role in defending the digestive tract and hence the whole organism. After all, the gastrointestinal tract presents the largest permeable surface of any organ system to the outside world, in humans about 300 square meters of vulnerable luminal surface. This defensive role matches the occurrence within the intestines of 70–80% of the lymphocytes in the body. I have chosen not to present a detailed analysis of the interactions between the enteric nervous system and the gut immune system in this book. It is nevertheless an important topic. Although the information available is already extensive, in my view a comprehensive understanding of gastrointestinal neuro-immune interactions has not yet been achieved. The secretomotor reflexes, especially in the small intestine, have the important role of returning water to the lumen. A large amount of water is absorbed with nutrients, and around 9 liters per day is returned to the lumen through active secretion by the enterocytes; this return is controlled through enteric secretomotor reflexes and sympathetic neurons. Dysregulation of secretomotor control, for example in cholera intoxication, is life threatening. Transepithelial fluid fluxes are too great to be ignored by the rest of the body. This is why they are regulated by the central nervous system through the sympathetic innervation of the gut. This sympathetic regulation plays one part in the homeostatic regulation of body water, which is linked to the control of blood volume, blood delivery to the tissues, fluid intake, and fluid excretion. Many abnormalities of gastrointestinal motility occur, and in some cases these can be traced to malfunction within the enteric nervous system. This emphasizes the need to understand the mechanisms through which enteric neurons contribute to enteric neuropathology, and to use the recently acquired, extensive basic knowledge of the enteric nervous system to provide solutions to motility disorders. Part of the answer will be to understand more adequately how transitions between motility patterns occur. The small intestine exhibits several stereotyped patterns of movement: mixing activity, propulsive movements during normal digestion, the migrating myoelectric complex, rapid propulsive movement (the peristaltic rush) when there are noxious materials to be propelled, and retropulsive movement as a component of vomiting. All of these depend on the enteric nervous system, which
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must thus be programmed to exhibit each of these patterns. The challenge remains to uncover how the enteric nervous system generates the different motility patterns, and what signals are critical to switching between one activity and another. Solutions to these puzzles may point to ways to control the patterns artificially, and thus modify intestinal behavior to suit patients with motility disorders. Another challenge that lies ahead is to determine which nerve growth, guidance, and differentiation factors influence the maturation of each of the neuron types, about 15 in the small intestine, that constitute the enteric nervous system. If these mechanisms were properly understood, it might be possible to treat enteric neuropathologies in which one or more neuron type is defective in its function. In summary, now that there is a substantial, even though not absolutely complete, knowledge of the functional organization of the enteric nervous system, the stage is set to put that knowledge to advantage in improving the treatment of patients with gastrointestinal disorders.
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Index
Note: Page numbers in italics refer to illustrations; those in bold refer to tables. accommodation reflex 97, 141 acetylcholine inhibitor of excitatory transmitter release 108 interneurons 126 major neurotransmitter 32, 33, 54, 103–4, 113 release 93, 108–9 stimulates gastric secretion 189–91 acetylcholine esterase (AChE), inhibition 104 achalasia 206 action potentials 43, 44, 47–8, 139, 140 adenylyl cyclase (AC) 115–16 after-depolarizing potential 48–9 after-hyperpolarizing potential (AHP) 44, 45, 48–52, 154, 203 absence 47, 70 aganglionosis 25, 201 AH neurons absent or very rare in stomach 146–7 Dogiel type II morphology 64 electrophysiological properties 45–52 terminology 43–4 amino acid transport 181 anal sphincter 162, 164, 169 anti-neoplastic drugs 201 anticholinesterases 202 antidromic and orthodromic activation 112 apamin 110 aquaporins 181 area postrema 157 arterioles, innervation 90, 92, 99 ascending excitatory reflexes 121–2, 125 ATP inhibitory action 111, 124 in motor neurons 103 as neurotransmitter 54, 74, 107–9, 114 receptor stimulant 202–3 Auerbach’s plexus see myenteric plexus auto-immune enteric neuritis 201
benzamide derivatives 201–2 bicarbonate secretion 182–3, 185, 194, 195, 196–7 bile salts 195–6 biliary system 16, 25–6 see also gall bladder BK channels 48, 50 blood volume, control 184, 212 bombesin 32, 33, 130 mammalian see gastrin-releasing peptide (GRP) brainstem reflex centers 145–7, 159 Brunner’s glands, innervation 14 c-kit 134, 136 calbindin 33, 86 calcineurin (protein phosphatase 2B) 50, 51 calcitonin gene-related peptide (CGRP) 27, 32–3, 93, 103, 122, 123, 188 calcium removal by mitochondria 49, 50 role in generation of late AHP 49 calcium channels blockers 49 high voltage activated (HVA) 47 calretinin 32, 33, 59–60, 64, 86 capsaicin 188 carbon monoxide 111 carnivores 147, 148, 151 central nervous system afferent signals 69 bypassing 69 control of stomach reflexes 97, 98 Chagas’ disease 157–8 Champy zinc iodide-osmium method 136 charybdotoxin 50 chemical coding 32–3, 103–4, 211 chemical stimulation see mechanical or chemical stimulation cholecystokinin (CCK) 32, 33, 103, 115, 160–1, 165, 196–8 267
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choledeco-duodenal sphincter see sphincter of Oddi cholera toxin 90, 91, 180 choline acetyltransferase (ChaT) 27, 32–3, 77, 87, 103, 106 choline acetyltransferase (ChaT)/5-HT neurons 123–4 choline acetyltransferase (ChaT)/NOS neurons 88, 124 choline acetyltransferase (ChaT)/SOM neurons 123–4 cholinergic nerves, inhibitory action on gastrin release 193 cholinergic transmission 87 circular muscle contraction 81, 96, 155–6 innervation 6, 10, 11, 12, 14, 95 motor neurons 54–6, 57, 85, 88 motor unit 95–6 movements in vivo 86 cisapride 202 clotrimazole 203 cocaine and amphetamine-regulated transcript peptide (CART) 31, 32, 33 coeliac plexus 176 colon extrinsic reflex 89 interneurons 63, 87 mixing 158 motility 157–9 slow wave activity 158 types of neurons 78 computer modeling 94 constipation chronic 201 slow-transit (STC) 201, 206 contractile events, speeds of conduction 143 corticotrophin-releasing factor (CRF) 31, 32, 33 Crohn’s disease 208 Cryptosporidium 90 crypts of Lieberkuhn 61 cystic fibrosis 180 cystic fibrosis transmembrane conductance regulator (CFTR) 181, 182 deep muscular plexus 11–12, 95–6 defecation 87, 158, 164 dendrites 29, 31, 40 depolarization, transient 49 descending excitatory/inhibitory reflexes 122–6 diabetes-related motility disorders 201 diarrhea 74, 186, 204, 205 chronic 201 life-threatening 182, 186, 205 secretory 186, 205–6 toxin-mediated 74, 186 digestive system, monitoring and control by neurons 69
digestive tract see intestine distension 73, 83–4, 88, 92, 95 Dogiel type I neurons 31, 32, 33, 34–5, 55, 77 small 40–1 Dogiel type II neurons 35–40 absent or rare in stomach 52, 147 action potentials 44 AH neurons 47 dendritic type 40 description 31, 32, 33 in humans 78 IK channels 50 rich in mitochondria 39, 52 ultrastructure 19, 39 Dogiel type III neurons 31, 32, 40–1, 63 duodenum 63, 185 dynorphins 32, 33, 103 dyspepsia, functional (non-ulcer) 201 electrolytes 59, 180–9 emesis see vomiting endocrine cells 13, 62 see also entero-endocrine cells endothelins and their receptors 24–5 enkephalin 27, 32–3, 108 enteric disease, sporadic and acquired 200 enteric ganglia development 23 distribution 7 drug effects 19 structure 1, 17–20 enteric glial cells see glial cells enteric interneurons 63–4 enteric nervous system age-related changes 27 defensive role 212 development 23–7 evolutionary aspects 21–3, 211 genetic basis of disorders 24 historical evolution of ideas 80 independent action 80 interaction with CNS 1 invertebrate 21, 22 plasticity 200, 207, 208–10 reflex circuitry 80–102 structure 1–28 types of neurons 32–3 vertebrate 81 enteric neurons chemical coding 31, 32–3 classification 29–30, 40, 43–53 conduction speeds 44 electrophysiological properties 43 functionally defined 53–76 human 76–8 neurochemical signature 79 neurotransmitters 103 pre-vertebral synaptic connections 174 precursors 25 shapes 31–43
IND EX
as therapeutic targets 200 enteric neuropathies 200, 201 enteric neurotransmitters and their receptors 32–3 enteric plexuses, structure 2, 3–14 entero-endocrine cells 62, 66–7, 126–7 entero-enteric reflexes 69, 171–6 entero-gastric reflex 176, 178 enterochromaffin cells 90, 126 enterochromaffin-like (ECL) cells 189, 190 esophagus control of motility 96–7 innervation of glands 14 myenteric plexus 96–7 neural control 159–60 neurons 52–3, 78 paucity of ganglia in submucosa 9 peristalsis and swallowing 159–60 striated muscle, innervation 58, 97 excitation, propagation 81, 88 excitatory junction potentials 108 excitatory motor neurons circular muscle 55 co-transmitters 106–8 density 94 neurotransmitters 54, 104–6 excitatory post-synaptic potentials (EPSPs) 44 fast 111, 112–14, 161, 174–5 slow 111, 115–16 excitatory transmission, non-nicotinic 93 exocrine secretion 58–9 fed and unfed states 147, 148, 151 feedback inhibition 105 fluid balance 58–9, 101, 188–9 fluid secretion see diarrhea forskolin 115 GABAA receptors 51–2, 203 galanin 32–3, 103 gall bladder contraction 160–1 intrinsic reflex pathways 161 neurons 25–6, 53, 98–9 secretion into 195–6 see also biliary system gamma-amino butyric acid (GABA) 32, 33, 108 ganglia, in the mucosa 9 ganglionated plexuses 1, 2, 3–9, 15–17 gastric acid secretion 62, 189–90, 192–3, 194 gastric corpus, S and AH neurons 52 gastric inhibition, reflex 176 gastric mixing 142–5 gastric motility (peristalsis) 97–8, 140–7, 143 gastric mucosal hyperemia 187–8 gastric muscle preparations 97–8 gastric protection 187–8
269
gastric reflexes, intrinsic 193–4 gastric reservoir function 140–2 gastric secretomotor neurons 62 gastric vasodilator neurons 60–1 gastric volume, adjustment 97 gastric, see also stomach gastrin 62, 130, 189–91 gastrin-releasing peptide (GRP) 32–3, 115, 130, 162, 191, 195 gastroenteropancreatic endocrine system 67 gastrointestinal muscle, rhythmic activity 132–4 gastrointestinal tract see intestine gastrointestinal veins, innervation 61–2 genetic studies 25 giant migrating complex see peristaltic rush glands of Lieberkuhn 13 glial cells 17, 18, 20–1 glial-derived neurotrophic factor (GDNF) 24 gliofilaments 21 glucose 88, 181, 182, 184 glutamate 114 granisetron 88, 90, 125, 205 guinea-pig classification of neurons 29, 30, 31, 32–3 model species 53, 54, 86 gut see intestine haustral contractions 158 HCN2 channels 51 heat-stable enterotoxin 90 herbivores 147 hexamethonium 90–1, 92 Hirschsprung’s disease (colorectal aganglionosis) 24, 25, 157–8, 164, 200, 201, 206 histamine 115, 189–91 5-hydroxytryptamine (5-HT) as co-transmitter 114 in enteric reflexes 74, 126–7 in entero-endocrine cells 62, 126 nerve-mediated release 62 as neurotransmitter 27, 32, 33 in transduction of mucosal distortion 92 5-hydroxytryptamine (5-HT) receptor agonists 202 blocker 89 hyperpolarization-activated cation current 51 iatrogenic or drug-induced disorders 201 IK (intermediate conductance potassium) channels 48, 50–1, 50, 154, 155, 203 ileo-cecal junction 163–4 ileo-cecal sphincter 161–2 ileo-colic sphincter 169 ileum, neurons 63, 64 ileus 177–8, 201, 204 immune system cells 13, 67–8
270
I NDE X
infective agents, intraluminal application 90 inflammation 209–10 inhibitory motor neurons circular muscle 55 continuously active 154 morphology 55 neurotransmitters 54, 105, 109–11 sympathetic inputs 88 inhibitory post-synaptic potentials (IPSPs), slow 111, 118 inhibitory transmitters, relative roles 54 interganglionic strands (connectives) 4 interneurons ascending 86–7 assemblies 95 descending 87–8, 123–4 electrophysiological studies 87 enteric 63–4 motility reflexes 86–8 in myenteric plexus 85 pharmacological studies 87, 88 properties 78 synaptically connected chains 85, 86 transmission between 123 VIP 64 interstitial cells of Cajal (ICC) electrical activity 137 in generation of slow waves 97, 134 innervation 104, 105 in neuromuscular transmission 12, 127–8 NK1 receptors 107 structure and properties 134–7 subgroups 136 intestinal crypts (glands of Lieberkuhn) 13 intestine acute or chronic reactions 200 afferent neurons 68 enteric ganglia 7 extrinsic denervation 82 fetal development 26 inflammation 208–10 irritation or handling 176, 177, 178 motility 81 abnormalities 212 mucosal innervation 13–14 slow waves 138–9 sphincters 161–5 stationary contractions 151–2 sympathetic innervation 99–101 see also large intestine; small intestine; and individual organs intestino-intestinal inhibitory reflex, spinal 177 intestinofugal afferent neurons (IFANs) 76, 79, 174–5 intrinsic primary afferent neurons (IPANs) absent or rare in stomach 146 AH electrophysiological characteristics 38, 64, 70
anally directed processes 95 c-Fos induction 74 calbindin immunoreactive 88 characteristics 33, 69–70 chemical coding 33 chemical markers 31 connections 83 currents and channels 46 detection of activity 66 Dogiel type II morphology 38, 64, 69 efferent effects 66 EPSPs 114, 115 excitants 202–3 extrinsic and intrinsic 68–9 group activation 94–5 human equivalents 76 immunoreactivity 93 innervate submucosal neurons 92 intermediates in activation 74 as interneurons 66 length of projections 122–3 mechanosensitivity 70–3 neurotransmitters 33, 94 as nociceptors 66, 73–4 and peristaltic reflex 154 polymodal nature 74–5 response to stimuli 66, 83–4, 203 rich in mitochondria 208 submucosal 94 sustained slow post-synaptic excitation 116–17 synaptic connections 75–6, 94, 122, 123, 130 terminology 29, 65–6 transmission from entero-endocrine cells 126–7 intrinsic sensory neurons see intrinsic primary afferent neurons (IPANs) invertebrates, enteric nervous system 21, 22 ion channels, drug targeting 203 irritable bowel syndrome (IBS) 201, 208–10 irritants, secretory responses 186–7 isolectin B4 (IB4) 31, 32–3, 74 KATP channels, in AH neurons 52 lamina propria, nerve fibers 13 large intestine motility control 81–8 water and electrolyte secretion 180–9 laxative abuse colonic neuropathy 201 longitudinal muscle contraction 155–6 innervation 10, 56, 58 movements in vivo 86 lower esophageal sphincter 161, 162–3 lungs, ganglia 16–17 lymphatic system 13, 62–3
IND EX
mammalian bombesin see gastrin-releasing peptide (GRP) Mash-1 24 mast cells innervation 63 mecamylamine 93 mechanical or chemical stimulation 83, 88, 92–3, 187 Meissner’s plexus see submucosal plexus MELAS 208 metoclopramide 202 migrating myoelectric complexes (MMCs) in carnivores 147, 148, 151 characteristics 32 dependent on enteric nervous system 124, 149–50, 212 fetal 26 function 151 generation 95 initiation 150 in intestinal motility 147–51 migration 63, 82, 87, 124 persistence 149, 170 phases 147–8, 150, 151 in ruminants and other herbivores 147 speed of propagation 143, 150 mini-neurons 40–1 mitochondrial neurogastrointestinal encephalomyopathy (MNGIE) 207–8 mitochondriopathies 201, 207–8 mixing activities 142–5, 158, 212 motilin 62, 150 motility control 81–8 disorders 201–4 inhibition in non-sphincter regions 166–9 physiological effects of noradrenergic neurons 170–1 reflexes 80, 84, 86–8 sympathetic inhibition 166–9, 171–8 motor neurons assemblies 95–6 characteristics 53–63, 78 muscle 53–4 projections 55–6 transmitters 104–11, 130 mucosa innervation 13–14 mechanical or chemical stimulation 83, 88, 92–3, 187 mechanoreceptors 73 motor neurons 59, 60 muscles 165–6 mucus-secreting glands, innervation 14 multiple endocrine neoplasia 2B 201 muscarinic antagonists 60, 104 muscarinic receptors 92, 105, 106 muscle motor neurons 53–4, 56, 57
271
noradrenergic innervation 99 stimulants 202 muscularis mucosae contractions 165–6 innervation 12–13 motor neurons 56–7 myenteric ganglia see myenteric plexus myenteric neurons 31, 32 IPANs 69 in the esophagus 97 slow EPSPs 115 slow IPSPs 118–19 ultrastructure 19 myenteric plexus components 4–6, 10, 58 extent 15 fetal development 26 gastric 97–8 innervation of circular muscle 54 interconnections 14–15 interneurons 63 motor neurons 56, 57 number of nerve cells 15, 93–4 esophageal 96–7, 159 severed 85, 86, 92 species differences 3–4, 5 structure 2, 3–6, 39 tertiary component 6, 10, 58 vasodilation control pathways 92 NADPH reductase, fetal activity 26–7 nausea 69, 157 netrins 26 neural crest cells 23, 207 neural markers, fetal 27 neuro-neuronal synapses post-synaptic events 111 presynaptic inhibition 119–20 neurofilament protein (NFP) 32–3 neurokinin (NK) 32, 33, 106, 107 neurokinin (NK) receptors 32, 105, 107, 115 neuromuscular transmission 127–8 neuronal intestinal dysplasia (NID) 201 neuronal nuclear protein (NeuN) 31, 32, 33 neuronal tracing experiments 98 neurons assemblies 93–6 change of phenotype 206–7 degenerative section 82 density in the gut 93–4 displacement 64 functional classes 78 giant 43 in human intestine 76–8 inexcitable 43 loss 206–207 see also aganglionosis; Hirschsprung’s disease mechanosensitivity 72–73
272
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shape and function 29 see also Dogiel type neurons neuropathies, congenital or developmental 201 neuropeptide γ 106 neuropeptide K 106 neuropeptide Y (NPY) 27, 32, 33, 103, 205–6 neurotransmitters at neuro-neuronal synapses 111–20 and co-transmitters 32, 33 conserved across mammalian species 103 multiple 103 nicotinic receptors 113 nicotinic synapse 91 nitric oxide (NO) as co-transmitter 32, 109 immunoreactivity 77 inhibitory transmitter 54, 103, 110–11 mediates relaxation of the gastric fundus 142 retrograde transmitter 125 nitric oxide (NO) receptor 111 nitric oxide synthase (NOS) activation 110–11 conserved 103 fetal 27 in inhibitory motor neurons 55 in interneurons 32, 86, 88 non-adrenergic, non-cholinergic (NANC) transmission 109 noradrenaline 99, 101 noradrenergic neurons 100–1, 170–1 opiates, used in treatment of diarrhea 204 opioid antagonists 108 opioid peptides and receptors 108, 204 see also enkephalin opioid receptor antagonists, as prokinetics 204 opioid-induced dysfunction 201 oral ganglia 15 oral rehydration therapy 181, 187, 205 orexin 31, 32, 33 orthologues 29 pain, sensation 69 pancreas ganglia 16 neurons 25–6, 53, 98–9 secretions 196–8 paralytic secretion 183 paravascular nerve fibers 14 paravertebral ganglia 100 partitioned organ baths 120–1 pelvic nerve 17 penetrating fiber bundles (vertical fibers) 15 pepsin and pepsinogen 194–5 peptide histidine isoleucine (PHI) 110 peptide histidine methionine (PHM) 110
peptide YY 62, 205–6 peptides opioid 108, 204 regulatory 31 periglandular plexus 13 peripheral inhibitory reflexes 185 peristalsis colon 158 during normal digestion 212 gastric 97–8, 140–7 in vivo studies 152–4, 170 intestinal 81 esophageal 97, 159–60 speed of conduction 143 see also propulsive reflexes peristaltic reflexes 49, 113, 126–7, 158 peristaltic rush 143, 156, 212 peritonitis 176, 178 perivascular plexus 14 Peyer’s patches 13, 62–3 pharyngeal ganglia 15 phospholipase C (PLC)–diacyl glycerol (DAG)–PKC pathway 115–16 physostigmine 112 pituitary adenylyl cyclase activating peptide (PACAP) 32, 33, 106, 110, 115 plexus sous-musculeux see submuscular plexus plexus of the villous core 13 plexuses, interconnections 14–15 postoperative ileus see ileus presynaptic inhibition 119–20 prevertebral ganglia 100 primary afferent neurons 65, 66–7 primary transmitters, modulation of release 108–9 prokinetics 201–2 propulsive reflexes 74, 81, 82, 84–5 protein kinases A and C 50–1 protein phosphatase 2B (calcineurin) 50, 51 prucalopride 202 pseudo-obstruction 201, 206 pseudounipolar neurons 35 purine receptors 105, 110, 202–3 purinergic transmission 93 pyloric sphincter 161, 163 pyloric stenosis 200, 201, 206 receptive relaxation reflex 97 receptor stimulants 202–3 receptor-tyrosine kinase (Ret) 24 recording conditions, influence on electrophysiological properties 52 recto-anal reflex 158–9, 164, 165 reflex control of water and electrolyte secretion 182–9 reflex pathways, monosynaptic 87 reflexes co-ordination 98
IND EX
intrinsic 88–93, 212 polarized 81, 83 retropulsive movement 212 rotavirus 90, 91 ruminants MMC 147 S neurons 43–5, 112, 118 S-100 calcium binding protein 20 Salmonella 90 satiety 69 Schwann cells 20 second messenger systems 203 secondary plexus 6 secretin 194, 196–7 secretion 94, 187–9 secretomotor neurons cholinergic 59–60, 61, 62, 90 density 94 innervated by IPAN 92 intrinsic restraint by PYY 185–6 non-cholinergic 90 projections 60 role in secretion 58–60 small intestine 61, 90 synapses 130 secretomotor reflexes benign stimuli 182–3 drug target 205 early ideas 80 important role 212 initiated by 5-HT 126–7 initiated by pathogens or toxins 90–1 intrinsic 88–93 local nature 89–90 noxious stimuli 182, 186–7 sympathetic regulation 183–5 secretomotor transmission 128–9 secretomotor/vasodilator control 188–9 secretomotor/vasodilator neurons 58–9, 60, 61 SGLT1 181 sham feeding 192, 197 SK channels 50 slow synaptic transmission, S neurons 118 slow waves amplitude 139, 140 colon 158 depolarizing potentials 132–3 early studies 132–4 generate gastric peristalsis 142, 147 ICC generated 137 intestine 138–9 and neural control 138 properties 139 speed of conduction 143, 153 small intestine ascending interneurons 86–7, 95 contractile events 143
273
density of enteric neurons 93–4 descending interneurons 87–8, 95 emesis and retropulsion 157 innervation 11 motility 81–8, 147–57 secretomotor neurons 90 transit times 155 types of neurons 78 water and electrolyte secretion 180–9 small simple neurons 40–1 smooth muscle, innervation 53–4, 104–11 somatostatin (SOM) 32, 33, 103, 119, 124, 193 sphincter of Oddi (choledeco-duodenal sphincter) 39, 98–9, 162, 164–5 sphincters 99, 161–5, 169, 171 splanchnic nerves 172, 176, 177 stem cell therapy 207 stomach contractile events 143 distention 146, 147 functions 97, 140 intrinsic reflexes 212 motility 97 myenteric plexus 97–8 paucity of Dogiel type II neurons 52 paucity of ganglia in submucosa 9 receptive relaxation 141 regional differences 140 slow waves 138, 140 types of neurons 78 vagus-mediated reflexes 97, 98 see also entries under gastric stretch-activated channels (SACs) 71 subglandular plexus 13 submucosal plexus control of motility and secretion 92 development 25–6 Dogiel type II neurons 39 early studies 2, 3 extent and density 15 inner and outer 8–9, 54 innervation of circular muscle 10, 54 innervation of longitudinal muscle 56 innervation of mucosa 60 interconnections 14–15 motor neurons 56, 84 NOS-immunoreactive neurons 64 secretomotor neurons 90 severed 85 slow IPSPs 118–19 species differences 8, 9 structure 6–9 types of neurons 33 ultrastructure of neurons 19 submuscular plexus 11–12 subserosal plexus 10, 14–15 substance P 106, 107
274
I NDE X
suramin 110 sustained slow post-synaptic excitation (SSPE) 116–17 swallowing, esophageal peristalsis 159–60 sympathetic inhibition 101, 176–8 sympathetic neurons ganglionic 100 post-ganglionic 99 sympathetic reflexes 176 sympathetic regulation 212 synaptic inputs, in myenteric and submucosal plexuses 19 synaptic vesicles 19–20 tachykinins (TK) cause secretion 93 as co-transmitters 107 contribute to gastric protection 188 defective response 206–7 excitatory transmitters 54, 103, 106–7 in inhibitory reflexes 124 as transmitters 32, 33, 124 tachykinin (TK) receptors 107, 115, 124 tachykinin (TK)-immunoreactive neurons, in humans 77 tegaserod 202 tertiary plexus 6, 10, 58 tetrodotoxin (TTX) 45, 46, 47, 88 tissue fluid volume, conservation 184 tissue protection 187–8 toxins expulsion 156, 157 intraluminal application 90, 91 secretory responses 186–7 tracer studies 19 trachea and bronchi, ganglia 16–17 transcription factors 25 transit abnormalities 201 times 155 transmitters see neurotransmitters Trendelenburg method 81–2, 83 Type IV neurons 33, 41–2 Type V neurons 42 Type VI neurons 42–3 Type VII neurons 43
ultrastructural studies, quantitative 55 undigested material, removal by MMC 151 Ussing chamber 59, 89 vagal motor neurons 58 vagotomy 145, 146, 172, 176, 193 vagus nerve control of esophagus 159, 160 innervation of Brunner’s glands 14 innervation of enteric neurons 97 intramural extensions 17 reflexes mediated by 97, 98, 145–7, 192 and reverse peristalsis of vomiting 157 stimulates release of gastrin 190–1 stimulation 60, 62 vascular innervation 14 vasoactive intestinal peptide (VIP) antagonist 205 as co-transmitter 109, 110 fetal 27 in gall bladder 195–6 as inibitory transmitter 54, 103, 111 mimics slow EPSPs 116 neurons 31, 59–60, 77, 101 as non-cholinergic transmitter 61, 182 secretomotor and vasodilator transmitter 128–9 stimulant of secretion 59 as transmitter 32, 33 vasodilation 59, 92 vasodilator neurons 33, 60, 61, 90 vasodilator transmission 129, 130 vasomotor circuits, intrinsic 88–93 vertebrates, enteric nervous system 81 vesicular acetylcholine transporter (VAChT) 93 villi smooth muscle, contractions 165–6 structure 13 subepithelial plexus 13 vomiting 157, 212 VPAC receptors 105, 110 water secretion 180–9 transport 59
