The Role of Submucosal Neurons in Physiological and Pathophysiological Intestinal Secretion
Candice Fung
BBiomedSc, MSc, UMelb.
ORCID iD: 0000-0002-4277-3664
Submitted in total fulfillment of the requirements of the degree of Doctor of Philosophy
October 2016
Department of Physiology
The University of Melbourne
Produced on archival quality paper
ABSTRACT
The enteric nervous system (ENS) has two major types of secretomotor neurons, which contain either acetylcholine (i.e. are cholinergic) or vasoactive intestinal peptide (VIP). These are well conserved across species, and are key mediators of physiological and pathophysiological intestinal secretion. Notably, VIP is a potent secretogogue implicated in cholera toxin (CT)-evoked hypersecretion – the most widely-studied model of bacterial toxin-induced diarrhoea. As diarrhoeal disease remains a significant health problem worldwide, a better understanding of these secretomotor pathways and mediators in physiological and pathophysiological secretion is crucial for advancing therapeutic treatments. While acetylcholine is an established enteric neurotransmitter, VIP has long been considered a putative transmitter. VIP acts via VPAC1 and VPAC2 receptors, but their physiological role within the enteric circuitry remains unclear. Despite previous efforts to better characterize the role of VIP, a number of discrepancies arose from these studies. These issues were mainly due to the lack of selective pharmacological tools, as well as technical limitations. Furthermore, although the secretory effects of CT have been studied extensively, the degree of enteric neural involvement, and the relative contribution of cholinergic and VIP secretomotor neurons remains debatable. Thus, the aim of my PhD studies was to address these longstanding questions which have impeded our understanding of the control of intestinal secretion by employing specific agonists and antagonists for VPAC receptors and more advanced experimental techniques that are now available.
In Chapter 3, I localized VPAC1 receptors to a subset of cholinergic secretomotor neurons and cholinergic excitatory longitudinal muscle motor neurons. Further, I demonstrated that VIP acts via VPAC1 receptors on cholinergic secretomotor neurons to stimulate secretion and longitudinal muscle contraction. This was done using a combination of molecular, immunohistochemical and functional studies, as well as selective antagonists. There was no evidence of VPAC2 receptor involvement.
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In Chapter 4, I identified a novel role of VIP and VPAC receptors in modulating neuro-glial communication in the mouse submucosal plexus. This was achieved by performing Ca2+ imaging on transgenic Wnt1-Cre;R26R-GCaMP3 mice, which express a fluorescent Ca2+ indicator in their ENS (enteric neurons and glia). VIP application to submucosal ganglia exclusively evoked Ca2+ responses in neurons, but surprisingly, using specific agonists and antagonists for VPAC1 and 2 revealed a role for VIP as a transmitter in signaling from neurons to glia. Activating VPAC1 receptors initiates neuron-to-glia signaling by stimulating purine release. Whereas, activating VPAC2 receptors suppresses this signaling pathway. Thus, VIP may have a dual role through its activation of VPAC1 and 2 receptors in fine-tuning purinergic neuron-glia communication.
In Chapter 5, I used an in vivo mouse ileal loop model of CT-incubation, followed by a series of in vitro analysis to investigate the relative contributions of cholinergic and VIP secretomotor neurons to CT-evoked hypersecretion. Following the incubation, I found an increased fluid accumulation in CT-treated loops which corresponded to an increase in basal secretion measured using Ussing chambers. This hypersecretory effect did not appear to have a significant neuronal component. However, I found that CT induced a sustained increase in the excitability of cholinergic submucosal neurons by utilizing Wnt1-Cre;R26R- GCaMP3 mice to examine Ca2+ activity within the enteric circuitry. Increased spontaneous activity and enhanced responses to neural stimulation was observed in cholinergic submucosal neurons following CT-exposure. Further, CT was found to activate most submucosal neurons with the increased expression of an activity-dependent marker pCREB. I also found that the myenteric plexus did not significantly contribute to the sustained hypersecretion. Collectively, I showed that CT induced sustained hypersecretion by a direct effect on the mucosa, but also partly by increasing the excitability of cholinergic submucosal neurons.
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Overall, my thesis presents the first evidence for neural VPAC1 receptor expression in the ENS and demonstrates their function in the regulation of intestinal secretion and motility. Furthermore, I identified a novel modulatory role of VIP and VPAC receptors in signaling within the enteric neuro-glial circuitry. Finally, my work also extended our knowledge of CT-evoked hypersecretion, highlighting the significant contribution of cholinergic submucosal neurons in the control of intestinal secretion.
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DECLARATION
This is to certify that:
this thesis comprises only my original work except where indicated in the preface due acknowledgement has been made in the text to all other material used; and this thesis is less than 100,000 in length, inclusive of footnotes, but exclusive of tables, maps, appendices and bibliography
Candice Fung
October 2016
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PREFACE
Under my direction, the following people have made the following contributions to my work:
Chapter 3: About 90% of the Ussing chamber experiments and all of the PCR studies were conducted during my Masters (MSc). Technical assistance and advice for PCR studies were provided Prof. Laura Parry, Ms. Maria Jelinic, and Dr. Yu May Soh. Technical assistance for Ussing chambers was provided by Dr. Jaime Foong. Some Ussing chamber experiments were performed by Ms. Petra Unterweger. Some technical assistance and advice for muscle contractility studies was provided by Ms. Rachel Gwynne. Assistance in preparing the manuscript of this chapter for publication was provided by Dr. Jaime Foong, Prof. Laura Parry, and Prof. Joel Bornstein.
Chapter 4: Technical assistance and advice for calcium imaging experiments and analyses were provided by Dr. Carla Cirillo and A/Prof. Pieter Vanden Berghe. Some advice for immunohistochemistry was provided by Dr. Carla Cirillo and Ms. Mathusi Swaminathan. Electrical stimulation calcium imaging experiments for this chapter was performed by Dr. Werend Boesmans.
Chapter 5: The majority of mouse surgeries were performed by Ms. Petra Unterweger. Some technical assistance with mouse surgeries was also provided by Prof. Andrew Allen and Ms. Katerina Koussoulas. Technical assistance and advice for c-Fos labelling experiments were provided by Ms. Petra Unterweger.
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Publications arising from this thesis include:
C. Fung, P. Unterweger, L.J. Parry, J.C. Bornstein, J.P.P. Foong. (2014) VPAC1 receptors regulate intestinal secretion and muscle contractility by activating cholinergic neurons in guinea pig jejunum. American Journal of Physiology – Gastrointestinal and Liver Physiology, 306(9), G748-58.
Abstracts (refereed*):
C. Fung, C. Cirillo, J.C. Bornstein, J.P. Foong, P. Vanden Berghe. (2016) VPAC1 activation reveals neuron-glia interactions in the submucosal plexus of mouse jejunum. 2nd Federation of Neurogastroenterology and Motility Meeting, San Francisco, USA.
C. Fung, P. Unterweger, K. Koussoulas, A.M. Allen, J.C. Bornstein, J.P. Foong. (2016) Mechanisms underlying cholera toxin-induced hypersecretion in mouse ileum. 2nd Federation of Neurogastroenterology and Motility Meeting, San Francisco, USA.
C. Fung, P. Unterweger, J.P.P. Foong, L.J. Parry, J.C. Bornstein (2014) VPAC1 receptors regulate intestinal secretion and muscle contractility by activating cholinergic neurons in guinea pig jejunum. Australasian Neurogastroenterology and Motility Association Inc. meeting, Melbourne, AU.
C. Fung, J.P.P. Foong, L.J. Parry, J.C. Bornstein. (2013) VPAC1 receptors expressed on cholinergic secretomotor neurons regulate secretion in the guinea-pig jejunum. 33rd Annual Meeting of the Australian Neuroscience Society, Melbourne, AU.
C. Fung, J.P.P. Foong, L.J. Parry, J.C. Bornstein. (2013) VPAC1 receptors localised to cholinergic secretomotor neurons regulate intestinal secretion in guinea-pig jejunum. Gastroenterology 144(5), S-296.*
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ACKNOWLEDGEMENTS
I have been very fortunate to have worked with a number of highly skilled, knowledgeable, and generous people over the course of my Ph.D. These people have contributed significantly to the work presented in this thesis and I express my thanks and gratitude here.
Firstly, my sincere thanks are due to my supervisors Joel Bornstein and Jaime Foong. This work would not have been possible without their expertise, support, and guidance throughout the course of my Ph.D. Joel, your wealth of knowledge is astounding and all your advice has been invaluable. Jaime, all your efforts in pushing me forward to finish this Ph.D. and in this career are much appreciated.
I would like to thank Laura Parry and members of her laboratory, in particular, Maria Jelinic and Yu May Soh, for their time in teaching me PCR techniques for Chapter 3.
I owe thanks to Pieter Vanden Berghe for his generosity in hosting me in his laboratory to learn calcium imaging and for making Chapter 4 possible. I am also indebted to Carla Cirillo who taught me calcium imaging, and to Werend Boesmans for performing the additional electrical stimulation experiments for Chapter 4. Many thanks to Marlene Hao and Micky Moons for organising the mice for experiments and for technical advice. I would like to extend my thanks to all LENS (Laboratory for Enteric Neuroscience) members for their support and hospitality during my time in Leuven.
I would also like to acknowledge Mastura Monif, Daniel Poole, and Trung Nguyen for their technical advice on calcium imaging.
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I am grateful to Petra Unterweger, who performed most of the mouse surgeries and made Chapter 5 possible. I would also like to acknowledge Katerina Koussoulas for her assistance with surgeries. I thank Andrew Allen for sharing his expertise and for assisting with setting up this surgical technique. I am grateful to Annette Bergner and Heather Young for providing the Wnt1-Cre;R26R-GCaMP3 mice, as well as Mai Xuan Tran for providing technical assistance and advice on genotyping. I also thank Colin Anderson for generously donating his last aliquot of anti-CT-B.
I would like to acknowledge members of the Department of Physiology, particularly the Physiology Student’s Society, for their support and for providing a friendly environment in which to work and study.
Special thanks are due to past and present members of the Bornstein lab for their technical assistance, invaluable feedback, support and friendship. In particular, I would like to again thank Petra Unterweger and Katerina Koussoulas for their contribution to the cholera toxin work and for helping me see the light. I am also grateful to Mathusi Swaminathan for technical advice on immunohistochemistry and microscopy, as well as all the coffees. I also thank my lab members for their consideration for my mental wellbeing, the laughs, and generally for keeping the lab a lively workplace.
Finally, I would like to thank my family for all their patience, encouragement and support.
Many thanks,
Candice Fung October 2016
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TABLE OF CONTENTS
TABLE OF CONTENTS ...... ix
LIST OF TABLES ...... xvii
LIST OF FIGURES ...... xviii
LIST OF ABBREVIATIONS ...... xxi
CHAPTER 1: INTRODUCTION ...... 1
1. The Enteric Nervous System ...... 2
1.1 Structure and function ...... 2
1.2 Species and region differences ...... 3
2. The Basic Mechanisms of Intestinal Secretion ...... 6
2.1 The mucosa and Cl- secretion ...... 6
2.2 Sensory transduction ...... 11
3. The Classification of Enteric Neurons ...... 13
3.1 Morphological classification of enteric neurons ...... 13
3.2 Electrophysiological classification of enteric neurons ...... 15
3.3 Neurochemical coding of enteric neurons ...... 17
3.4 Functional classes of enteric neurons ...... 20
Intrinsic sensory neurons ...... 20
Interneurons ...... 22
Secretomotor neurons ...... 24
Motor neurons...... 26
4. Enteric Reflex Circuits ...... 26
4.1 Secretomotor reflexes ...... 27
4.2 Recurrent networks of submucosal neurons ...... 28
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4.3 Motility reflexes ...... 30
4.4 Co-ordination of secretomotor and motility reflexes ...... 31
5. Enteric Glia ...... 33
5.1 Enteric glial subtypes ...... 34
5.2 Enteric glia are involved in regulating intestinal motility and barrier function .... 36
5.3 The role of enteric glia in gastrointestinal diseases ...... 37
5.4 Protective effects of enteric glia and neurogenesis ...... 37
5.5 Enteric neuron-glia signaling ...... 38
6. Neural and Glial Transmitters and Receptors ...... 39
6.1 Acetylcholine, nicotinic receptors (nAChRs), and muscarinic receptors (mAChRs) ...... 39
6.2 Noradrenaline and α2-adrenoceptors ...... 41
6.3 Somatostatin (SOM) and SST1 and SST2 receptors ...... 41
6.4 Tachykinins and neurokinin (NK) receptors ...... 42
6.5 Serotonin (5-HT) and 5-HT receptors ...... 43
6.6 ATP and purinergic P2Y1 receptors ...... 45
6.7 Nitric oxide (NO) ...... 48
6.8 Vasoactive intestinal peptide (VIP) and VIP receptors ...... 49
7. Effects of Cholera toxin (CT) ...... 51
7.1 Cholera toxin-evoked hypersecretion ...... 51
7.2 ENS involvement in CT-evoked hypersecretion ...... 52
7.3 Models of neural mechanisms underlying CT-evoked hypersecretion...... 53
7.4 Effects of cholera toxin on intestinal motility ...... 57
8. Thesis Aims ...... 58
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CHAPTER 2: MATERIALS AND METHODS ...... 59
1. Animals ...... 59
2. Identification and quantification of Vipr1 and Vipr2 messenger RNA ...... 59
2.1 Tissue preparation ...... 59
2.2 RNA extraction ...... 60
2.3 First strand complementary DNA synthesis ...... 60
2.4 Reverse transcriptase polymerase chain reaction ...... 61
2.5 Quantitative real time polymerase chain reaction (qPCR) ...... 65
3. Immunohistochemistry ...... 67
3.1 Tissue preparation ...... 67
3.2 Microscopy ...... 67
3.3 Cell counts ...... 68
4. In vitro measurement of short-circuit current in Ussing chambers ...... 68
4.1 Tissue preparation ...... 68
4.2 Electrical measurements ...... 69
4.3 Data analysis ...... 70
5. In vitro measurement of longitudinal muscle contractility ...... 70
6. Genotyping ...... 71
7. Calcium imaging ...... 75
7.1 Tissue preparation ...... 75
7.2 Imaging ...... 75
7.3 Data Analysis ...... 76
8. In vivo incubation of cholera toxin (CT) in a ligated ileal loop ...... 77
9. Cryosectioning and anti-CT-B labelling ...... 78
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10. Drugs ...... 79
CHAPTER 3: VPAC1 RECEPTORS REGULATE INTESTINAL SECRETION AND MUSCLE CONTRACILITY BY ACTIVATING CHOLINERGIC NEURONS IN GUINEA PIG JEJUNUM ...... 81
1. ABSTRACT ...... 81
2. INTRODUCTION ...... 82
3. MATERIALS AND METHODS ...... 84
3.1 Identification and quantification of Vipr1 and Vipr2 mRNA ...... 84
Tissue preparation ...... 84
RNA extraction ...... 85
Reverse transcriptase-PCR ...... 85
Quantitative real-time polymerase chain reaction ...... 85
3.2 Immunohistochemical localization of VPAC1R ...... 86
3.3 Cell counts ...... 87
3.4 In vitro measurement of longitudinal muscle contractility ...... 87
Tissue preparation ...... 87
3.5 In vitro measurement of short-circuit current in Ussing chambers ...... 88
Experimental protocol ...... 88
3.6 Data measurement and statistics ...... 89
3.7 Drugs used ...... 89
4. RESULTS ...... 90
4.1 Differential expression of Vipr1 and Vipr2 across the gut layers ...... 90
4.2 Cholinergic excitatory motor neurons and secretomotor neurons express VPAC1R ...... 91
4.3 Role of VPAC1R in longitudinal muscle contraction and intestinal secretion ...... 96
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4.4 VIP evokes longitudinal smooth muscle contractions by activating VPAC1Rs on neurons ...... 96
4.5 VIP-mediated secretion involves activation of cholinergic secretomotor neurons. 98
4.6 VIP evokes a small neurogenic secretory component involving VPAC1R ...... 99
4.7 Neurally mediated basal secretion involves VPAC1R ...... 101
5. DISCUSSION ...... 101
5.1 Exogenous VIP activates VPAC1R to evoke longitudinal muscle contraction ..... 101
5.2 Exogenous VIP activates VPAC1R on cholinergic secretomotor neurons ...... 103
5.3 Signaling pathways involving VIP, PACAP, and NO ...... 104
5.4 VPAC1R is the major receptor subtype in the mucosa ...... 105
5.5 Enhanced secretory responses with repeated agonist exposure ...... 106
5.6 Conclusion ...... 106
CHAPTER 4: VPAC RECEPTOR SUBTYPES TUNE PURINERGIC NEURON-TO- GLIA COMMUNICATION IN THE MURINE SUBMUCOSAL PLEXUS ...... 107
1. ABSTRACT ...... 107
2. INTRODUCTION ...... 108
3. MATERIALS AND METHODS ...... 109
3.1 Tissue preparation and Ca2+ imaging in submucosal plexus ...... 109
3.2 Drugs ...... 110
3.3 Analysis ...... 112
3.4 Immunohistochemistry ...... 112
3.5 Data and statistics ...... 113
4. RESULTS ...... 116
2+ 4.1 VIP exclusively evokes neuronal [Ca ]i transients in submucosal plexus ...... 116
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4.2 Inhibiting VPAC2 receptors reveals a VIP-evoked glial response ...... 116
2+ 4.3 Selective VPAC1 activation predominantly evokes [Ca ]i transients in enteric glia ...... 122
4.4 The VPAC1 receptor is expressed on cholinergic submucosal neurons and nerve fibers ...... 124
4.5 Inhibiting VPAC2 receptors reveals delayed glial responses to electrical stimulation ...... 125
4.6 VPAC1 agonist-evoked glial activation involves a TTX-insensitive mechanism .. 125
2+ 4.7 VPAC1 receptor activation stimulates glial [Ca ]i responses via purinergic signaling ...... 130
5. DISCUSSION ...... 133
5.1 VPAC2 receptors inhibit VIP-evoked glial responses...... 133
5.2 Activation of VPAC1 receptors on cholinergic neurons stimulates glial responses ...... 134
5.3 VPAC1 receptor activation stimulates glial responses via purine release ...... 135
5.4 Conclusion ...... 137
CHAPTER 5: CHOLERA TOXIN INDUCES SUSTAINED EXCITABILITY OF CHOLINERGIC SUBMUCOSAL NEURONS IN MOUSE ILEUM ...... 138
1. ABSTRACT ...... 138
2. INTRODUCTION ...... 139
3. MATERIALS AND METHODS ...... 141
3.1 Mice ...... 141
3.2 In vivo incubation of CT in a ligated ileal loop ...... 141
3.3 In vitro measurement of short-circuit current in Ussing chambers ...... 141
3.4 Ca2+ imaging and analysis ...... 142
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3.5 Drugs ...... 143
3.6 Immunohistochemistry ...... 143
3.7 Confocal imaging and cell counts ...... 144
3.8 Data and statistics ...... 146
4. RESULTS ...... 146
4.1 Acute CT exposure induces a sustained increase in basal secretion ...... 146
4.2 CT-incubation induced pCREB expression in submucosal neurons ...... 150
4.3 CT-exposure increases sustained excitability in cholinergic submucosal neurons ...... 152
2+ 4.4 CT-exposure did not alter VPAC1 agonist-evoked glial [Ca ]i responses ...... 155
4.5 CT-exposure induces c-Fos expression in myenteric plexus ...... 155
4.5 The myenteric plexus displays an overall reduction in neuronal excitability following CT-exposure ...... 157
4.6 Effects of CT were confined to the ileal loop ...... 159
5. DISCUSSION ...... 163
5.1 CT primarily acts on the mucosa epithelium to induce hypersecretion ...... 163
5.2 CT induced sustained hyperexcitability in cholinergic submucosal neurons ...... 164
5.3 Some myenteric neurons may be hyperactive during but not following CT incubation ...... 167
5.4 Conclusion and future directions ...... 168
CHAPTER 6: DISCUSSION ...... 170
1.1 Neural VPAC1 receptors regulate intestinal secretion and muscle contractility . 171
1.2 VIP and VPAC receptors regulate neuron-to-glia signaling ...... 172
1.3 ENS involvement in the sustained effects of acute cholera toxin exposure ...... 174
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1.4 The role of cholinergic and non-cholinergic secretomotor neurons in the regulation of intestinal secretion ...... 177
1.5 Conclusion ...... 177
CHAPTER 7: LIST OF REFERENCES ...... 179
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LIST OF TABLES
Table 1.1. A comparison of the types of submucosal neurons found in guinea pig vs. mouse ileum...... 18
Table 1.2. A comparison of the types of myenteric neurons found in guinea pig vs. mouse ileum...... 19
Table 2.1. RT-PCR primer sequences for guinea pig Vipr1, Vipr2, and gapdh...... 63
Table 2.2. Guinea pig Vipr1 and Vipr2 receptor nucleotide and amino acid sequences...... 64
Table 2.3. qPCR primer and probe sequences for guinea pig Vipr1, Vipr2, and rat Rn18s...... 66
Table 2.4. Drugs and their targets...... 80
Table 4.1. Primary antibodies used for immunohistochemistry...... 114
Table 4.2. Secondary antibodies used for immunohistochemistry...... 115
Table 5.1. Primary and secondary antibodies used for immunohistochemistry...... 145
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LIST OF FIGURES
Figure 1.1. Diagram of a peeled apart wall of the small intestine...... 5
Figure 1.2. Components of the small intestinal mucosa...... 7
Figure 1.3. Schematic diagram of the signaling systems regulating Cl- secretion (crypt cells) and release of 5-HT (EC cells) in the intestinal epithelium ...... 10
Figure 1.4. Confocal micrographs depicting the morphology of myenteric neurons found in the guinea pig small intestine: Dogiel types I and II, and filamentous neurons...... 14
Figure 1.5. Intracellular recording traces comparing the electrophysiological properties of an S neuron and AH neurons...... 16
Figure 1.6. Morphological subtypes of enteric glia...... 35
Figure 1.7. A simplified model of the enteric circuitry activated by CT...... 55
Figure 2.1. Identifying the presence or absence of Wnt1-cre expression...... 74
Figure 3.1. mRNA expression of vasoactive intestinal peptide receptor 1 gene (Vipr1) and vasoactive intestinal peptide receptor 2 gene (Vipr2) in the gut layers of guinea pig jejunum...... 93
Figure 3.2. Localizing VPAC1 receptor (VPAC1R) in the myenteric plexus of guinea pig jejunum...... 94
Figure 3.3. Localizing VPAC1R in the submucosal plexus of guinea pig jejunum...... 95
Figure 3.4. Effects of various antagonists on vasoactive intestinal peptide (VIP)- induced contraction in longitudinal muscle and myenteric plexus preparations...... 97
Figure 3.5. Effects of various antagonists on the second phase of VIP-induced change in short-circuit current (ΔISC)...... 100
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2+ Figure 4.1. VIP-induced [Ca ]i transients in enteric neurons and glia...... 119
Figure 4.2. VIP (100 µM)-evoked glial and neuronal responses in the presence of various antagonists...... 121
2+ Figure 4.3. VPAC1-induced [Ca ]i transients in enteric neurons and glia...... 123
Figure 4.4. Localizing VPAC1 receptor (VPAC1R)-expression in the submucosal plexus of mouse jejunum...... 127
Figure 4.5. Confocal images of VPAC1 receptor (VPAC1R)-immunoreactive nerve fibers and varicosities in the submucosal plexus of mouse jejunum taken at a single optical plane...... 128
2+ Figure 4.6. VPAC1 agonist (100 µM)-evoked glial and neuronal [Ca ]i responses in the presence of antagonists...... 129
2+ Figure 4.7. P2Y1 agonist 2MeSADP (100 µM)-evoked glial [Ca ]i responses...... 132
Figure 5.1. CT-treated vs. control ileal loops following 3.5 h incubation in vivo...... 149
Figure 5.2. pCREB labelling of submucosal neurons in control vs CT-treated ileal loops...... 151
Figure 5.3. Spontaneous activity in submucosal neurons following CT-incubation. . 153
2+ Figure 5.4. DMPP and electrically-evoked [Ca ]i responses in submucosal neurons are increased following CT-exposure...... 154
Figure 5.5. c-Fos labelling in myenteric neurons in control vs. CT-treated ileal loops...... 156
2+ Figure 5.6. DMPP- and electrically-evoked [Ca ]i responses in myenteric neurons are decreased following CT-exposure...... 158
Figure 5.7. c-Fos labelling of myenteric neurons were confined to the CT-treated ileal loop...... 161
Figure 5.8. Changes in excitability of enteric neurons following CT-exposure were confined to the ileal loop...... 162
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LIST OF ABBREVIATIONS
5-HT 5-hydroxytryptamine, serotonin
ACh Acetylcholine
AHP After-hyperpolarizing potential
ATP Adenosine triphosphate bp Base pairs
Ca2+ Calcium
2+ [Ca ]i Intracellular calcium cAMP Cyclic adenosine monophosphate
CCK Cholecystokinin
CFTR Cystic fibrosis transmembrane conductance regulator cGMP Cyclic guanosine monophosphate
CGRP Calcitonin gene-related peptide
ChAT Choline acetyltransferase
Cl- Chloride
CM Circular muscle
CMMC Colonic migrating motor complex
CNS Central nervous system
CT Cholera toxin
CT-B Cholera toxin (B-subunit)
EC Enterochromaffin
ENS Enteric nervous system
EPSP Excitatory postsynaptic potential
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DMPP 1,1-dimethyl-4-phenylpiperazinium
GFAP Glial fibrillary acidic protein
GPCR G protein-coupled receptor
HEX Hexamethonium
HYO Hysocine
IPAN Intrinsic primary afferent neuron
ISC Short circuit current
ISN Intrinsic sensory neuron
IPSP Inhibitory postsynaptic potential
IR Immunoreactive, immunoreactivity
KCl Potassium chloride
LMMP Longitudinal muscle-myenteric plexus
MMC Migrating myoelectric complex
MP Myenteric plexus mRNA Messenger ribonucleic acid
NA Nicotinic acetylcholine receptor nAChR Noradrenaline
NO Nitric Oxide
NOS Nitric oxide synthase
NPY Neuropeptide Y
PACAP Pituitary adenylate cyclase activating peptide
PBS Phosphate buffered saline
PCR Polymerase chain reaction
PD Potential difference
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PPADS Pyridoxal phosphate-6-azophenyl-2’,4’-disulphonic acid qPCR Quantitative polymerase chain reaction
RT-PCR Reverse-transcriptase polymerase chain reaction
SEM Standard error of the mean
SMP Submucosal plexus
SOM Somatostatin
SP Substance P
TH Tyrosine hydroxylase
TTX Tetrodotoxin
VPAC VIP and PACAP receptor
VIP Vasoactive intestinal peptide
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CHAPTER 1: INTRODUCTION
The human gastrointestinal tract receives up to 8 L of secreted fluid daily – secretions from the small intestine largely contribute to this volume, but it also consists of gastric, pancreatic and biliary secretions (Barrett and Keely, 2000; Murek et al., 2010). While the majority of fluid is re-absorbed in the small intestine, the concurrent process of active fluid secretion is important for intestinal function. This secreted fluid facilitates digestion by providing a solvent in which enzymes, as well as the water-soluble products of broken- down carbohydrates and proteins, can diffuse (Barrett and Keely, 2000). Further, the fluid reduces the viscosity of the luminal contents and aids in mixing. Physiological fluid secretion also contributes to maintaining an optimal pH in the luminal environment and provides lubrication to the mucosal epithelium to aid in the movement of contents along the gastrointestinal tract (Murek et al., 2010). However, intestinal secretion can become dysregulated with exposure to noxious agents such as bacterial enterotoxins, viruses, and parasites. These pathogens can trigger hypersecretion in the gastrointestinal tract, leading to diarrhoea and rapid dehydration with an uncontrolled loss of the body’s fluid and electrolytes (Lundgren and Jodal, 1997; Lundgren et al., 2000). It has been proposed that this is an inherently protective mechanism and attempt to eliminate the source of irritation. Yet, if fluids are not replenished, it can be life-threatening. Diarrhoeal disease is a global problem with over a billion cases reported annually, and is a leading cause of death in children under five (WHO, 2013). Thus, intestinal secretion is a tightly regulated process important for maintaining body fluid balance.
While there are various causes of diarrhoeal disease, a commonality of many diarrhoea- inducing agents is that they act by disrupting the normal activity of the enteric nervous system (ENS) – the intrinsic nervous system of the gastrointestinal tract. It is well established that the ENS is important in diarrhoeal disease (Lundgren, 2002), yet the neural circuits involved are yet to be fully characterized. Cholera has been considered the model
1 for diarrhoeal disease and is caused by various strains of the bacterium Vibrio cholerae. Cholera is an acute infection of the small intestine which results in watery diarrhoea, vomiting, and subsequent dehydration. While this may be treated with oral rehydration therapy, it remains an issue particularly given that cholera typically presents in areas lacking a clean water supply. There are also vaccines available but presently these provide protection for only a limited time and booster doses are necessary to maintain protection. More effective treatments are necessary in the event of outbreaks.
Animal model studies have highlighted the effectiveness of selective antagonists targeting specific enteric pathways which drive hypersecretion, as potential anti-secretory therapies (Mourad et al., 1995; Turvill and Farthing, 1997; Turvill et al., 2000b; Farthing et al., 2004; Banks et al., 2005).
Thus, the aim of this thesis is to gain a more comprehensive understanding of the enteric circuitry underlying physiological intestinal secretion, and of how this system is disrupted in pathophysiology, as it is imperative to advancing the development of anti-diarrhoeal therapeutics (Farthing, 2002).
1. The Enteric Nervous System
1.1 Structure and function
The ENS is situated within the walls of the gastrointestinal tract and controls physiological intestinal function. It is a highly elaborate network comprising vast numbers of neurons and glial cells; the ENS contains some 200 – 600 million neurons in humans (Furness et al., 2014) and over 1.2 million neurons in mice (Obermayr et al., 2013). Enteric neurons and
2 glia are arranged in the form of small clusters or ‘ganglia’, which are interconnected by nerve strands and form the basic elements of this network. The ENS consists of two major ganglionated nerve plexuses: the submucosal and myenteric plexuses (Figure 1.1). The submucosal plexus lies between the circular muscle and mucosal epithelium, and regulates intestinal secretion. The myenteric plexus is embedded between the longitudinal muscle and circular muscle layers, and is primarily responsible for regulating intestinal motility. The two plexuses are interconnected and this is necessary for coordinating changes in secretion and motility for normal intestinal function. The ENS is unique in its ability to operate independently of central nervous control, although the two systems are integrated via the extrinsic nerves.
1.2 Species and region differences
While the plexuses of the ENS can be readily identified by their characteristic meshwork, enteric ganglia vary in size, shape, and organization, both between different species and intestinal regions (Furness, 2006). Further, the constituent neurons may also differ in function and neurochemistry, thus reflecting functional specialization (McConalogue and Furness, 1994; Furness et al., 1995b).
The submucosal plexus of guinea pig and other small mammals comprises a single layer of submucosal ganglia containing neurons involved in secretion, but not motor neurons that innervate muscle (Furness et al., 1984). By contrast, the submucosal plexus of larger animals (e.g. pig and human) consists of two or three interconnected layers of ganglia (Figure 1.1), with each layer containing different complements of neurons, which may be involved in both secretion and motility (Timmermans et al., 1997; Hens et al., 2001; Timmermans et al., 2001). Such differences in the arrangement of submucosal ganglia and the functional types of neurons they contain make it challenging to extrapolate from one animal model to another.
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The myenteric plexus can be divided into a primary, secondary, and tertiary plexus (Furness, 2006). The myenteric ganglia and interconnecting fiber tracts make up the primary plexus. The secondary plexus runs parallel to the circular muscle and comprises of finer nerve strands arising from ganglia or primary fiber tracts and which generally do not connect adjacent ganglia. Finally, the tertiary plexus is formed of fine fiber strands that branch from the primary and secondary fiber tracts, or arise from ganglia, and are found in the spaces between the primary meshwork. In species or intestinal regions with a thickened longitudinal muscle layer, a longitudinal muscle plexus is observed. While in regions where the longitudinal muscle is thin, it is exclusively supplied by the tertiary plexus (Furness, 2006). Hence, species and region differences are important considerations in interpreting data and generalizing findings.
Much of the current knowledge about the ENS is based on that of the guinea pig intestine; it was the preferred animal model due to the ease in separation of its gut layers and accessibility of its neurons. In fact, the constituent neurons of guinea pig small intestine are fully characterized (Furness et al., 2003) and the guinea pig ENS remains the most well understood compared to that of other species. Many of the peptides which form the chemical coding of enteric neurons differ significantly in cross-species comparisons (Keast et al., 1985a; Bornstein et al., 2012), but the guinea pig ENS remains a useful model as the primary enteric neurotransmitters identified in the guinea pig are conserved across mammalian species (Furness et al., 1995b). Nonetheless, with the recent advances in gene technologies and the ever-increasing lines of transgenic mouse strains, there is a rapid shift towards using mutant mouse models of gastrointestinal diseases or mice that are encoded with reporter genes. Moreover, mice are a good animal model given their genetic similarities to humans (Waterston et al., 2002). In addition, mice share common immune responses with humans and have been used extensively in the study of inflammatory bowel disease (Mizoguchi et al., 2003; Lin and Hackam, 2011; Jiminez et al., 2015). Thus, there is a need to better characterize the murine ENS.
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Figure 1.1. Diagram of a peeled apart wall of the small intestine.
This illustrates the representative organization of the Enteric Nervous System (ENS) of human and larger animals. The mucosa lines the lumen of the gut and is situated adjacent to the submucosal plexus; together they mediate secretory function. The submucosal plexus may contain two or three interconnected layers of ganglia. Submucosal neurons contained in the outer plexus situated closest to the circular muscle, project to the muscle layers and to the mucosa. This outer plexus may be involved in the control of both intestinal secretion and motility. Submucosal neurons in the inner plexus closest to the mucosa, primarily innervate the mucosa and are mainly involved in secretion. The myenteric plexus lies between the circular and longitudinal muscle layers, and this arrangement attests to its primary role in controlling intestinal motility. The small intestines of small laboratory animals generally have a similar structure; however, a key difference is that the submucosal plexus is formed from one single layer of ganglia primarily containing neurons involved in secretion, and is similar to the inner submucosal plexus of larger mammals. Figure reproduced from (Furness, 2012).
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2. The Basic Mechanisms of Intestinal Secretion
2.1 The mucosa and Cl- secretion
The mucosa comprises the muscularis mucosae, lamina propria (connective tissue), and the mucosal epithelium. The mucosal epithelium lining the lumen of the gastrointestinal tract is the site at which absorption and secretion of fluid and electrolytes occur. It also acts as a barrier to the external environment (Cooke, 2000). In the small intestine, the mucosa forms a series of folds from which arise villi that project into the lumen, separated by crypts where the epithelium dips into the lamina propria (Laster and Ingelfinger, 1961) (Figure 1.2). A continuous epithelial layer lines the villi and the crypts, but epithelial cells are heterogeneous. The epithelium comprises of absorptive columnar cells (enterocytes) which line the villi, crypt cells, sparsely interspersed enteroendocrine cells, and other specialized cells (e.g. mucous-producing goblet cells) (Madara, 2010). Villi have a major role in absorption, and the crypts are generally associated with secretion. Enteroendocrine cells have chemosensory functions; they can detect and respond to nutrients via the release of various peptides and hormones (Gribble and Reimann, 2016). Within the lamina propria is an extensive network of predominantly enteric nerve fibers that supply the mucosa; the majority of nerve fibers supplying the mucosa originate from submucosal neurons, although there are some projections from myenteric neurons (Keast, 1987) as well as extrinsic nerves.
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Figure 1.2. Components of the small intestinal mucosa.
Light micrograph (A) and corresponding trace drawing (B) illustrating components of the small intestinal mucosa. The mucosa comprises of muscularis mucosa, lamina propria, and epithelium. The intestinal epithelium can be divided into two major compartments: the villi and crypts. Villi project into the lumen and increase the absorptive epithelial surface area. Separating the villi are crypts, which dip into the lamina propria. The epithelium lining the villi and crypts forms one continuous layer, however, they are composed of distinctly different cell types. The major cell types of villi are absorptive enterocytes and goblet cells, whereas crypt cells include undifferentiated cells, goblet cells, and basally located Paneth cells. Other specialized epithelial cell types such as enteroendocrine cells may be scattered throughout epithelium but are not well depicted here. Figure taken from Madara (2010).
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Fluid flux is driven by the osmotic gradient established by increasing electrolyte concentrations and water follows passively down this concentration gradient across the mucosa into the intestinal lumen (Burleigh and Banks, 2007; Murek et al., 2010). Actively secreted chloride (Cl-) is the primary ion maintaining the osmotic gradient and hence the major determinant of net fluid secretion in the intestine (Barrett and Keely, 2000; Burleigh and Banks, 2007; Xue et al., 2007; Murek et al., 2010). Cl- uptake into mucosal epithelial cells occurs via Na+/K+/2Cl-(NKCC1) co-transporters on the basolateral membrane (Figure 1.3). The accumulated Cl- is then secreted via apical Cl- channels (Barrett and Keely, 2000).
Three channels through which Cl- secretion may occur have been identified to date. The most extensively studied is the cystic fibrosis transmembrane regulator (CFTR) Cl- channel which is activated by cAMP or cGMP (Xue et al., 2007; Murek et al., 2010) (Figure 1.3). CFTR mutants and CFTR knockout mouse models show a decrease in, or total absence of, cAMP-mediated Cl- secretion in both in vivo and in vitro studies (Clarke et al., 1992; Ratcliff et al., 1993; Grubb and Gabriel, 1997; Murek et al., 2010). Further, CFTR inhibitors effectively reduce CT-induced secretion in rodent models (Thiagarajah et al., 2004; Muanprasat et al., 2012). cAMP-mediated Cl- secretion has a slow onset, taking up to 15 min to reach a peak response (Xue et al., 2007), which is also compatible with the slow time course of CT-evoked secretion (Carey and Cooke, 1986; Mourad et al., 1995).
Another channel is a Ca2+-activated Cl- channel (CACC) (Murek et al., 2010). Whereas bacterial enterotoxins have been associated with CFTR, viral enterotoxins and in particular rotavirus have shown to exert secretory effects involving CACCs (Thiagarajah and Verkman, 2013; Ko et al., 2014). Ca2+-mediated Cl- secretion is transient and has a fast onset (Xue et al., 2007). However, the molecular identity of the CACC involved is unknown. The human CACC hClCA1 was previously considered a candidate, as it was shown to be expressed by crypt cells (Gruber et al., 1998); however, subsequent studies showed that it is a secretory protein rather than an integral membrane protein and hence is
8 not considered a proper functional channel (Gibson et al., 2005; Murek et al., 2010). TMEM16A has also been assessed as a potential candidate, but while it plays a role in intestinal motility (Singh et al., 2014), its contribution to Ca2+-activated Cl- conductance in the intestinal epithelium is less clear (Namkung et al., 2011; Huang et al., 2012; Thiagarajah et al., 2015).
A third channel identified is chloride channel type-2 (CIC-2). However, its role remains controversial given its unusual expression along the lateral membrane of enterocytes of rodents (Gyomorey et al., 2000; Gyomorey et al., 2001), as well as discrepancies when compared to human tissue where CIC-2 was localised to an intracellular compartment of colonic cells (Lipecka et al., 2002; Murek et al., 2010). Further, compared to CFTR knockout mice, ClC-2 – CFTR double knockout mice lacking both channels did not appear to show any further alterations to Cl- transport across colonic epithelia measured in vitro (Zdebik et al., 2004; Murek et al., 2010). Therefore, the physiological contribution of CIC- 2 to intestinal Cl- secretion remains unclear.
All mucosal epithelial cells perform both absorptive and secretory functions to an extent. Nevertheless, the tips of the villi which are directly exposed to the luminal contents are the specialised structures for absorption of fluid and nutrients, whereas the crypts are the primary effector cells mediating Cl- flux (Barrett and Keely, 2000). Secretion is thought to occur in a decreasing gradient toward the tips of the villi. This matches the graded pattern of CFTR expression, where it is most highly expressed in the crypts and becomes lower towards the villi (Strong et al., 1994; Ameen et al., 1995). The location of the villi also allows the local control of transport processes through a direct interaction with the luminal contents. In contrast, the crypts being less accessible to luminal contents depend on indirect regulatory mechanisms, namely secretomotor pathways within the ENS (Lundgren and Jodal, 1997) and hence the dense network of nerve fibers supplying the mucosa (Furness, 2006).
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CIC-2 CACC
Figure 1.3. Schematic diagram of the signaling systems regulating Cl- secretion (crypt cells) and release of 5-HT (EC cells) in the intestinal epithelium.
A. Cl- secretion through Cl- channels including CFTR and others (CIC-2 and CACC). Cl- secretion depends on the electrochemical driving forces. B. Acetylcholine activates muscarinic M3 receptors on the basolateral membrane of crypt cells to stimulate secretion via intracellular Gαq signaling and mobilization of Ca2+. VIP activates VPAC1 receptors to stimulate secretion via Gαs signaling pathways and elevation of cAMP levels. C. Enterochromaffin cell model (human BON cell). ATP is thought to be involved in transducing the signal from mechanical stimulation of the mucosa and in activating secretory reflex pathways by triggering 5-HT release. Figure adapted from Xue et al. (2007).
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2.2 Sensory transduction
Sensory nerve endings are protected from the hostile environment of the lumen by the intestinal epithelial barrier, thus “sensors” in the mucosa are necessary to detect luminal stimuli and serve as an interface connecting the ENS to the external environment (Blackshaw et al., 2007). However, some nutrients are actively transported across the epithelium (e.g. glucose) and lipophilic solutes may cross by passive diffusion (e.g. capsaicin) (Bertrand, 2009). Enterochromaffin (EC) cells are considered to be one such “sensor” – they are a prominent type of enteroendocrine cell in the intestine. EC cells are widely distributed, though they are interspersed between enterocytes and other enteroendocrine cells (Schafermeyer et al., 2004; Kidd et al., 2006). EC cells synthesize and store serotonin (5-HT) in high concentrations (Gershon and Tack, 2007). In response to chemical and mechanical stimuli, EC cells release 5-HT (Kirchgessner et al., 1992; Foxx- Orenstein et al., 1996; Bertrand, 2009), which is thought to be a key mediator in linking the luminal environment to the enteric circuitry. 5-HT released across the basolateral membrane stimulates 5-HT1P (Pan and Gershon, 2000) and/or 5-HT4 (Kellum et al., 1999) receptors on terminals of submucosal sensory neurons supplying the mucosa, as well as 5-
HT3 receptors on terminals of myenteric sensory neurons (Bertrand et al., 2000a; Gershon, 2004). This may in turn activate secretomotor reflexes (Kirchgessner et al., 1992; Sidhu and Cooke, 1995; Cooke et al., 1997a; Kellum et al., 1999; Pan and Gershon, 2000), and/or facilitate motility reflexes (Foxx-Orenstein et al., 1996; Tuladhar et al., 1997; Bertrand, 2009; Gwynne et al., 2014) in human and guinea pig intestine. There is also some evidence for ATP as another sensory mediator that stimulates mucosal sensory nerve endings via P2X receptors (Bertrand and Bornstein, 2002; Bertrand, 2003). ATP may be released from enterocytes in response to mechanical stimulation (Cooke et al., 2003). While 5-HT and ATP can both activate sensory nerve endings supply the mucosa directly, ATP appears to also stimulate 5-HT release (Bertrand, 2003).
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Other types of enteroendocrine cells may release a variety of mediators in response to nutrients, including cholecystokinin (CCK), secretin, glucose-dependent insulinotropic polypeptide (GIP), neurotensin, peptide YY (PYY), and glucagon-like peptides 1 and 2 (GLP-1 and GLP-2) (Engelstoft et al., 2013; Gribble and Reimann, 2016). Receptors for many of these mediators have also been identified on enteric nerves (Jackerott and Larsson, 1997; Schutte et al., 1997; Guan et al., 2006; Richards et al., 2014). For instance, CCK release from enteroendocrine cells is associated with sensing lipids and proteins (Raybould, 1999; Gribble and Reimann, 2016), and there is some evidence that CCK may activate intrinsic enteric reflexes via CCK-1 and CCK-2 receptors (Schutte et al., 1997; Sayegh and Ritter, 2003; Gulley et al., 2005; Ellis et al., 2013). GLP-1 release can also be stimulated by the presence of lipids or protein, as well as carbohydrates, and their receptor has been localised to enteric neurons (Richards et al., 2014). Further, GLP-1 has been shown to modulate neurogenic secretion (Baldassano et al., 2012). However, the understanding of the role of these mediators in nutrient-sensing, the neural pathways that they may activate and the signaling mechanisms involved is still limited.
Subsequent to activation of intrinsic pathways by stimulation of nerve endings in the mucosa, secretomotor neurons – the efferent neurons contained within the submucosal plexus – can stimulate intestinal secretion. The two major transmitters are released by secretomotor neurons at epithelial junctions are acetylcholine (ACh) and vasoactive intestinal peptide (VIP) (Figure 1.3). ACh stimulates muscarinic M3 receptors and signals via the Ca2+ pathway to evoke a rapid and transient response, while VIP acting at mucosal VPAC1 receptors stimulates cAMP-mediated Cl- secretion with slower kinetics (Xue et al., 2007).
To appreciate the complexities of intestinal secretion and its regulation, it is necessary to understand the specific components of secretomotor pathways activated in physiological
12 and pathophysiological contexts. The classification of enteric neurons, with an emphasis on neurons involved in secretory reflexes, is summarized below.
3. The Classification of Enteric Neurons
The ENS contains many different types of neurons which can be classed according to their morphology, electrical properties, function, or neurochemistry (Tables 1.1 and 1.2). While these classification systems are largely established on work conducted in the guinea pig, there are efforts to better characterize the mouse ENS with the increasing use of transgenic and mutant mouse models. Where possible, comparisons between the two animal models are included in the following sections, though many inferences are drawn from knowledge of the guinea pig circuitry.
3.1 Morphological classification of enteric neurons
The characterization of enteric neurons by morphology is based on the work of Dogiel (1899), who was first to illustrate and define three neuron types in the ENS: Dogiel types I, II, and III. However, other neuronal morphologies have been identified since and this classification system has been extended to include types IV, V, VI, and VII, as well as filamentous neurons and small neurons (Brehmer et al., 1999).
Type I neurons are characterized by a uniaxonal appearance with numerous short lamellar dendrites, and Type II neurons have a smooth soma and multiple axons that tend to project circumferentially (Figure 1.4). The presence of both type I and type II neurons has been confirmed using modern methodologies across a range of species (Furness, 2006). Unlike the first two types, Dogiel’s description of type III neurons has not been definitively
13 confirmed by current methodologies, nonetheless type III is occasionally used to define filamentous neurons with long branched processes (Furness, 2006).
Figure 1.4. Confocal micrographs depicting the morphology of myenteric neurons found in the guinea pig small intestine: Dogiel types I and II, and filamentous neurons.
A. Type II neurons are described as having a smooth soma and multiple long axons that tend to project circumferentially. B. Type I neurons are defined by a uniaxonal appearance with numerous short lamellar dendrites. Unlike the other types, Dogiel’s description of type III neurons has not been definitively confirmed by current methods. Nevertheless type III is sometimes used to define (C) filamentous neurons with long branched processes (Furness, 2006). Figure taken from Clerc et al. (1998).
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3.2 Electrophysiological classification of enteric neurons
Another means of classifying enteric neurons is by examining their electrical behaviour (Figure 1.5). Based on a system introduced by Hirst and colleagues (1974), enteric neurons can be grouped into AH- and S-type neurons, depending on their electrophysiological properties. AH neurons are typified by their larger action potentials with an inflection on the falling phase, and these are generally followed by a long late after-hyperpolarizing potential (AHP; >2s) (Furness, 2006). By contrast, S neurons characteristically have short action potentials that lack slow AHPs, and exhibit fast excitatory postsynaptic potentials (EPSPs). This classification scheme also corresponds well with Dogiel’s morphological classes, where AH neurons have Dogiel type II morphology, while S neurons are uniaxonal and include Dogiel type I and filamentous neurons (Bornstein et al., 1994).
The electrophysiological properties of enteric neurons in guinea pig small intestine have been extensively characterized (Bornstein et al., 1988; Bornstein and Furness, 1988; Iyer et al., 1988; Clerc et al., 1998). However, there is limited literature detailing the electrical properties of enteric neurons in mouse small intestine (Ren et al., 2003; Mao et al., 2006; Foong et al., 2012), and this is particularly so for submucosal neurons. In fact, the electrophysiology of murine submucosal neurons has only been examined in the colon to date. Interestingly, these studies identified S-type, but not AH-type, neurons using electrophysiological classification (Wong et al., 2008; Foong et al., 2014). Further, although most neurons were uniaxonal, both studies reported a small population of neurons with two axons. By contrast, the electrophysiology and correlating morphological classifications demonstrated in mouse myenteric plexus appears to correspond well with what is established in the guinea pig (Nurgali et al., 2004; Mao et al., 2006).
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Figure 1.5. Intracellular recording traces comparing the electrophysiological properties of an S neuron and AH neurons.
Intracellular recording traces of action potentials fired by an S neuron (A) and AH neurons (B – D). This electrophysiological classification for enteric neurons was first introduced by Hirst et al. (1974). (A) S neurons display short action potentials followed by short duration (20 – 100 ms) after-hyperpolarizing potentials (AHPs). By contrast, AH neurons exhibit larger action potentials with an inflection on the falling phase and generally followed by an early AHP and a longer late AHP (2 – 30 s) as shown in B – D. Figure taken from Furness (2006).
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3.3 Neurochemical coding of enteric neurons
Different subtypes of enteric neurons are equipped with unique sets of molecules – such as the primary transmitters used by the neurons and their synthesizing enzymes, or calcium- binding proteins – which provide them with a distinct neurochemical identity that can be identified by immunohistochemical methods (Furness, 2006). This unique neurochemical coding of neurons from both enteric plexuses can be used to discern different functional subtypes, as summarized for the guinea pig and mouse ileum in Tables 1.1 and 1.2. This is a useful means of identifying neurons as primary transmitters are generally well conserved between gut regions and between species; however, the specific neurochemical coding can vary. For instance, neuropeptide Y (NPY) marks a specific subset of cholinergic secretomotor neurons in guinea pigs, while NPY co-localizes with VIP rather than choline acetyltransferase (ChAT, synthesizing enzyme for ACh) in secretomotor neurons in rats (Cox et al., 1994) and mice (Mongardi Fantaguzzi et al., 2009). Another key difference is the expression of calcitonin gene-related peptide (CGRP) in myenteric type II neurons of the mouse, but not the guinea pig (Qu et al., 2008).
Of note, as with the lack of electrophysiological evidence for the presence of AH neurons in the submucosal plexus of mouse colon, the immunohistochemical study by Mongardi Fantaguzzi et al. (2009) also reported an absence of type II neurons in the submucosal plexus of mouse ileum. This suggests that the wiring of the murine submucosal circuitry may differ significantly to that of the guinea pig.
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Table 1.1. A comparison of the types of submucosal neurons found in guinea pig vs. mouse ileum.
Guinea pig Ileum Mouse Ileum
Functional type Neurochemical code Morphology Proportion* Functional type Neurochemical code Proportion†
ChAT/TK/NeuN/ Intrinsic sensory Non-cholinergic Type II 11% neuron calbindin secretomotor VIP/NPY/calretinin ~30% neuron
Non-cholinergic VIP/CRF/GAL/ secretomotor/ Type I 45% Non-cholinergic VIP/NPY/calretinin/TH ~20% vasodilator PACAP vasodilator neuron neuron
Cholinergic secretomotor/ ChAT/calretinin Stellate 15% ChAT/SOM/CGRP +/- NPY ~30% vasodilator Cholinergic neuron secretomotor neuron Cholinergic ChAT/NPY/CCK/SOM/ secretomotor Type IV 29% ChAT ~10% CGRP neuron
Interneuron Uni-axonal projecting to the VIP with thin 1% Unknown Neither ChAT nor VIP ~8% myenteric plexus dendrites
Abbreviations: ACh acetylcholine; CCK cholecystokinin, ChAT choline acetyltransferase, CGRP calcitonin gene-related peptide; CRF corticotrophin- releasing factor; GAL galanin; NeuN neuronal nuclear protein; NYP neuropeptide Y; PACAP pituitary adenylyl cyclase activating peptide; SOM somatostatin; TH tyrosine hydroxylase; TK tachykinin; VIP vasoactive intestinal peptide. Guinea pig small intestine table adapted from Furness (2006). Mouse small intestine table based on work by Mongardi Fantaguzzi et al. (2009). *Proportion of total submucosal neurons marked by a non-specific stain for cholinesterases (Wilson et al., 1981; Furness et al., 1984); †Proportion of total submucosal neurons as marked by Hu (Mongardi Fantaguzzi et al., 2009).
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Table 1.2. A comparison of the types of myenteric neurons found in guinea pig vs. mouse ileum. Guinea pig Ileum Mouse Ileum Functional type Neurochemical code Morphology Proportion* Neurochemical code Morphology Proportion†
Intrinsic sensory ChAT/TK/NeuN/ ChAT/NF-M/CGRP/ Type II 26% Type II 26% neuron 80% calbindin calbindin +/- calretinin
Inhibitory circular muscle motor NOS/VIP Type I 16% NOS/VIP +/- NPY Type I 23% neuron Inhibitory Small, no longitudinal muscle NOS/VIP Small type I ~2% NOS/VIP obvious 3% motor neuron dendrites
Excitatory circular Small/medium, muscle motor ChAT/TK Medium Type I 12% ChAT/TK +/- calretinin no obvious 21% neuron dendrites
Excitatory Small, no longitudinal muscle ChAT/TK/calretinin Small Type I 25% ChAT/calretinin +/- TK obvious 13% motor neuron dendrites
ChAT/NOS/VIP Type I 5% ChAT/NOS/VIP Type I 3%
Descending ChAT/5-HT Type I 2% ChAT/5-HT Type I 1% interneurons Filamentous Filamentous ChAT/SOM 4% ChAT/SOM/calretinin 4% dendrites dendrites
Ascending ChAT/TK/calretinin Type I 5% ChAT/TK +/- calretinin Type I 4% interneurons
Abbreviations: 5-HT serotonin; ACh acetylcholine; CGRP calcitonin gene-related peptide; NeuN neuronal nuclear protein; NF-M neurofilament (medium); NYP neuropeptide Y; NOS nitric oxide synthase; SOM somatostatin; TK tachykinin; VIP vasoactive intestinal peptide. Table adapted from Qu et al. (2008), and incorporates findings from Sang & Young (1996, 1998). *Proportion of total myenteric neurons labelled with a NCB (neuronal cell body) antibody raised against the N-terminal of neurokinin B (Costa et al., 1996); †Proportion of total myenteric neurons marked by Hu (Qu et al., 2008).
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The ENS contains all the basic elements for a complete neural circuit: intrinsic sensory neurons (ISNs), interneurons, and motor neurons.
3.4 Functional classes of enteric neurons
Intrinsic sensory neurons
The intrinsic sensory neuron (ISN) is the afferent limb of enteric circuits necessary for monitoring the state of the gut. ISNs may also be referred to as intrinsic primary afferent neurons (IPANs). ISNs can transduce and encode the intensities, durations and patterns of physiological stimuli including mechanical distortion and luminal chemicals, and can also detect and trigger protective responses to noxious stimuli (Furness et al., 2004a). The term ISNs generally corresponds to AH neurons with Dogiel type II morphology (Bornstein et al., 1994; Furness et al., 2004a).
Myenteric ISNs detect mechanical distortion of their processes by mechanosensitive ion channels that open with muscle distortion to cause local depolarization and hence initiate action potential firing (Kunze et al., 1998; Kunze et al., 2000). These neurons also receive synaptic inputs; most myenteric AH neurons display slow EPSPs, whereas fast EPSPs are less common and, when observed, tend to be smaller than those seen in S neurons (Kunze et al., 1993; Furness et al., 1998; Blackshaw et al., 2007; Foong et al., 2012). Functionally and morphologically analogous neurons have been identified across different species and regions, although they may differ in their neurochemistry (Furness et al., 2004a). In particular, equivalent myenteric AH/Dogiel type II neurons have been characterized in the mouse small and large intestines. The murine neurons are mechanosensitive and have similar electrophysiological properties (Nurgali et al., 2004; Bornstein, 2006; Mao et al., 2006; Qu et al., 2008). However, while calbindin is considered a useful marker of AH/Dogiel type II neurons in guinea pig ileum, CGRP or neurofilament M are more specific markers of these neurons in mouse ileum (Qu et al., 2008) (Tables 1.1 and 1.2).
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Myenteric ISNs project to neurons within the myenteric ganglia including other ISNs, interneurons, and motor neurons, to submucosal ganglia (Furness et al., 2004a), and to the mucosa (Song et al., 1994). Type II neurons in the mouse colon also project within the myenteric ganglia and through a combination of retrograde tracing and lesion studies, have been shown to directly innervate the mucosa of the colon and ileum (Furness et al., 2004b; Qu et al., 2008). In guinea pig small intestine, myenteric type II neurons were mainly found to either project circumferentially (Bornstein et al., 1991), or had descending projections up to 110 mm anally, while short oral projections were less than 2 mm (Brookes et al., 1995).
In guinea pigs, submucosal ISNs can detect mechanical distortion of the mucosa (Kirchgessner et al., 1992; Pan and Gershon, 2000). Submucosal ISNs project to the mucosa (Furness et al., 2004a), to other submucosal neurons (Evans et al., 1994), and to myenteric ganglia (Song et al., 1998). Similar to myenteric ISNs, projections of submucosal ISNs within the plexus display an oral to anal polarity, although they project considerably shorter distances (up to 3 mm) circumferentially or longitudinally (Moore and Vanner, 1998).
As previously highlighted, immunohistochemical (Mongardi Fantaguzzi et al., 2009) and electrophysiological studies (Wong et al., 2008; Foong et al., 2014) of the murine intestine did not identify submucosal ISNs. Qu et al (2008) reported the presence of neurofilament M-immunoreactive nerve cell bodies in the submucosal plexus in sections of mouse ileum, where neurofilament M was used as a marker of type II neurons. However, in wholemount preparations, it was apparent that only neurofilament M-immunoreactive nerve fibers and no cell bodies were present in submucosal ganglia (Mongardi Fantaguzzi et al., 2009). Taken together, this suggests that reflexes in the murine intestine may run via the myenteric plexus as observed in the guinea pig ileum (Reed and Vanner, 2007). However, despite the apparent lack of submucosal type II/AH neurons, there is a possibility that submucosal neurons with sensory functions are present in mouse. Mechanosensitive myenteric S-type have been described in both the
21 small and large intestines of guinea pig and mouse (Spencer and Smith, 2004; Mazzuoli and Schemann, 2009, 2012; Mazzuoli-Weber and Schemann, 2015).
Interneurons
The myenteric plexus of guinea pig small intestine contains one class of cholinergic ascending interneurons immunoreactive for calretinin and substance P (SP), and three classes of descending interneurons distinguished by immunoreactivity for nitric oxide synthase (NOS)/VIP, somatostatin (SOM), or 5-HT (Furness, 2000). Neurons with similar neurochemical coding have been identified in the mouse ileum (Sang and Young, 1996; Sang et al., 1997; Sang and Young, 1998; Qu et al., 2008) (Table 1.2). Nevertheless, much of the current understanding of their connectivity is based on correlated electrophysiological and immunohistochemical studies on the guinea pig circuitry.
Within the myenteric plexus, each class of interneuron forms connections with like interneurons, extending orally and anally along the intestine (Furness, 2006). Ascending interneurons receive inputs from local sensory neurons and project to excitatory motor neurons (Pompolo and Furness, 1995). NOS/VIP- and SOM-immunoreactive descending interneurons project to and receive inputs from other NOS/VIP- and SOM- immunoreactive interneurons. These descending interneurons also project to inhibitory motor neurons (Bornstein et al., 2004; Neal and Bornstein, 2008). NOS/VIP interneurons, like ascending interneurons, also receive inputs from local sensory neurons (Li and Furness, 2000). Unlike NOS/VIP- and SOM-immunoreactive interneurons, 5-HT descending interneurons do not contact inhibitory motor neurons, and primarily form connections with other 5-HT interneurons (Young and Furness, 1995). These interneurons are the only source of neuronal 5-HT in the ENS (Furness et al., 1995b). A small subset of S neurons exhibit 5-HT3-mediated fast EPSPs, which suggests a role for 5-HT interneurons in descending excitation (Zhou and Galligan, 1999; Gwynne and Bornstein, 2007b). There is further evidence showing that these interneurons contact myenteric AH neurons (Erde et al., 1985) and evoke slow EPSPs in
22 these neurons via 5-HT7 receptors (Monro et al., 2005; Tonini et al., 2005; Gwynne and Bornstein, 2007b).
Each of the three types of descending interneurons also projects to submucosal ganglia (Li et al., 1995; Song et al., 1997; Meedeniya et al., 1998), however ascending interneurons do not (Furness, 2006). Intracellular recordings from guinea pig ileum preparations containing both the submucosal and myenteric plexuses have demonstrated functional connections between the two (Moore and Vanner, 2000). This study identified both fast and slow excitatory synaptic inputs to submucosal S-type neurons anal to the site of stimulation in the myenteric plexus. Slow EPSPs were not seen when the distance between the recording site and point of stimulation exceeded 10 mm, whereas fast EPSPs were detected at distances of up to 25 mm. Interestingly, they reported a lack of IPSPs in the submucosal neurons. There is also evidence for projections running from the submucosal plexus to the myenteric plexus. A small number of submucosal neurons in guinea pig ileum (<1%) have single axons that project to myenteric ganglia, but not to the mucosa or to submucosal arteries, indicating that they are interneurons connecting the two plexuses (Song et al., 1998). An electrophysiological study further showed that focal electrical stimulation of submucosal neurons can evoke fast and slow EPSPs in myenteric S neurons, and slow EPSPs in myenteric AH neurons (Monro et al., 2008). However, relatively few neurons were reported to display synaptic potentials (7/29 neurons studied) which suggests functional connections from the submucosal plexus to the myenteric plexus are sparse.
The evidence for the presence of interneurons in the submucosal plexus is less clear. A study by Evans et al. (1994) in guinea pig small intestine showed that NPY- and VIP- secretomotor neurons provide varicose nerve terminals to other submucosal neurons and therefore, may possibly function as interneurons. Reed and Vanner (2001) have also showed that cholinergic and a small proportion of VIP-neurons can evoke fast EPSPs in an adjacent VIP-neuron in submucosal plexus preparations. However, a separate study reported that only ISNs make obvious synaptic contacts with other neurons within the
23 submucosal plexus (Furness et al., 2003). The discrepancies between these are unclear. ISNs under certain circumstances can also evoke nicotinic fast EPSPs in S-neurons (Pan and Gershon, 2000), which suggests that in some cases secretomotor reflexes may circumvent the intermediate neuron and ISNs can directly signal to secretomotor neurons. In this sense, ISNs may also act as interneurons.
Whether sensory neurons or interneurons are present in the murine submucosal plexus remains elusive. Nonetheless, electrophysiological studies have reported the occurrence of spontaneous fast EPSPs and fast EPSPs which were not tightly stimulus locked in submucosal plexus preparations from mouse colon, implying that functional contacts between submucosal neurons may be present and allow for neural integration (Wong et al., 2008; Foong et al., 2014). However, this is an area that requires further investigation.
Secretomotor neurons
Secretomotor neurons are the final motor neurons which mediate intestinal secretion and vasodilation, and they reside in the submucosal plexus. However, these neurons have only been completely characterized in the guinea pig small intestine (Furness, 2006). These include a major class of VIP-containing secretomotor/vasodilator neurons and one class of calretinin-containing cholinergic secretomotor/vasodilator neurons, which innervate both the mucosal epithelium and submucosal arterioles, as well as a separate class of cholinergic NPY-containing secretomotor neurons which only supply the mucosa (Furness et al., 2003) (Table 1.1). By comparison, in the mouse small intestine, secretomotor and vasodilator neurons form distinctly separate classes, that is, specific subsets of submucosal neurons project to either the mucosa or blood vessels, but not to both (Mongardi Fantaguzzi et al., 2009). This suggests that perhaps some secretomotor neurons and vasodilator neurons receive common diverging inputs in order to synchronously regulate localized blood flow and intestinal secretion. In addition, the specific neurochemical coding of submucosal neurons also differs with that of the guinea pig, where NPY and calretinin both colocalize with VIP in the mouse
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(Mongardi Fantaguzzi et al., 2009). Finally, there is a small population (~8%) of submucosal neurons that are neither ChAT- or VIP-immunoreactive, and are yet to be defined neurochemically.
Despite these differences, the two major populations of secretomotor neurons: cholinergic (ChAT-immunoreactive) and non-cholinergic (VIP-immunoreactive), are generally well-conserved across species (Hubel, 1985; Keast et al., 1985a, b; Porter et al., 1996; Porter et al., 1999; Krueger et al., 2016). The division of cholinergic and non- cholinergic secretomotor neurons is consistently observed in small animals, including guinea pigs and mice (Furness, 2000; Mongardi Fantaguzzi et al., 2009). However, this segregation is less clear in humans, where ChAT co-localizes with VIP in some submucosal neurons (Schneider et al., 2001; Anlauf et al., 2003; Krueger et al., 2016).
VIP-secretomotor neurons have been the focus of the many secretion studies and are considered to have a more prominent role in regulating secretion, over cholinergic secretomotor neurons. There are several reasons for this. Firstly, they are the major site of integration of a number of intrinsic and extrinsic neuronal inputs, at least in the guinea pig (Bornstein et al., 2010). This includes inhibitory synapses from sympathetic nerves and inhibitory myenteric interneurons, whereas cholinergic secretomotor neurons do not appear to have inhibitory inputs (Bornstein et al., 1986, 1988; Evans et al., 1994). Secondly, VIP-secretomotor neurons have been implicated in the hypersecretory response evoked by cholera-toxin (CT) in rat small intestine studies (Mourad and Nassar, 2000; Banks et al., 2005; Kordasti et al., 2006), and this will be further discussed later.
In contrast with the lab animal studies, a comprehensive study by Krueger and colleagues (2016) using over 2200 surgical samples showed that cholinergic secretomotor neurons are the key mediators of electrically-stimulated intestinal secretion in human small intestine. However, human studies of CT-evoked
25 hypersecretion implicate VIP in pathophysiological secretion (Bloom et al., 1976; Banks et al., 2005). A further complication is the co-expression of ChAT and VIP in some submucosal neurons in the human small intestine (Schneider et al., 2001; Anlauf et al., 2003; Krueger et al., 2016). Thus, the relative contribution of ACh and VIP to the control of intestinal secretion remains unclear.
Motor neurons
The myenteric plexus contains both excitatory and inhibitory motor neurons, which supply the longitudinal and circular muscle. Excitatory motor neurons use ACh and tachykinins as cotransmitters, while inhibitory motor neurons use NO, with the cotransmitters ATP, VIP, and pituitary adenylyl cyclase activating peptide (PACAP) (Bornstein et al., 2004). Both patterns of expression of excitatory and inhibitory transmitters appear to be well conserved between the guinea pig and mouse small intestine (Qu et al., 2008) (Table 1.2). Both excitatory and inhibitory motor neurons receive excitatory inputs from interneurons, as well as direct inputs from calbindin- immunoreactive ISNs, indicating that there are monosynaptic ISN to motor neurons reflex pathways.
4. Enteric Reflex Circuits
Mechanical or chemical stimulation of the mucosa and distension elicit polarized reflexes. The most basic elements of a reflex circuit comprise the afferent limb – a sensory neuron, and the efferent limb – a motor neuron, which are connected either directly or via an interneuron.
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4.1 Secretomotor reflexes
Distension (Frieling et al., 1992; Weber et al., 2001), mechanical distortion of the mucosa (Sidhu and Cooke, 1995; Kellum et al., 1999; Christofi et al., 2004; Cooke et al., 2004), chemicals and nutrient applied to the mucosa (See and Bass, 1993; Hansen and Witte, 2008), and various noxious agents (Lundgren and Jodal, 1997; Lundgren et al., 2000) all trigger secretomotor reflexes. Different forms of stimulation may activate different sensory elements and hence it follows that different pathways are stimulated accordingly (Grider and Jin, 1994; Cooke et al., 1997b). Nevertheless, the key components of the nerve circuitry involved are contained intrinsically within the walls of the gastrointestinal tract, as secretory reflexes can still be evoked after extrinsic denervation (Caren et al., 1974; Lundgren et al., 1989). Further, the efferent neurons are largely contained within the submucosal plexus, since neurogenic secretory responses can be recorded in mucosa and submucosal plexus preparations from which the myenteric plexus has been removed (Carey et al., 1985; Carey and Cooke, 1986; Johnson et al., 1994).
The submucosal plexus may also contain both the efferent and afferent elements of the secretomotor circuit, depending on the region and species. For instance, complete secretomotor circuits have been described in the submucosal plexus of guinea pig and rat colon (Cooke et al., 1997b; Christofi et al., 2004; Cooke et al., 2004). These in vitro secretion studies using preparations lacking the myenteric plexus showed that mucosal stroking excites submucosal ISNs and stimulates the release of endogenous purines to activate postsynaptic P2Y receptors on cholinergic and VIPergic secretomotor neurons. Subsequent ACh- or VIP-release activates mucosal muscarinic M3 receptors or VPAC1 receptors respectively, to trigger a secretory response.
In addition to this conventional circuit, the activation of axon reflexes may cause SP release from ISNs with collaterals in the mucosa to directly stimulate NK1 receptors and evoke a secretory response (Cooke et al., 1997c; Cooke et al., 1997b). Furthermore,
27 there is an extrinsic innervation of the mucosa and stimulation of these can also elicit a secretory response (Vanner and Macnaughton, 2004).
Retrograde tracing of submucosal neurons innervating the mucosa of guinea pig colon showed that the majority of their projections were less than 5 mm longitudinally or circumferentially (Neunlist et al., 1998). This is consistent with the proposal that intrinsic secretomotor reflexes are predominantly localised and confined to the site of the stimulus, i.e. are short reflexes (Hubel et al., 1991; Furness, 2006). However, there is also evidence for long secretomotor reflex pathways that run through the myenteric plexus in the guinea pig small intestine (Reed and Vanner, 2007). This study showed that applying an electrical stimulus to intact mucosa evoked fast EPSPs in submucosal neurons located 0.7 – 1 cm anally from the point of stimulation. The number of neurons displaying fast EPSPs was reduced by severing the connections between the mucosa/submucosa and the underlying layers. Furthermore, fast EPSPs were absent after a lesion in the myenteric plexus was made between the stimulus site and the recording site, but were still observed after lesioning the submucosal plexus. Thus these experiments show that the reflex examined likely involves myenteric sensory afferents innervating the mucosa, and that the circuit activated runs through the myenteric plexus. While this was not examined, it is likely that these long reflexes are polysynaptic (Reed and Vanner, 2007) and may involve any of the 3 classes of descending interneurons described earlier (Li et al., 1995; Song et al., 1997; Meedeniya et al., 1998; Moore and Vanner, 2000). Given that the mouse submucosal plexus appears to lack ISNs as discussed above, it is conceivable that long secretomotor reflexes also occur in the murine ENS.
4.2 Recurrent networks of submucosal neurons
In the guinea pig small intestine, submucosal ISNs identified by SP-immunoreactivity typically project in a circumferential direction within the plexus and for distances of up to 3 mm (Song et al., 1992). Evans et al. (1994) and Reed and Vanner (2001) also showed that submucosal ISNs provide dense networks of varicosities within the same
28 ganglia and to adjacent ganglia. This arrangement indicates that submucosal ISNs may form excitatory self-reinforcing networks by signaling via slow EPSPs (Chambers et al., 2005) as seen in the myenteric plexus with myenteric ISNs (Pompolo and Furness, 1988; Kunze et al., 1993; Thomas et al., 2000). These recurrent networks likely provide a significant source of excitation for secretomotor neurons, even in the absence of a sensory stimulus (Chambers et al., 2005). As basal secretion is at least partially neurogenic (Carey and Cooke, 1989b), and VIP neurons receive diverging input from local submucosal ISNs (Reed and Vanner, 2001), these networks may contribute to the tonic drive of VIP secretomotor neurons (Chambers et al., 2005).
It has also been proposed that submucosal VIP neurons may form recurrent networks (Chambers et al., 2005). They have processes that travel up to 3.5mm roughly circumferentially (Song et al., 1992) and can traverse up to 7 submucosal ganglia (Reed and Vanner, 2001). These neurons provide varicosities along the length of their axons which may contact cell bodies within a ganglion as they pass through and have occasional axon collaterals (Evans et al., 1994, Reed and Vanner, 2001). Thus, it is feasible that submucosal VIP neurons innervate other submucosal neurons via these varicose projections and may act as interneurons. However, this proposal has been challenged by reports that VIP-immunoreactive submucosal neurons do not form varicosities around other submucosal neurons (Costa and Furness, 1983; Furness et al., 2003). On the other hand, VIP has also been shown to depolarize submucosal neurons in guinea pig caecum (Mihara et al., 1985). Dual impalement intracellular recordings from pairs of submucosal VIP neurons showed that stimulating one VIP neuron evoked a slow depolarizing response in the neighboring VIP neuron (Reed and Vanner, 2001). Hence, this electrophysiological evidence supports the argument for the presence of recurrent VIP networks in the submucosal plexus. Computer modeling studies suggest that these excitatory self-reinforcing VIP-networks have the potential to become hyperexcitable and may have a role in the neural mechanism underlying pathological hypersecretion (Chambers et al., 2005).
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4.3 Motility reflexes
Many of the stimuli which trigger secretomotor reflexes, such as mechanical or chemical stimulation of the mucosa, as well as distension, also elicit motility reflexes (Smith and Furness, 1988; Smith et al., 1990, 1991). Motility reflexes can be evoked in guinea pig small intestine following chronic extrinsic denervation, indicating that they can be generated by intrinsic mechanisms (Furness et al., 1995a). Local motility reflexes underlie complex motor patterns such as segmentation, which is associated with the mixing motions for nutrient absorption (Cannon, 1912; Gwynne et al., 2004; Gwynne and Bornstein, 2007a), and peristalsis for the propulsion of luminal content along the gastrointestinal tract (Bayliss and Starling, 1899; Bornstein, 1994; Furness et al., 1995a). Segmentation and peristalsis are observed in the fed state and are patterns that are also observed in isolated intestinal segments (Mall, 1896; Bornstein, 1994; Gwynne and Bornstein, 2007a). In addition, migrating myoelectric complexes (MMCs) are observed in the small intestine and occur in the fasted state in humans and other species that feed intermittently (Telford and Sarna, 1991; Kunze and Furness, 1999; Furness, 2006; Deloose et al., 2012). MMCs can also occur in the fed state, though less frequently, in species which feed constantly such as guinea pigs (Galligan et al., 1985). MMCs comprise of a set of phasic motor patterns where there are alternating periods of quiescence (Phase I) and strong propulsive contractions that traverse the length of the small intestine in the anal direction (Phase III). These are separated by sporadic motor activity (Phase II and IV) (Bornstein et al., 2002). MMCs are not readily observed in vitro (Bornstein, 1994; Thomas et al., 2004) and have mainly been characterized based on in vivo studies (Sarna et al., 1981; Galligan et al., 1985; Galligan et al., 1986, 1989). Thus, while these studies show that intrinsic neural circuitry is important in mediating MMCs, these pathways involved are not well defined.
Stretch-sensitive myenteric ISNs (Kunze et al., 1998), myenteric interneurons and motor neurons are sufficient to generate motility reflexes in response to distension, and removing the submucosal plexus does not interrupt the propagation of this reflex (Smith et al., 1990; Bornstein, 1994). The basis of most propulsive motility patterns are peristaltic reflexes, which involve an ascending excitatory pathway and a descending
30 inhibitory pathway, and evokes circular muscle contraction oral to the stimulus and relaxation immediately anal to the stimulus (Mall, 1896; Bayliss and Starling, 1899). With correlated electrophysiological, morphological and immunohistochemical analyses, various elements of the circuit underlying these simple motor reflexes were deduced (Bornstein et al., 1991; Bornstein et al., 2002). The model proposed involves the activation of ascending interneurons which synapse excitatory motor neurons innervating the longitudinal and circular muscle to evoke a contraction immediately oral to the stimulus site. There is also a concurrent activation of descending interneurons contacting motor neurons to evoke circular muscle relaxation and longitudinal muscle contraction anal to the stimulus (Bornstein et al., 1991; Bornstein et al., 2004).
4.4 Co-ordination of secretomotor and motility reflexes
It is clear that similar types of stimuli can trigger motility and secretomotor reflexes, and the co-ordination of intestinal secretion and motor activity is thought to be important for normal gastrointestinal function (Furness, 2006). Furthermore, secretomotor and motility reflexes may involve common neurotransmitters such as ACh (Cassuto et al., 1982a; Galligan et al., 1986; Spencer et al., 2000; Bian et al., 2003; Vanner and Macnaughton, 2004; Reed and Vanner, 2007), 5-HT (Keast et al., 1985b; Johnson et al., 1994; Monro et al., 2002; Monro et al., 2005; Tonini et al., 2005; Gwynne and Bornstein, 2007b; Foong et al., 2010a), and ATP (Cooke et al., 2004; Thornton et al., 2013) to name a few. VIP is another putative transmitter of interest given its expression in both secretomotor neurons and myenteric motor neurons (Tables 1.1 and 1.2).
Despite this, there are only limited studies that have investigated the coupling of intestinal secretion and motility. Evidence for synchronized changes in intestinal secretion and motility is derived from a number of studies in the human, canine, guinea pig, and rat small intestine, where a temporal correlation between spontaneous motor activity and changes in transmural potential difference (a measure of electrogenic ion secretion) was observed (Read et al., 1977; Greenwood et al., 1986; Greenwood and
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Davison, 1987; See et al., 1990). It is thought that in the fed state, synchronized changes in secretion and motility are important for optimal digestion and absorption of nutrients (Greenwood and Davison, 1987). It has been further speculated that intestinal secretions may have a protective role against mechanical stresses imposed by the luminal contents on the mucosal epithelium (Barrett and Keely, 2000; Murek et al., 2010). Changes in transmural potential difference have also been associated with phase III of the MMC (Read, 1980; Mellander et al., 2000; Mellander et al., 2001). In the fasted state, intestinal secretion may be necessary for lubricating the intestinal lumen to assist in the transit of luminal debris along the intestinal tract (Greenwood and Davison, 1987). In addition, pathological hypersecretion such as that evoked by cholera toxin (CT), may also be linked with altered motility (Kordasti et al., 2006), and together these responses may act as a protective mechanism to eliminate the noxious agent.
It has been suggested that long secretomotor reflexes running through the myenteric plexus may be involved in integrating secretion and motility in the guinea pig ileum (Vanner and Macnaughton, 2004; Reed and Vanner, 2007). Yet a study examining the coupling of circular muscle contraction and electrogenic ion secretion in rat jejunum showed that this process was unaffected by chronically ablating the myenteric plexus (See et al., 1990). It was concluded that integration of the motility and secretory reflex occurs in the submucosal plexus. However, it has also been shown using the same animal model that a degree of neuroplasticity is apparent following myenteric denervation (Luck et al., 1993). While the submucosal plexus does provide some innervation of the circular muscle in rat jejunum, these fibers were seen to proliferate in the absence of the myenteric plexus. Thus, the possibility that under normal conditions, the myenteric plexus is involved in linking secretion and motor activity in rat jejunum cannot be ruled out.
Progress in this area had largely been hindered by the technical limitations associated with simultaneously measuring both the rapid changes in motor activity and secretion. Moreover, the methods used were lacking in the temporal and spatial resolution
32 necessary to identify the specific neural elements involved. However, with recent advances in optical imaging techniques and development of transgenic reporter mice, there is potential to better examine integrated network activity and the corresponding functional output associated with complex gut behaviours (Vanden Berghe et al., 2001; Zariwala et al., 2012; Boesmans et al., 2015a; Hennig et al., 2015; Mutoh et al., 2015). For instance, a relatively recent study using calcium imaging with the fluorescent indicator Fluo-4, has shown a degree of synchrony in activity of myenteric and submucosal neurons during colonic migrating motor complexes (CMMCs) in mice (Okamoto et al., 2012). Interestingly, activity within the enteric glial network also appears to be synchronized with nerve activity during CMMCs (Broadhead et al., 2012). This raises the possibility that glia might also interact with neurons and participate in coordinating enteric reflexes, thus adding a further level of complexity to the system. Although detailed analysis of the enteric circuitry remains a challenge with studies of this nature as movement artefacts are still a limiting factor (Boesmans et al., 2015a), new analytical methods are emerging to address these issues (Hennig et al., 2015).
5. Enteric Glia
Depending on the species, enteric glia can be found in equivalent or greater numbers than neurons (Gabella and Trigg, 1984; Hoff et al., 2008). By comparison with enteric neurons which been extensively studied, the role of enteric glia is far less well understood. This is largely attributed to the limited methods that were available to examine glial activity, and for a significant period of time glia were dismissed as simply being support cells. However, with the imaging technologies and techniques now available (Boesmans et al., 2013a), it is becoming apparent that enteric glia are actively involved in the control of gut function (Neunlist et al., 2014). Emerging data suggest that enteric glial cells also have important roles in regulating physiological colonic motility (Nasser et al., 2006b; Broadhead et al., 2012; Gulbransen and Sharkey, 2012; McClain et al., 2015) and in pathophysiology such as colitis (Nasser et al., 2007; MacEachern et al., 2015; Brown et al., 2016; Ochoa-Cortes et al., 2016). Whether
33 enteric glia are also involved in intestinal secretion is unclear. Furthermore, much of the current literature detailing the role of enteric glia mainly focuses on the myenteric plexus of mouse intestine, and few studies have examined enteric glia in the submucosal plexus.
5.1 Enteric glial subtypes
Enteric glial cells have been classified into subtypes (I – IV) based on their location within the layers of the gastrointestinal tract and distinct morphology (Boesmans et al. 2014) (Figure 1.6). Type I glial cells, which have been likened to astrocytes, are intraganglionic and possess numerous highly branched processes that project out in multiple directions from a central nucleus (Hanani and Reichenbach, 1994). Type II glial cells are interganglionic and appear elongated with processes running parallel along with the internodal strands. Type III glial cells are those found within the level of the plexuses or lamina propria, are extraganglionic, and tend to have 4 major processes with secondary branching (Boesmans et al., 2014). Type IV glial cells display a bipolar morphology and are intramuscular (Gulbransen and Sharkey, 2012; Vanderwinden et al., 2003; Boesmans et al., 2014).
As with enteric neurons, enteric glia show differential expression of various chemical markers. For instance, the majority of type I glial cells in the myenteric plexus of mouse colon were found to co-express glial fibrillary acidic protein (GFAP) and S100 calcium- binding protein β (S100β), while the remainder of type I cells expressed either GFAP or S100β (Boesmans et al., 2015b). The co-expression of GFAP or S100β and the transcription factor Sox10 were similarly examined, but ultimately, the various patterns of GFAP, S100β, and/or Sox10 were not unique to any one morphological glial subtype (Boesmans et al., 2015b). Moreover, whether these different glial subtypes are functionally distinct remains obscure.
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Figure 1.6. Morphological subtypes of enteric glia.
A. Individual green fluorescent protein (GFP)-expressing enteric glia (green) situated within a ganglion, outside ganglia, or within the interconnecting fibre tracts of the myenteric plexus (Tuj1, red). B – E. Different enteric glial subtypes (type I–IV) expressing MADM (mosaic analysis with double markers)-induced GFP (green) with the glial marker S100β (gray). (F – I) Deconvolved maximum projections (top) and 3-D reconstructions (bottom) of type I (F), II (G), III (H), and IV (I) enteric glial cells identified in Sox10Cre|MADM mice. Figure taken from Boesmans et al. (2015b).
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5.2 Enteric glia are involved in regulating intestinal motility and barrier function
Insights on the functional roles of enteric glia have been gleaned from studies examining the effects of perturbing glial activity using gliotoxins (Nasser et al., 2006b; MacEachern et al., 2015) or using genetic approaches (Aube et al., 2006; Savidge et al., 2007; McClain et al., 2014). The most prominent effects observed were typically alterations in motility measured as slowed transit in the small and/or large intestines (Aube et al., 2006; Nasser et al., 2006b; McClain et al., 2014). Calcium imaging studies have also demonstrated that glial activation can be driven by intrinsic excitatory motor pathways associated with CMMCs (Broadhead et al., 2012). Thus, there is increasing evidence showing that enteric glia may be actively involved in the physiological regulation of intestinal motility (Sharkey, 2016).
Disrupting glial activity also alters intestinal barrier function and increases epithelial permeability (Aube et al., 2006; Savidge et al., 2007; Neunlist et al., 2013; MacEachern et al., 2015). However, there is no clear evidence of changes to intestinal secretion (Nasser et al., 2006b; MacEachern et al., 2015). It has been reported that glial-specific connexin-43 knockout mice, in which the propagation of glial Ca2+ signals is disrupted, produced faecal pellets with higher water content (McClain et al., 2014). While this suggests that secretomotor function may have been affected, electrogenic ion secretion was not specifically examined. Mice administered a CNS gliotoxin were found to develop diarrhoea and this was associated with morphological abnormalities in myenteric glia in the colon (Aikawa and Suzuki, 1985). However, intestinal function was not directly assessed and whether similar effects were observed in the submucosal plexus or in other intestinal regions is unclear. Hence, while enteric glia may be involved in the regulation of intestinal motility and barrier function, their role in the control of intestinal secretion remains to be elucidated.
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5.3 The role of enteric glia in gastrointestinal diseases
Enteric glia are implicated in various pathological states and it has been proposed that a “reactive enteric glial cell phenotype” may contribute to gastrointestinal diseases, such as irritable bowel syndrome and inflammatory bowel disease (Cabarrocas et al., 2003; Ochoa-Cortes et al., 2016). This “reactive phenotype” is associated with an increased production of pro-inflammatory mediators such as cytokines IL1b, IL-6, and tumor necrosis factor-α (Cabarrocas et al., 2003; Ochoa-Cortes et al., 2016), and is typically marked by the upregulation of GFAP (Cabarrocas et al., 2003; Neunlist et al., 2013). For instance, in ulcerative colitis, GFAP upregulation is linked to increased S100β production and release, which in turn increases NO production and may have detrimental effects (Kolios et al., 2004; Cirillo et al., 2009; Cirillo et al., 2011; Neunlist et al., 2013). A recent study has also suggested that enteric glial activation may contribute to neuronal cell death in colitis (Brown et al., 2016). However, whether changes in enteric glial phenotype occur as a consequence of disease, or if enteric glia are actively involved in initiating disease and/or its progression, remain to be determined.
5.4 Protective effects of enteric glia and neurogenesis
Enteric glia may have a detrimental role under some conditions (Gulbransen et al., 2012; Brown et al., 2016). However, they are also capable of exerting protective effects through the release of a range of factors such as glial-derived S-nitrosaglutathione (GSNO) – which enhances epithelial barrier function (Savidge et al., 2007) – and glial- derived neurotrophic factor (GDNF) (Anitha et al., 2006; Zhang et al., 2010; Baudry et al., 2012; Neunlist et al., 2013). Further, mouse enteric glial progenitors have the potential to undergo neurogenesis in vivo in response to ENS injury (Laranjeira et al., 2011). This raises the possibility of using glia in diseases associated with neurodegeneration or aganglionosis (Ochoa-Cortes et al., 2016).
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5.5 Enteric neuron-glia signaling
The general consensus is that enteric glial cells are not electrically excitable (Hanani et al., 2000); although, glial cells in the central nervous system possess a range of ion channels similar to neurons (Verkhratsky and Steinhäuser, 2000). Glial activity is reflected by changes in intracellular [Ca2+], and information is integrated and transmitted via various Ca2+ signals (Ochoa-Cortes et al., 2016). However, the mechanisms for the bi-directional communication between enteric neurons and glia are not well understood. Ultrastructural studies demonstrating that nerve endings terminating on enteric glia have presynaptic specializations (Gabella, 1981) provided the earliest anatomical evidence for connections between enteric neurons and glia. The nerve axons at these neuron-glia junctions were shown to predominantly contain small agranular vesicles and some larger granular vesicles. In addition, synaptic release sites have been identified on enteric glia by a combination of immunohistochemical analysis and imaging using styryl dye (Vanden Berghe and Klingauf, 2007). Further, close apposition of serotonergic varicosities to enteric glia have been identified (Okamoto et al., 2014). There is also some evidence suggesting that enteric glia are innervated by sympathetic nerves (Gulbransen et al., 2010) and intrinsic serotonergic inputs (Okamoto et al., 2014).
Enteric glia express a variety of receptors for neurotransmitters (Nasser et al., 2006a; Nasser et al., 2007) and many of these are purinergic (Gomes et al., 2009; Gulbransen and Sharkey, 2009) thus implicating purinergic signaling in neuron-glia communication. In addition to ATP, enteric glia can respond to acetylcholine and serotonin, which are major enteric neurotransmitters (Boesmans et al., 2013b). Enteric glia in turn have the potential to release neuroactive “gliotransmitters” such as ATP (Zhang et al., 2003) to modulate neural activity (Ochoa-Cortes et al., 2016), but whether “gliotransmission” occurs within the ENS remains elusive. Nevertheless, a recent study by McClain et al. (2015) suggests that selective activation of enteric glia can evoke neurogenic muscle contractions. Common transmitters and receptors that may be utilized by enteric glia and neurons will be further discussed in the following section.
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6. Neural and Glial Transmitters and Receptors
Activity of enteric neurons and intestinal function can be regulated by a range of transmitters and receptors. Many of these transmitters have also been identified in enteric glia and their release from glia have the potential to modulate neural activity. A discussion of this is incorporated in the following sections. At least four distinct post- synaptic events are observed at enteric neuro-neuronal synapses and can be broadly divided into: fast EPSPs, slow EPSPs, intermediate EPSPs and slow inhibitory postsynaptic potentials (IPSPs) (Bornstein and Furness, 1988; Gwynne and Bornstein, 2007b; Foong et al., 2010b). The transmitters involved in mediating these synaptic potentials, will be discussed. The majority of this work was conducted in guinea pig intestine, thus the following studies described largely refer to findings based on the guinea pig circuitry, unless stated otherwise.
6.1 Acetylcholine, nicotinic receptors (nAChRs), and muscarinic receptors (mAChRs)
Cholinergic transmission is the major form of excitatory neurotransmission within the ENS (Galligan and North, 2004). This is primarily mediated through nicotinic ACh receptors (nAChRs), which are ligand-gated ion channels (Kirchgessner and Liu, 1998; Obaid et al., 2005). This form of communication occurs between most enteric neurons (Surprenant, 1984; Bornstein et al., 1986; Bornstein et al., 1987; Kunze and Furness, 1999; Foong et al., 2015). Activation of nAChRs evokes fast EPSPs in submucosal and myenteric neurons which can be blocked by nicotinic antagonists such as tubocurarine, hexamethonium, or mecamylamine (Nishi and North, 1973; Hirst et al., 1974; Hirst and McKirdy, 1975; Surprenant, 1984; Pan and Gershon, 2000). Submucosal neurons receive fast excitatory inputs mainly from myenteric neurons (Moore and Vanner, 1998, 2000), though there is some contribution from cholinergic submucosal neurons (Bornstein et al., 1987; Reed and Vanner, 2001). Further, secretomotor reflexes, excitatory ascending reflex contractions triggered by mucosal stimulation, peristalsis and MMCs have all been shown to involve nicotinic cholinergic neurotransmission in
39 various species including guinea pig, cat, rat, and mouse (Cassuto et al., 1982a; Galligan et al., 1986; Spencer et al., 2000; Bian et al., 2003; Vanner and Macnaughton, 2004; Reed and Vanner, 2007).
While fast EPSPs in submucosal and myenteric neurons commonly involve nAChRs (Galligan and Bertrand, 1994; Moore and Vanner, 1998; Galligan et al., 2000), they are not exclusively mediated by cholinergic transmission. Purinergic P2X receptors and 5- HT3 receptors also have important roles (LePard et al., 1997; Johnson et al., 1999; Galligan et al., 2000; Galligan and North, 2004; Monro et al., 2004; Gwynne and Bornstein, 2007b). As in the guinea pig, nAChR-mediated fast EPSPs have been described in submucosal and myenteric neurons of the mouse small and/or large intestine (Bian et al., 2003; Nurgali et al., 2004; Wong et al., 2008; Foong et al., 2015). There is further evidence that ATP also acts at P2X receptors as a fast synaptic transmitter in the myenteric plexus of mouse intestine (Bian et al., 2003; Ren et al., 2003; Nurgali et al., 2004).
The other major subtype of ACh receptors is muscarinic receptors (mAChRs), which are G-protein coupled receptors (Caulfield and Birdsall, 1998). The localization and function of mAChRs on smooth muscle are well established and are important in directly mediating ACh-evoked muscle contraction. Presynaptic mAChRs on the nerve fibers innervating muscle are also involved in modulating ACh-release (Harrington et al., 2010). mAChRs mediate slow EPSPs and may be involved in transmission between myenteric sensory neurons, sensory neurons to ascending interneurons and descending interneurons (Harrington et al., 2010).
In addition to its role in enteric neurotransmission, ACh may also be involved in neuron to glia signaling, as it has been shown to directly evoke Ca2+ responses in rat and human enteric glia, albeit in culture (Boesmans et al., 2013b). The response was found to be
40 mediated by mAChRs, rather than nAChRs. How this may contribute to signaling within the intact circuitry remains to be determined.
6.2 Noradrenaline and α2-adrenoceptors
Almost all VIP submucosal neurons, but few cholinergic neurons, display inhibitory synaptic inputs in response to electrical stimulation (Bornstein et al., 1986, 1988; Evans et al., 1994; Hu et al., 2003; Foong et al., 2010b). Noradrenaline is the primary transmitter of post-ganglionic sympathetic neurons, and mediates slow IPSPs in VIP- secretomotor neurons via α2-adrenoceptors (Mihara et al., 1987; Bornstein et al., 1988; Shen and Surprenant, 1990; Foong et al., 2010b). This was first demonstrated by North and Surprenant (1985) who showed that 6-hydroxydopamine, toxic to adrenergic neurons, abolished the IPSP. The hyperpolarizing potentials evoked by local application of noradrenaline mimicked the IPSP and this was blocked by various α2-adrenoceptor antagonists (North and Surprenant, 1985). This form of inhibition is important in the tonic control of intestinal secretion (Hubel, 1976; Furness, 2006).
Expression of α2-adrenoceptors has also been observed in enteric glia in the myenteric plexus of rat and mouse, but not guinea pig, small intestine, as well as in the submucosal plexus of the mouse, though less consistently (Nasser et al., 2006a). It is unclear whether these have any physiological roles, but raises the possibility that glial activity may also be under sympathetic control.
6.3 Somatostatin (SOM) and SST1 and SST2 receptors
In addition to adrenergic IPSPs, VIP-submucosal neurons receive intrinsic inhibitory inputs since IPSPs can be induced by transmural stimulation after extrinsic denervation (Mihara et al., 1987; Bornstein et al., 1988; Shen and Surprenant, 1993). Somatostatin (SOM) was first proposed to be the transmitter underlying this non-adrenergic IPSP as it was blocked by SOM-desensitization (Shen and Surprenant, 1993). More recent studies
41 demonstrate that SOM mediates part of the non-adrenergic IPSPs via STT1 and STT2 receptors (Foong et al., 2010a; Foong et al., 2010b). While sympathetic fibers use SOM as a co-transmitter (Costa and Furness, 1984), there are also intrinsic sources arising from descending interneurons in the myenteric plexus, as well as cholinergic NPY- secretomotor neurons (Furness, 2000). However, it is most likely that VIP-secretomotor neurons receive this form of inhibitory input from descending interneurons as they are abolished with extrinsic and myenteric denervation (Bornstein et al., 1988).
SOM has been shown to have anti-secretory actions by modulating neurogenic secretion via SST1 and SST2 receptors (Foong et al., 2010a). Veratridine (activates voltage- dependent Na+ channels) evoked a secretory response that was largely suppressed by
SOM, and this effect was abolished by a SST1 antagonist and inhibited by a SST2 antagonist (Foong et al., 2010a). Further, SOM alone reduced baseline secretion, an effect inhibited by tetrodotoxin (TTX; Na+ channel blocker). SOM also selectively inhibited the non-cholinergic component of the DMPP-evoked secretory response, indicating that SOM primarily acts to reduce the excitability of VIP secretomotor neurons (Foong et al., 2010a).
6.4 Tachykinins and neurokinin (NK) receptors
Neurokinin A and substance P (SP) are tachykinins contained within enteric neurons and extrinsic primary afferent neurons (Costa et al., 1980; Holzer and Holzer-Petsche,
1997b). In the myenteric plexus, NK1 and NK3 receptors are largely involved in mediating slow EPSPs in AH neurons (Alex et al., 2001; Johnson and Bornstein, 2004;
Gwynne and Bornstein, 2009). Tachykinins acting on NK3 receptors may also be involved in transmission between myenteric sensory neurons and ascending interneurons, and ascending interneurons to circular muscle excitatory motor neurons (Johnson et al., 1998; Gwynne and Bornstein, 2007b).
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In the submucosal plexus, cholinergic NPY secretomotor neurons express NK1 and NK3 receptors, and AH neurons express NK1 receptors (Portbury et al., 1996; Lomax et al., 1998; Jenkinson et al., 1999). SP depolarizes submucosal neurons (Surprenant et al., 1987). Further, SP has been shown to evoke secretory responses in mucosa-submucosa preparations from guinea pig small intestine by activating a neural component that is partly sensitive to muscarinic blockade, as well as by a direct action on the mucosa (Keast et al., 1985c; Perdue et al., 1987). Findings from these studies suggest that SP stimulates a secretory response via both cholinergic and non-cholinergic secretomotor neurons. Similar findings were reported using SP and a NK1 agonist in preparations from mouse small intestine (Wang et al., 1995). By contrast, Reddix and Cooke (1992) found that both SP and a NK1 agonist evoked a secretory response in guinea pig small intestine that was unaffected by nicotinic and muscarinic antagonists, suggesting that the response was predominantly mediated via non-cholinergic VIP secretomotor neurons. Thus, the role of cholinergic secretomotor neurons remains elusive. The involvement of NK3 receptors in the small intestine is also unclear, as NK3 agonist was found to have minimal secretory effects (Reddix and Cooke, 1992; Holzer and Holzer-
Petsche, 1997a). However, NK3 receptors have been implicated in the hypersecretory response evoked by CT (Gwynne et al., 2009).
6.5 Serotonin (5-HT) and 5-HT receptors
In addition to EC cells, 5-HT is also contained within a subset of myenteric descending interneurons. The involvement of 5-HT in synaptic transmission and the receptor subtypes expressed by submucosal and myenteric neurons are not well understood. Electrophysiological studies have demonstrated that 5-HT depolarizes all submucosal neurons (Surprenant and Crist, 1988) and some myenteric neurons (Wood and Mayer,
1979; Johnson et al., 1980), and elicits fast EPSPs via ionotropic 5-HT3 receptors (Mawe et al., 1986; Vanner and Surprenant, 1990). By comparison, relatively few neurons exhibit 5-HT3-mediated fast EPSPs evoked by electrical stimulation (Zhou and Galligan, 1999; Monro et al., 2004). 5-HT may also evoke slow EPSPs in myenteric AH neurons (Wood and Mayer, 1979; Johnson et al., 1980; Mawe et al., 1986; Monro et al., 2005). However, the receptor involved remains elusive. There is pharmacological
43 evidence implicating 5-HT1P receptors (Mawe et al., 1986), but 5-HT1P receptor expression is not well defined as the receptor is yet to be cloned. By contrast, 5-HT7 receptors have been characterized and are expressed on myenteric AH neurons (Hedlund and Sutcliffe, 2004; Tonini et al., 2005). Further, using a selective antagonist,
5-HT7 receptors were shown to mediate a slow EPSP previously attributed to 5-HT1P receptors (Monro et al., 2005). Some suggested possibilities are that the antagonist used may target both 5-HT1P and 5-HT7 receptors, or that they may even be the same receptor (Monro et al., 2005), but this remains to be determined. Nevertheless, 5-HT acting via
5-HT3 and 5-HT7 receptors is thought to be involved in descending reflex pathways in the guinea pig small intestine (Monro et al., 2002; Monro et al., 2005; Tonini et al., 2005; Gwynne and Bornstein, 2007b), as well as in generating CMMCs in the mouse
(Dickson et al., 2010). Further, 5-HT-containing descending interneurons and 5-HT3 receptors have been implicated in coordinating activity in myenteric and submucosal neurons during CMMCs in mouse colon (Okamoto et al., 2012; Okamoto et al., 2014).
Myenteric neurons exhibit IPSPs, but unlike those observed in the submucosal plexus, they are insensitive to α2-adrenoceptor blockade (Johnson and Bornstein, 2004;
Gwynne and Bornstein, 2007b). Rather, 5-HT1A receptors have been shown to mediate IPSPs in myenteric AH neurons (Johnson et al., 1980; Galligan, 1996; Johnson and Bornstein, 2004). Such inhibitory inputs may contribute to regulating activity within recurrent networks, enabling the encoding of graded responses that reflect the magnitude of stimuli and are physiologically meaningful (Thomas et al., 2000; Thomas and Bornstein, 2003). 5-HT1A-mediated IPSPs have also been described in non- cholinergic VIP-secretomotor neurons and may be important in modulating the activity of these neurons (Foong et al., 2010b).
In addition to postsynaptic effects, 5-HT acting on presynaptic 5-HT4 receptors has a major role in facilitating excitatory transmitter release in the ENS of guinea pigs and mice (Pan and Galligan, 1994; Galligan, 1996; Liu et al., 2005; Gwynne and Bornstein,
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2007b). Thus, 5-HT4 receptors may be involved in strengthening activated peristaltic or secretomotor reflex pathways (Gershon, 2005; Poole et al., 2006).
In mucosa-submucosa preparations, 5-HT added to the serosal side can act on 5-HT3 receptors to stimulate secretion via activation of both cholinergic and non-cholinergic secretomotor neurons (Hendriks et al., 1989; Johnson et al., 1994). However, findings by Hendriks et al. (1989) indicate that the 5-HT3 receptor-mediated component of the 5- HT secretory response predominantly involves cholinergic secretomotor neurons, since blocking 5-HT3 receptors did not affect the residual response after muscarinic blockade with hyoscine. This is also consistent with anatomical data showing that cholinergic NPY secretomotor neurons have numerous serotonergic inputs (Neal and Bornstein, 2007). These data suggest that a different 5-HT receptor is expressed by non-cholinergic secretomotor neurons. In accordance with this, numerous studies have shown that a low concentrations of 5-HT (10 nM – 500 nM) preferentially excites non-cholinergic neurons, while a higher concentration of 5-HT activates both populations of secretomotor neurons (Keast et al., 1985b; Johnson et al., 1994; Foong et al., 2010a). Whether this applies in other species is unclear.
Cultured enteric glia also respond to 5-HT (Kimball and Mulholland, 1996), and this is inhibited by 5-HT2 antagonism (Boesmans et al., 2013b). In addition, 5-HT2A receptor expression has been described in mouse ileum and was localized to cholinergic submucosal neurons projecting to the mucosa (Gross et al., 2012). Activation of these 5-
HT2A receptors has been shown to be involved in promoting mucosal growth and proliferation. Given that glial cells may also produce trophic factors (Rühl, 2005), it is possible that glial responses to 5-HT contribute to epithelial turnover.
6.6 ATP and purinergic P2Y1 receptors
Activation of P2Y1 receptors by ATP has been associated with various forms of excitatory synaptic potentials in both submucosal and myenteric neurons (Hu et al.,
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2003; Monro et al., 2004; Fang et al., 2006; Gao et al., 2006; Foong and Bornstein, 2009; Gwynne and Bornstein, 2009).
In the myenteric plexus, P2Y1 receptors mediate slow EPSPs in NOS-immunoreactive neurons and intermediate EPSPs in NOS-negative neurons (Gwynne and Bornstein, 2009), and have a major role in transmission between NOS-immunoreactive descending interneurons of inhibitory reflex pathways (Thornton et al., 2013). ATP is also well known to be a non-adrenergic non-cholinergic (NANC) inhibitory neurotransmitter at neuromuscular junctions within intestinal smooth muscle (Burnstock et al., 1970; Burnstock, 2008), and the action of ATP on circular muscle is mediated by the P2Y1 receptor (Wood, 2006; Wang et al., 2007).
The role of P2Y1 receptors in the submucosal plexus is less clear. Hu and colleagues (2003) identified slow EPSPs mediated by ATP acting at P2Y1 receptors in submucosal neurons of guinea pig small intestine using a P2Y1-specific antagonist MRS2179. Local application of 5-HT (100 μM) to submucosal plexus preparations, which depolarizes cholinergic submucosal neurons via 5-HT3 receptors (Surprenant and Crist, 1988; Cooke et al., 1997a; Pan and Gershon, 2000), evoked MRS2179-sensitive slow EPSPs in neighbouring ganglia (Hu et al., 2003). 5-HT applied directly onto the neurons that displayed purinergic EPSPs induced depolarizing responses that were MRS2179- insensitive (Hu et al., 2003). This suggests that ATP is released by submucosal neurons and acts at postsynaptic P2Y1 receptors to excite neighbouring ganglion cells. Using electrical stimulation, they identified two other possible sources of purinergic inputs. Sympathetic fibers and myenteric neurons were both shown to evoke MRS2179- sensitive slow EPSPs in submucosal neurons when stimulated. There is also evidence for ATP as a co-transmitter with noradrenaline from sympathetic nerves supplying the gut (Burnstock, 2000). The majority of the neurons studied in these experiments that displayed slow EPSPs with a purinergic component (>87%) were VIP-secretomotor neurons.
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Fang et al., (2006) showed that applying ATP to the submucosal side of submucosal plexus and mucosa intestinal preparations evoked a secretory response that was inhibited by MRS2179. This indicates that ATP-induced secretion is primarily mediated by P2Y1 receptors and is compatible with the electrophysiology (Hu et al., 2003). The response was effectively suppressed by TTX, indicating that the secretory effects of ATP are predominantly neurally-mediated. The VIP-receptor antagonist PG-97 269 also significantly inhibited the response whereas hyoscine did not (Fang et al., 2006). These studies provide evidence supporting the proposal that P2Y1 receptors are exclusive to VIP-secretomotor neurons.
However, there are points of difference between these findings and data showing that there are P2Y1-mediated intermediate EPSPs that occur in non-VIP submucosal neurons (Bornstein et al., 1986; Monro et al., 2004). These results suggest the presence of P2Y1 receptors on cholinergic submucosal neurons. In support of this, it has been shown that P2Y1 receptors are strongly expressed by cholinergic secretomotor neurons in guinea pig colon and are involved in mediating secretomotor reflexes (Cooke et al., 2004).
ATP has a prominent role in glial signaling and various P2 receptors have shown to be expressed by enteric glia (Kimball and Mulholland, 1996; Rühl, 2005; Gomes et al., 2009; Gulbransen and Sharkey, 2009). While studies in guinea pig have shown that functional P2Y1 receptors are expressed by enteric neurons (Hu et al., 2003; Gao et al., 2006), studies in mice show that the activation of P2Y1 receptors primarily stimulates enteric glia, but not neurons (Gomes et al., 2009; Gulbransen et al., 2012; McClain et al., 2015). Rather, P2X receptors appear to have a more prominent role in neuronal signaling in the mouse intestine (Bian et al., 2003; Ren et al., 2003). Hence, there may be significant differences between the enteric circuitry of the guinea pig and mouse intestines. Enteric glia may also express other purinergic P2Y receptors including P2Y2 and P2Y4 (Kimball and Mulholland, 1996; Nassauw et al., 2006). Interestingly, studies in mouse small intestine demonstrate that both of these receptor subtypes have roles in
47 intestinal secretion (Cressman et al., 1999; Robaye et al., 2003; Ghanem et al., 2005); however, potential glial involvement in these responses remains to be elucidated. Moreover, despite the common transmitters and receptors shared by enteric neurons and glia, the mechanism by which neurons and glia communicate, and how glia may participate in transmission are largely unknown.
6.7 Nitric oxide (NO)
NO is another important inhibitory transmitter and mediates intestinal relaxation (Sanders and Ward, 1992). Two forms of NANC inhibitory transmission to circular muscle have been identified (Costa et al., 1986), one involves ATP as described earlier (Wood, 2006; Burnstock, 2008) and the other involves NO and VIP (Crist et al., 1992; He and Goyal, 1993; Holzer et al., 1997). NO may also be involved in murine colonic myoelectric complexes (Lyster et al., 1995; Spencer et al., 1998; Bornstein et al., 2010).
Inhibitory motor neurons and a subset of descending interneurons express NOS (Costa et al., 1992) (Table 1.2). However, the role of NO in synaptic transmission is not well understood. The NO donor sodium nitroprusside (SNP) did not alter resting membrane potential except for depressing slow EPSPs in some myenteric neurons in guinea pig small intestine (Tamura et al., 1993), yet NO was shown to be involved in mediating IPSPs in the distal colon (Dickson et al., 2007; Bornstein et al., 2010). NOS-expressing neurons in the submucosal plexus are rare, although NOS-immunoreactive terminals arising from myenteric descending interneurons are found in submucosal ganglia (Costa et al., 1992; Furness et al., 1994). In submucosal neurons, NO was found to enhance IPSPs and depressed non-purinergic slow EPSPs, but increased the overall excitability of VIP secretomotor neurons (Bornstein et al., 2010). As VIP secretomotor neurons receive numerous excitatory and inhibitory inputs, it was proposed that these seemingly contradicting effects of NO may in fact act to fine-tune the responses of these neurons to physiological stimuli (Bornstein et al., 2010).
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Additionally, enteric glia can also express inducible NOS (iNOS) and glial NO release has been associated with colitis (Cirillo et al., 2009; Cirillo et al., 2011; MacEachern et al., 2015; Brown et al., 2016; Ochoa-Cortes et al., 2016).
6.8 Vasoactive intestinal peptide (VIP) and VIP receptors
VIP is an important secretory mediator released by the final secretomotor neurons. In the gut, VIP is contained exclusively within neurons and in addition to secretomotor neurons, VIP-immunoreactivity occurs in myenteric interneurons that also supply submucosal neurons (Bornstein and Furness, 1988). However, whether VIP is a neurotransmitter within the circuitry is yet to be elucidated.
VIP is also co-expressed with NOS in inhibitory motor neurons (Table 1.2) and has been shown to be a mediator of inhibitory neuromuscular transmission. Activating inhibitory motor neurons that supply the circular muscle by electrical stimulation, hyperpolarizes and relaxes the smooth muscle (Crist et al., 1992; He and Goyal, 1993; Bornstein et al., 2004). Further, VIP and NO are thought to interact to induce smooth muscle relaxation, whereby NO acts presynaptically to enhance VIP release from myenteric neurons (Grider and Jin, 1993; Allescher et al., 1996; Kurjak et al., 2001) and VIP in turn stimulates NO production in smooth muscle (Rekik et al., 1996; Bornstein et al., 2004). However, the VIP receptor involved remains unknown (Van Geldre and Lefebvre, 2004).
Three VIP receptor subtypes have been identified to date: VPAC1, VPAC2, and VIPs (Harmar et al., 1998; Zhou et al., 2006). Surprisingly the exact localization of these receptors is not well understood despite the apparent importance of VIP. Various reports implicate VPAC1 receptors in mediating mucosal secretion, which suggests that they are expressed on the mucosa (Banks et al., 2005; Kordasti et al., 2006). The other site of interest is within the ENS, as it has been shown that VIP can depolarize VIP neurons (Mihara et al., 1985; Palmer et al., 1987). However, whether VIP receptors are
49 expressed on enteric neurons is unclear. Data from a radioligand-binding study in mouse small intestine suggest that VPAC1 receptors are primarily expressed in the mucosa while VPAC2 receptors are found in smooth muscle (Harmar et al., 2004). Activation of VPAC2 receptors (VPAC2R) is thought to directly relax smooth muscle as suggested by findings in the rat stomach (Robberecht et al., 1998). Whereas VIP was found to stimulate neurogenic contraction in longitudinal muscle preparations of guinea pig small intestine (Kusunoki et al., 1986; Katsoulis et al., 1992). However, the neural receptor subtype involved was not identified.
There is another neuropeptide that displays affinity for VIP receptors similar to VIP, that is, pituitary adenylate cyclase activating peptide (PACAP), which shares 68% homology with VIP (Harmar et al., 1998; Laburthe et al., 2002). Hence, VIP receptors are termed VPAC (a contraction of VIP and PACAP) receptors and likely share these two physiological ligands (Laburthe et al., 2002). There is also a PACAP-specific receptor PAC1 which does not respond to physiological concentrations of VIP (Harmar et al., 1998). PACAP was shown to stimulate secretion in flat sheet preparations of rat small intestine partly via a selective high-affinity PACAP receptor on submucosal neurons (Cox, 1992). This was likely the PAC1 receptor. The presence of PACAP- immunoreactive fibers in submucosal ganglia (Portbury et al., 1995) also suggests a role for these receptors in secretory function.
An additional VIP receptor was cloned from guinea pig taenia coli: the VIPS receptor (Zhou et al., 2006). This subtype showed high affinity for VIP, but not PACAP, and potently stimulated cAMP-formation. Thus unlike VPAC1 and VPAC2, VIPS is specific for VIP. Being structurally similar to VPAC2 (they differ only by 2 residues), VIPS was predicted to have a role in inhibition of smooth muscle (Jin et al., 1994; Zhou et al.,
2006). However, whether VIPS receptors are expressed in gut layers other than smooth muscle or in other gut regions, and whether they also have a role in secretory function have not been investigated.
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7. Effects of Cholera toxin (CT)
7.1 Cholera toxin-evoked hypersecretion
CT is considered the archetypal diarrhoea-inducing enterotoxin and CT-induced secretion has been studied in attempt to better understand not only pathological secretion but also physiological secretion. CT has been a commonly-used tool to dissect the secretomotor reflex circuitry, yet these pathways are still unclear.
CT is composed of 5 beta-subunits and an alpha subunit comprising of an A1- and an A2-chain, where the A2-chain links the enzymatically-active A1-chain to the pentameric beta-subunit (Wernick et al., 2010),. The beta-subunits allow the toxin to attach to the enterocytes via a GM1 ganglioside receptor (Holmgren et al., 1975). The toxin-GM1 receptor complex is then endocytosed and retrogradely transported to the endoplasmic reticulum (ER) (Jobling et al., 2012). Within the ER, the alpha subunit is proteolytically cleaved into the A1- and A2-chains and dissociates from the rest of the toxin. The active A1-chain then translocates to the cytoplasm (Wernick et al., 2010) and catalyzes the ADP-ribosylation of Gsα to constitutively activate adenylate cyclase (Thiagarajah and Verkman, 2005; Jobling et al., 2012). This in turn increases cAMP levels to stimulate Cl- secretion across the mucosa through CFTR (Thiagarajah et al., 2004; Satitsri et al., 2016) and water follows osmotically (Burleigh and Banks, 2007). This was initially thought to be the sole mechanism underlying CT-evoked hypersecretion. However, it appears that little of the toxin ever reaches the crypts and the toxin is mainly in contact with the tips of villi (Weiser and Quill, 1975; Lundgren and Jodal, 1997). Yet, absorptive function is largely unaffected (Hallback et al., 1979), which provides the basis for oral rehydration therapy. Furthermore, CT does not penetrate the mucosa (Gwynne et al., 2009). Taken together, it is conceivable that CT is not simply directly exerting its secretory effects at the level of the mucosal epithelium, and suggests that it has indirect effects via the ENS.
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Vibrio cholerae- and CT-evoked hypersecretion have been studied extensively using in vivo or in vitro models. De and Chatterje (1953) first developed the rabbit ligated ileal loop model to study V. cholerae pathogenicity. Basu and Pickett (1969) extended on this and further compared this ileal loop model in various laboratory animals including gerbil, rat, hamster, guinea pig, chinchilla, cat, and mouse. They reported that all animals except mice showed susceptibility to various strains of V. cholerae or CT, and displayed consistent fluid accumulation in the ileal segments (10 cm long) exposed to these toxic agents. However, more recent studies have demonstrated that V. cholerae and CT can evoke significant fluid secretion with shorter (2 – 3 cm) small intestine loops in adult mice (Luerang et al., 2012; Muanprasat et al., 2012; Sawasvirojwong et al., 2013). In vivo studies have shown that acute exposure to CT for 30 min induces net secretion within 60 – 120 min (Cassuto et al., 1982a). Hence, subsequent studies of CT hypersecretion using in vivo models tend to begin 2 h after CT incubation (Mourad et al., 1995; Turvill et al., 2000a, b; Kordasti et al., 2006).
The in vivo models are advantageous in that the effects of the toxin can be monitored as it occurs in an intact system. Fluid secretion were monitored in various ways, by measuring the volume of fluid in the ileal loops (De and Chatterje, 1953), weighing the intestinal segments, collecting the effluent and analyzing it for ion concentrations, and/or monitoring the transmural potential difference throughout the procedure (Moore et al., 1971; Field et al., 1972; Cassuto et al., 1981a; Mourad et al., 1995; Turvill et al., 2000b; Kordasti et al., 2006). However, while in vitro models may not closely reflect the changes that are occurring in the whole animal, they enable more detailed analyses of the enteric network.
7.2 ENS involvement in CT-evoked hypersecretion
ENS involvement in CT-induced secretion is well documented and evidence for this is largely based on in vivo experiments conducted in rats and cats. These experiments showed that the CT-evoked secretory response can also be attenuated or even reversed by nerve activity-blocking agents: TTX, hexamethonium, and the local anaesthetic
52 lidocaine (Cassuto et al., 1981a; Cassuto et al., 1982a; Cassuto et al., 1983; Lundgren and Jodal, 1997; Lundgren, 2002). Further, in vivo experiments conducted in rats showed that chronic extrinsic denervation did not reduce CT-induced secretion so the nerves involved must be intrinsic (Sjöqvist, 1991). Jiang et al. (1993) proposed that CT specifically and directly binds VIP-containing enteric neurons to evoke a depolarizing response. However, there is little supporting evidence for CT actually reaching the nerve plexuses (Weiser and Quill, 1975; Gwynne et al., 2009).
Thus, the in vivo data generally indicated that CT-evoked secretion was predominantly due to enteric neural activity. Yet, in vitro data tends to suggest that while CT-induced hypersecretion may be partly neurally-mediated, a significant component of the response is due to the toxin directly affecting the mucosal epithelium (Field et al., 1972; Carey and Cooke, 1986; Moriarty et al., 1989). These studies were conducted on guinea pig and rabbit small intestinal preparations. Some of these discrepancies may be due to species differences, but are yet to be resolved.
7.3 Models of neural mechanisms underlying CT-evoked hypersecretion
A more recent model suggests that CT activates the ENS via 5-HT by depolarizing EC cells (Lundgren, 2002). The ongoing release of 5-HT from EC cells seen to accompany CT-induced secretion (Beubler et al., 1989b) is thought to then over-stimulate the secretomotor pathways at the input of the circuit that is, the 5-HT-sensitive nerve endings of ISNs via 5-HT3 receptors (Bertrand et al., 2000b) (Figure 1.7). In accordance with these findings, CT exposure to the mucosa induces a depletion of 5-HT in EC cells in cats (Nilsson et al., 1983). Further, luminal administration of 5-HT evokes a neurally- mediated secretory response in rat jejunum in vivo that is blocked by hexamethonium, thereby indicating that 5-HT acts upstream of a nicotinic synapse (Cassuto et al., 1982b;
Sjöqvist et al., 1992). Finally, the 5-HT3-receptor antagonist granisetron effectively attenuates CT-induced hypersecretion (Mourad et al., 1995).
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However, it has also been shown that in vitro incubation of CT in the lumen of guinea pig jejunum induces hyperexcitability in secretomotor neurons (Gwynne et al., 2009). This suggests an alternate mechanism involving changes in the circuit properties at the level of the final secretomotor neurons which is compatible with the observation that there was no increase in spontaneous firing; if there was over-activation of the ISNs, then an increase in spontaneous neural activity would be expected and this was not the case. Consistent with previous studies, hexamethonium prevented CT-induced hyperexcitability, indicating the presence of a nicotinic synapse in the reflex pathway
(Gwynne et al., 2004). A similar effect was observed by co-incubation with NK1 and
NK3 receptor antagonists. The involvement of NK1 receptors has also been reported in rat small intestine (Turvill et al., 2000a). However, the specific locations of these nicotinic and tachykinergic synapses are unknown. Both cholinergic NPY- and VIP- secretomotor neurons became hyperexcitable with luminal CT-incubation; although the role of cholinergic neurons is less clear given that atropine (muscarinic antagonist) showed no effect on CT-secretion in vivo in rats (Cassuto et al., 1982a). It is however consistent with the finding that the cholinergic component of the electrically-stimulated secretory response was enhanced with CT-incubation (Carey and Cooke, 1986). Given that CT-induces a sustained hyperexcitability in secretomotor neurons, this was suggested to be a key mechanism underlying CT-induced hypersecretion since the activation of these neurons will lead to an enhanced secretory response (Gwynne et al., 2009).
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Figure 1.7. A simplified model of the enteric circuitry activated by CT.
CT stimulates 5-HT release from EC cells to activate the afferent limb of the pathway via 5-HT3 receptors on the nerve endings of submucosal and/or myenteric ISNs. This in turn excites interneurons and then efferent secretomotor neurons and triggers the release of the secretagogue VIP to stimulate Cl- secretion across the mucosa. The excess release of 5-HT and subsequent over-activation of secretomotor pathways were proposed to underlie CT-evoked hypersecretion. However, it has also been suggested that CT may induce a sustained hyperexcitability in the output secretomotor neurons to enhance the secretory response. Figure by Gwynne (unpublished).
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Another study by Kordasti et al. (2006) proposed that separate pathways are involved in the induction and in the maintenance of hypersecretion, although 5-HT3-receptors may be involved in both. Notably, findings from this study showed that blocking nicotinic receptors inhibited the hypersecretion, but did not prevent the induction of the response. Contrastingly, VIP antagonism was shown to prevent the induction of the response but did not inhibit the hypersecretion. This suggests that cholinergic secretomotor neurons may play a role once the hypersecretion was established, yet atropine was also shown to be ineffective in inhibiting the hypersecretory response. Thus, the involvement of cholinergic and non-cholinergic secretomotor neurons needs to be clarified.
Nevertheless, VIP released by VIP-secretomotor neurons is considered to be the major secretory mediator as CT-exposure is associated with increased levels of VIP in the lumen, and VIP antagonists have been shown to attenuate CT-evoked hypersecretion (Mourad and Nassar, 2000; Banks et al., 2005; Kordasti et al., 2006). However, there are considerable discrepancies between studies relating to the specific VIP antagonist used and its efficacy. Mourrad and Nassar (2000) demonstrated that the general VIP antagonist [4Cl-D-Phe6, Leu17]-VIP dose-dependently reversed CT-induced secretion. However, Banks et al (2005) could not replicate these findings using similar protocols and both studies were conducted in rat jejunum in vivo. Moreover, they found [4Cl-D- Phe6, Leu17]-VIP ineffective against the secretory effects of VIP. Instead, they showed that the VPAC1 receptor (a VIP receptor subtype) selective antagonist PG 97-269 significantly attenuated CT-evoked hypersecretion, and antagonized VIP in monolayers of human colonic cell line T84, and in muscle-stripped preparations of human ileum and rat jejunum. Yet, in the previously described study by Kordasti et al. (2006), [4Cl-D- Phe6, Leu17]-VIP was found to prevent induction of CT-evoked hypersecretion but was ineffective once it had established. The literature detailing VIP antagonism in general has been inconsistent, not only in relation to CT-secretion. For instance, the VIP fragment VIP(10-28) inhibited the non-cholinergic component of electrically-stimulated secretion in guinea pig colon (Reddix et al., 1994), but this was not reproducible in rabbit colon (Banks et al., 2005). VIP(10-28) was also shown to be ineffective against VIP-evoked secretion in rat jejunum and colon (Cox and Cuthbert, 1989; Burleigh and
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Kirkham, 1993; Banks et al., 2005). Undoubtedly, this is an area of study which requires more thorough investigation and a better understanding of VIP receptors will be useful in the interpretation of such data.
7.4 Effects of cholera toxin on intestinal motility
Although much of the emphasis has been placed on the submucosal plexus, there is evidence highlighting the importance of the myenteric plexus in CT-evoked hypersecretion. Specifically, ablation of the myenteric plexus significantly attenuates the CT-induced secretion (Carey and Cooke, 1986; Jodal et al., 1993). CT has rapid effects on intestinal motility (Fung et al., 2010; Balasuriya et al., 2016), where an increase in propulsive contractions was observed in the guinea pig small intestine, and a decrease in CMMCs was reported in the mouse colon. Changes to motor activity were observed within the first 15 min of toxin exposure. The time course of these immediate motility effects differs drastically to that of the hypersecretory effect which takes hours to develop. The rapid onset of these effects indicates that delay in the binding of CT to the mucosa or in the transduction of the signal across the mucosa cannot sufficiently account for the delay in its hypersecretory effects (Fung et al., 2010). Whether these early changes to motor and presumably myenteric activity are in any way associated with later changes in secretion is unclear, but it is possible that these effects contribute to building up the neural activity which leads to eventual hypersecretion. On the other hand, the rapid increase in propulsive motor activity observed in the jejunum was shown to involve a pathway independent of 5-HT3 receptors (Fung et al., 2010), and is therefore a separate pathway to that which stimulates secretion (Mourad et al., 1995; Kordasti et al., 2006). Similar findings have been described in rats in vivo, where CT was seen to increase contractile activity evoked by distension, and this effect was more prominent after inhibiting 5-HT3 receptors (Kordasti et al., 2006). Although these effects were observed during the period when hypersecretion had established. Nevertheless, whether CT evokes changes to activity within the circuitry of the myenteric and submucosal plexus, the specific types of enteric neurons that may be involved, and how these may contribute to the pathogenic effects of CT remain to be elucidated.
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8. Thesis Aims
This thesis has two major aims. One is to elucidate the VIP and VPAC receptor function within the enteric circuitry. The first study assessed whether functional VPAC receptors are expressed in the ENS. The second study extends on this and examined the role of VIP and VPAC receptors in signaling the enteric circuitry by exploiting transgenic Wnt1-Cre;R26R-GCaMP3 mice in which enteric neurons and glia express a fluorescent calcium indicator. The second major aim is to investigate the effects of CT on the enteric nervous system and in particular, the involvement of cholinergic and non- cholinergic VIP secretomotor neurons. This was addressed in the third study using an in vivo ileal loop model of CT-incubation and Wnt1-Cre;R26R-GCaMP3 mice to assess the excitability of the enteric circuitry.
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CHAPTER 2: MATERIALS AND METHODS
As numerous experimental techniques were used to obtain the data in this thesis and several techniques were used for multiple studies, this chapter collates and describes all the methods employed. Included are methods for PCR experiments (Chapter 3), immunohistochemistry (Chapters 3 – 5), Ussing Chamber studies (Chapters 3 and 5), measuring muscle contractility (Chapter 3), genotyping (Chapter 5), calcium imaging (Chapters 4 and 5), ligated ileal loop surgery (Chapter 5), and cryosectioning (Chapter 5). However, technical specifications unique to individual results chapters are detailed in the respective chapters. Additionally, as many drugs were also used across multiple studies, they have been compiled and are listed in this chapter (Table 2.4).
1. Animals
Guinea pigs (180 – 350g) and mice (aged 6 – 12 weeks) of either sex, unless specified otherwise, were used for experiments in this thesis. Guinea pigs were first stunned by a rapid blow to the back of the head, and killed by severing the carotid arteries and spinal cord and mice were killed by cervical dislocation; all procedures were approved by the University of Melbourne Animal Experimentation Ethics Committee.
2. Identification and quantification of Vipr1 and Vipr2 messenger RNA
2.1 Tissue preparation
Guinea pigs of either sex were killed and the abdominal cavity was opened. A 4 – 5 cm segment of jejunum was isolated, flushed clean of intestinal contents, and placed in sterile phosphate buffered saline (PBS, 4°C). Using sterilised tools, the jejunal segment
59 was then cut along the mesenteric border, stretched and pinned flat on a silicone elastomer-lined dish (Sylgard 184, Dow Corning, North Ryde, NSW, Australia) chilled with ice. The 4 layers of the tissue were then separated by microdissection: samples of mucosa, submucosal plexus, longitudinal muscle and myenteric plexus, and circular muscle were obtained. The samples were frozen in liquid nitrogen immediately after they were dissected out and then stored at -80°C.
2.2 RNA extraction
The frozen intestinal samples were homogenized in a Wig-L-Bug® amalgamator (Dentsply-Rinn, Elgin, IL, USA). 1 mL TRI Reagent™ (Ambion, Applied Biosystem, Scoresby, VIC, Australia) was added per 100 mg tissue for total RNA extraction. The homogenised samples were thoroughly vortexed, incubated at room temperature for 10 min, and then centrifuged (12,000 x g) for 10 min at 4˚C. Chloroform (200 μl per 1 ml TRI Reagent™) was added to the resulting supernatant and incubated for 10 min at room temperature. Samples were then centrifuged (12,000 x g) for 15 min at 4˚C. The RNA-containing aqueous phase was mixed with isopropanol (500 μl per 1 ml TRI™ Reagent), while discarding the DNA-containing inter-phase and protein-containing organic phase. Samples were then centrifuged (12,000 x g) for 30 min at 4˚C for RNA precipitation. The RNA pellets obtained were washed twice with 0.5 ml 75% ethanol prior to resuspension in RNASecure™ water (Ambion). RNA purity and concentrations were measured on a NanoDrop® 1000 Spectrophotometer (NanoDrop Technologies Inc, Wilmington, DE, USA). Samples with A260:A280 absorbance ratios within the range of 1.8 and 2.1 were deemed sufficiently pure for further PCR analysis. A lower ratio may be indicative of contamination by protein, phenol or other organic chemicals that absorb strongly at 280 nm.
2.3 First strand complementary DNA synthesis
Each reaction used a total volume of 20 μl containing 1 μg total RNA diluted in nuclease free water, 4 μl 5X First Strand buffer (contains 250 mM Tris-HCl, 375 mM
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KCl, 15 mM MgCl2), 1 μL dNTP mix (10 mM), 1 μl random hexamers (50 ng/μl), 1 μl 0.1 M dTT, 1 μl SuperScript™ III Reverse Transcriptase enzyme (200 U/μl), and 1 μl RNaseOUT™ Recombinant RNase Inhibitor (40 U/μl, all from Invitrogen Australia, Mount Waverley, VIC, Australia). A negative control in which the reverse transcriptase enzyme was replaced with an equal volume of nuclease free water was also included to control for genomic DNA contamination. First strand cDNA synthesis was performed by incubating samples at 25°C for 5 min, 50°C for 60 min, and 70°C for 15 min to terminate the reaction. Samples were then stored at -20°C.
2.4 Reverse transcriptase polymerase chain reaction
Reverse transcriptase polymerase chain reaction (RT-PCR) was performed to determine the presence or absence of Vipr1 and Vipr2 gene transcripts (which encode the VPAC1 and VPAC2 receptors respectively) in the mucosa, submucosal plexus, longitudinal muscle and myenteric plexus, and circular muscle layers of guinea pig jejunum. Guinea pig specific forward and reverse primers were designed for Vipr1 and Vipr2 using the online tool Primer3 and synthesised by GeneWorks (Adelaide, SA, Australia). Each primer set was designed to span multiple introns (Table 2.1) in order to minimise transcripts amplified from genomic DNA, as synthesis of longer amplicons is less efficient.
Each reaction used 1 μl cDNA in a 25 μl reaction mixture containing 12.5 μl GoTaq Green Master Mix (Promega, Annandale, NSW, Australia), 1 μl forward and 1 μl reverse oligonucleotide primer (10 μM) for the gene of interest, and 9.5 μl nuclease free water. A negative control in which nuclease free water replaced the cDNA template was included in each RT-PCR experiment. A positive control for cDNA synthesis using oligonucleotide primers for the reference gene glyceraldehyde-3-phosphate dehydrogenase (gapdh) was also included. RT-PCR was performed in a MyCycler™ PCR machine (Bio-Rad Laboratories Pty., Ltd., Gladesville, NSW, Australia). PCR cycler was set to perform 1 cycle of initial denaturation (85°C for 2 min), 40 cycles of denaturing (94°C for 1 min), annealing (at specific annealing temperatures for each
61 primer set for 1 min; Table 2.1) and extension (72°C for 1 min), and 1 final cycle of extension (72°C for 10 min). Gel electrophoresis (2% 1X Tris-acetate-EDTA buffer agarose gels with ethidium bromide) was performed to resolve the PCR products (20 μl of each sample) at 100V for 50 min. A 100 bp DNA ladder (Hyperladder IV; Bioline Pty Ltd., Alexandria, NSW, Australia) was used as a reference to approximate PCR product size. Gels were viewed and imaged using the UVP BioDoc-IT™ system (2UV Transilluminates, Pathtech, Box Hill, VIC, Australia).
PCR product bands were cut from the gels for DNA extraction using a QIAquick Gel Extraction kit (Qiagen, Doncaster, VIC, Australia). Samples were then sequenced by the Applied Genetic Diagnostics laboratory (Department of Pathology, The University of Melbourne, VIC, Australia) to verify that the targeted genes were identified.
The sequenced PCR products were confirmed to be the genes of interest using BLAST (NCBI). The guinea pig Vipr1 sequence was shown to share 85% homology with that of mice, and 84% homology with that of rats. Similarly, the guinea pig Vipr2 sequence shares 89% and 87% homology with the mouse and rat Vipr2 sequences respectively. The nucleotide sequences obtained were then used to predict the amino acid sequence (Expert Protein Analysis System, ExPASy; Swiss Institute of Bioinformatics). Matching guinea pig amino acid sequences were identified for all predicted amino acid sequences using protein BLAST (NCBI), further verifying that targeted gene products were amplified (Table 2.2).
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Table 2.1. RT-PCR primer sequences for guinea pig Vipr1, Vipr2, and gapdh.
Guinea pig Vipr1, Vipr2 and gapdh sequences were obtained from the NCBI nucleotide database (see accession IDs). The PCR conditions for Vipr1 were more stringent (increased annealing temperature and reduced cycles) to eliminate non-specific amplification.
Gene Guinea pig Primer Sequences (5’ - 3’) Annealing Temp Location Accession ID
Vipr1 Forward TGTTTCACAGCGAGGAGTTG 57°C Exon 5 JN225407
Reverse TGCAGACGAACAGGACAAAG (reduced to 35 cycles) Exon 9
Vipr2 Forward CGTGCTGGTCAAGGATGATA 55°C Exon 5 NM_001173121
Reverse GGTGCTGTGGTCATTTGTGT Exon 8
gapdh Forward ACCCAGAAGACTGTGGATGG 55°C Exon 7 NM_001172951.1
Reverse TGCTGTAGCCGAACTCATTG Exon 11
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Table 2.2. Guinea pig Vipr1 and Vipr2 receptor nucleotide and amino acid sequences.
Gene Sequence (5’-3’)
Vipr1 Nucleotide GGCTGCAAGGCGGCCATGGTCTTCTTCCAGTACTGCGTCATGGCCAACTTCTTCTGGCTGCTGGTG GAGGGGCTCTATCTGCACATGCTGCTCGCCGTCTCCTTCTTCTCGGAGCGGAAGTACTTCTGGGG GTACATTCTCATCGGCTGGGGGGTGCCCAGCACATTCATCATGGTGTGGACTGTCGTCAGGATCC ACATTGAGGATGTTGGATGCTGGGACACCATTAACACCCCCGTGTGGTGGATCATCA
Amino acid GCKAAMVFFQYCVMANFFWLLVEGLYLHMLLAVSFFSERKYFWGYILIGWGVPSTFIMVWTVVRIHI EDVGCWDTINTPVWWII
Vipr2 Nucleotide ACTGCCTCAACCAGCCATCCTCTTGGGTTGGCTGCAAGCTCAGCCTGGTATTCTTCCAGTACTGCA TCATGGCAAACTTCTACTGGCTGCTGGTAGAGGGGCTGTACCTTCACACTCTCCTGGTGGCCATCT TCTCCCCCAGCAGATGCTTCCTAGCCTACCTGCTGATTGGATGGGGCATCCCCACCATTTGCATTG GAGTGTGGACGGCTGCCCGG
Amino acid CLNQPSSWVGCKLSLVFFQYCIMANFYWLLVEGLYLHTLLVAIFSPSRCFLAYLLIGWGIPTICIGVWT AAR
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2.5 Quantitative real time polymerase chain reaction (qPCR)
Forward and reverse primers and 6-carboxy fluorescein-labelled (FAM) TaqMan probes were designed using RealTimeDesign™ (Biosearch Technologies, Inc., Novato, CA, USA). For each gene of interest, at least one of the primers and probes was designed to either span an exon-exon junction or a long intron sequence to avoid amplification from genomic DNA (Table 2.2). qPCR was performed using an Opticon 2 PCR machine (Bio-Rad). Samples were prepared in triplicate containing 1 - 2 μl of cDNA, forward and reverse primers (10 μM), FAM-labelled probe (10 μM) and 1X SensiMix (Quantace Pty Ltd, Alexandria, NSW, Australia) in a total reaction volume of 20 μl. Rn18s was used as the reference gene. Samples were incubated at 95°C for 10 min for polymerase activation. The thermal cycler was programmed to perform 40 cycles of 95°C for 10 seconds (denaturation) and 60°C for 1 min (annealing and extension), and a fluorescence detection step following each cycle. All qPCR reactions for each gene of interest were performed in parallel to ensure comparable levels of gene expression between all tissue samples. Every qPCR included two negative controls: a sample that lacked reverse transcriptase during cDNA synthesis and a sample in which water substituted for the cDNA template.
Data was analysed using the comparative cycle threshold (2-ΔCT) method (Livak and Schmittgen, 2001). Each gene of interest was normalized to the endogenous reference gene (i.e. Rn18s), thus controlling for differences in the amount of total RNA between tissue types. This was calculated by first averaging all cycle threshold (CT) values for each triplicate. The averages of Rn18s CT values were then subtracted from the corresponding averages of the gene of interest CT values. Normalised gene expression was calculated as 2^-ΔCT*1000 (where ΔCT is the difference in threshold cycles between the gene of interest and reference).
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Table 2.3. qPCR primer and probe sequences for guinea pig Vipr1, Vipr2, and rat Rn18s.
TaqMan probes were labelled with FAM and BHQ1. Sequences for guinea pig Vipr1, Vipr2 and rat Rn18s were obtained from the NCBI nucleotide database (see accession IDs).
Gene Sequences (5’ - 3’) Location Accession ID
Guinea pig Vipr1 Forward CGGCTACACCATCGGCTAC Exon 4 JN225407
Reverse TGCAGTGCAGCTTCCTGA Exon 4/5
Probe TGTCCCTTGCCACCCTCCTG Exon 4
Guinea pig Vipr2 Forward CACCCAGAATGCCGATTTCATC Exon 1 NM_001173121
Reverse GCCAGGGCAGGATTTGTAC Exon1/2
Probe AGAAACAAAATGTTCAGAGCTTCTCAG Exon 1
Rat Rn18s Forward GCATGGCCGTTCTTAGTTGG V01270.1
Reverse TGCCAGAGTCTCGTTCGTTA
Probe TGGAGCGATTTGTCTGGTTATTCCGA
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3. Immunohistochemistry
3.1 Tissue preparation
The abdomen of the animal was dissected opened and a 4 - 5 cm segment of jejunum was excised. The segment was placed in PBS and flushed clean of any contents. The segment was then opened along the mesenteric border, and stretched and pinned flat on an elastomer-lined dish. Tissues were fixed in 4% formaldehyde/PBS either for 1 h 50 min at room temperature or overnight at 4°C. After clearing the fixative with 3 x 10 min PBS washes, preparations of submucosal plexus, and myenteric plexus with the attached longitudinal muscle were obtained by microdissection.
Preparations were permeabilized with 1% triton X-100/PBS (ProSciTech, Thuringowa, QLD, Australia) for 30 min at room temperature and washed in PBS (3 x 10 min). Preparations were double or triple-labelled by incubating with various combinations of primary antisera for 24 - 72 h at 4°C, washed in PBS (3 x 10 min), and then incubated in secondary antisera for 2 - 2.5 h at room temperature. Following PBS washes (3 x 10 min) to remove excess antisera, preparations were mounted onto slides using Dakocytomation fluorescence mounting medium (Carpinteria, CA, USA). Details of the specific antisera used are described in the respective results chapters (refer to Chapters 3 – 5).
3.2 Microscopy
Fluorescently labelled preparations were viewed through a 20x or 40x objective on a Axio Imager D.1 microscope and imaged with a AxioCam MRc5 camera using AxioVision software (all from Zeiss, Australia). Some preparations were viewed under an epifluorescence microscope (BX 41 Olympus, Olympus, Aartselaar, Belgium) and images
67 were acquired with an XM10 (Olympus) camera using Cell^F software. Confocal images were recorded using a Zeiss LSM780 confocal microscope (Cell Imaging Core, KU Leuven, Belgium) or Zeiss Pascal confocal microscope (Biological Optical Microscopy Platform, The University of Melbourne, VIC, Australia). Images were taken using 20x, 40x, or 63x objectives.
3.3 Cell counts
For double- or triple-labelling studies, the proportion of each neuronal subtype was calculated by examining co-expression with at least 75 Hu+ submucosal neurons or 200 Hu+ myenteric neurons in each preparation, where Hu labels all neurons. The mean proportion of each neuronal subtype was determined by averaging the proportions obtained from three to five animals. Data are expressed as means ± standard error of the mean (SEM).
4. In vitro measurement of short-circuit current in Ussing chambers
4.1 Tissue preparation
A segment of guinea pig jejunum or mouse ileum (guinea pig: 8 – 10cm; mouse: 4 – 5 cm in length) was excised and immediately placed in physiological saline (composition in mM:
NaCl 118, NaHCO3 25, D-glucose 11, KCl 4.8, CaCl2 2.5, MgSO4 1.2, NaH2PO4 1.0) bubbled with 95% O2: 5% CO2. The tissue was then opened along the mesenteric border and pinned flat in a silicone elastomer-lined dish. For mucosa-submucosa preparations, tissues were pinned serosal side up and the serosa, myenteric plexus and smooth muscle layers were removed by microdissection. Full thickness preparations refer to preparations in which all intestinal layers are intact. Each jejunal segment provided up to four Ussing chamber preparations. Each preparation was mounted across an opening (guinea pig: 12
68 mm pin circle diameter, 9 mm reservoir opening, CHM2; mouse: 5.5 mm pin circle diameter, 4 mm reservoir opening, CHM8; World Precision Instruments, Inc. (WPI), Sarasota, FL, USA) and divides the two halves of the Ussing-chamber. The chamber was then connected to the reservoir which contains the superfusate and these components are held in place with a custom-made apparatus (Department of Physiology, University of Melbourne, VIC, Australia). The exposed mucosal and serosal surfaces of the preparation were independently superfused in an equal volume (approximately 10 ml) of continuously circulating oxygenated physiological saline from separate reservoirs. The volumes of physiological saline in the two reservoirs were balanced to ensure equal hydrostatic pressure across the preparation. A supply of 95% O2: 5% CO2 via the injection ports of the reservoirs provided a “gas lift” enabling constant circulation of physiological saline. Reservoirs were also water-jacketed for the circulating solution to be maintained at 37°C.
4.2 Electrical measurements
Transepithelial voltage potential (Vt) was measured using a potentiometer (VCC600 Single Channel Voltage-Current Clamp; Physiological Instruments, Inc., World Trade Drive, San Diego, CA, USA) and two Vt-sensing electrodes (SR4 Reference Electrode Calomel Separable; ThermoFisher Scientific, Station Road, Auchtermuchty Fife, Scotland, UK) separately immersed in 3 M KCl, and connected to each half-chamber by salt bridges (polyethylene tubing OD 1.52 mm x ID 0.86 mm; Tyco Electronics, Huntingdale, VIC, Australia) containing 3% agar melted in 3 M KCl. Separate current-passing electrodes (Ag- AgCl electrodes; WPI) were connected to each half-chamber by larger salt bridges (ID 5 mm; WPI) also containing 3% agar melted in 3 M KCl. Ussing- chambers were first set up with physiological saline alone to eliminate electrical bias in measurements by “zeroing” the system (Clarke, 2009). This procedure involved offsetting the voltage difference between the Vt sensing electrodes and compensating for fluid resistance in the absence of tissue. The intestinal preparation was then mounted in the Ussing chamber with fresh
69 physiological saline and the spontaneous Vt was clamped to zero. Short-circuit current (ISC) was measured throughout the experiment.
Preparations were equilibrated for 40 min prior to collecting data to allow for the basal Isc of the tissue to stabilize. Experimental protocols were restricted to a maximum duration of 3 hours from the time of mounting the preparations to ensure minimal compromise to the integrity of the mucosal epithelium during experimentation. Drugs were added directly to, and diluted in, the reservoir of physiological saline superfusing the serosal half-chamber to achieve the final concentration. The total volume of physiological saline in each half- chamber was 10 ml. The maximum volume of each drug addition was limited to 100 μl to minimise imbalances in reservoir volumes.
4.3 Data analysis
Data collection and analysis were performed using AcqKnowledge 3.9.0 software (BIOPAC Systems, Inc., SDR Clinical Technology, Middle Cove, NSW, Australia). The maximum change in Isc from baseline (ΔIsc) induced by drug addition was calculated (i.e. the baseline was subtracted from the peak of the response). Data obtained from tissue of the same animal were matched; control data from adjacent segments of guinea pig jejunum or adjacent segments of mouse ileum were not significantly different.
5. In vitro measurement of longitudinal muscle contractility
The abdominal cavity of the guinea pig was opened and a segment of jejunum (8 – 10 cm in length) was isolated and placed in physiological saline bubbled with 95% O2: 5% CO2. The segment of jejunum was opened along the mesenteric border, and then pinned flat on a
70 silicone elastomer-lined dish. The mucosa, submucosa, and circular muscle layers were removed by microdissection. The remaining preparation of myenteric plexus and longitudinal muscle was cut into longitudinal strips (1.5 – 2 cm in length), then suspended in a 6 ml organ bath and attached to isotonic transducers (SDR Technology, Sydney, NSW, Australia) with a resting tension of 0.3 grams. Preparations were continuously bathed with physiological saline bubbled with 95% O2: 5% CO2 and maintained at 37°C. The longitudinal strips were equilibrated for 40 min to allow a stable baseline to be established before beginning the experiment.
Drugs were added directly to, and diluted in, the organ bath to achieve the final concentration. Drugs were washed out by 3 complete changes of physiological saline and each wash was separated by 5 min. The baseline was allowed to stabilize for 10 min after the final wash before further drug additions. Contractile responses were recorded using a Biopac M100A (SDR Technology) with a sampling rate of 10 s-1. Contractile responses to drugs were measured as the maximal change to muscle length (mm) from baseline. Data were analysed using Acqknowledge 3.2.4 software (SDR Technology). Data are presented as mean ± SEM.
6. Genotyping
Wnt1-Cre;R26R-GCaMP3 mice (C57BL6 background), which have the fluorescent Ca2+ indicator GCaMP3 encoded in neural crest-derived cells i.e. enteric neurons and glia, were used for calcium imaging experiments. Wnt1-Cre;R26R-GCaMP3 mice were generated by mating female homozygous floxed R26R-GCaMP3 mice with male heterozygous Wnt1-Cre mice (Danielian et al., 1998; Zariwala et al., 2012). The Wnt1-cre transgene allows for the specific expression of cre recombinase in neural crest- derived cells. As homozygous floxed R26R-GCaMP3 mice were used for breeding,
71 all offspring are expected to carry a copy of the floxed R26R-GCaMP3 allele. Thus, GCaMP3 expression depends on the presence of Wnt1-cre. Offspring carrying the Wnt1-cre transgene were identified by genotyping DNA extracted from mouse tails (Figure 2.1). Wnt1-Cre;R26R-GCaMP3 mice were generated by genetic crosses rather than by maintaining a breeding colony of Wnt1-Cre;R26R-GCaMP3 mice to avoid the possibility of transgene inactivation, which results in a loss of transgene expression over time, or genetic drift where modifying mutations may occur with continuous breeding within a colony (Davis et al., 2012).
A maximum of 2 mm of the tip of the tail was removed from 21 – 28 day old transgenic mice using sharp sterile scissors by the Biomedical Sciences Animal Facility staff (The University of Melbourne, VIC, Australia). Tail samples were stored at -20°C prior to DNA extraction. DNA was isolated from transgenic mouse tails (2 – 3 mm) using NaOH extraction. Tail samples were cut into small pieces and incubated in 100 µl 25 mM NaOH/0.2 mM (pH 12) on a heating block at 98°C for 90 min. Samples were vortexed twice during this incubation period. Samples were allowed cool before adding 100 µl 40 mM Tris.HCl (pH 5) to neutralise the extraction solution. The samples were centrifuged and then stored at 4°C until use.
The extracted tail DNA samples were genotyped for cre recombinase using the primers Cre 850 (5’- CGCCGTAAATCAATCGATGAGTTGCTTC -3’) and Cre 403 (5’- GATGCCGGTGAACGTGCAAAACAGGCTC -3’), which generates a 450 bp PCR product (Figure 2.1). Each reaction used 2 μl DNA in a 20 μl reaction mixture containing 4 μl 5 x MangoTaq coloured reaction buffer (Bioline), 0.5 μl dNTP mix (10 mM, Invitrogen), 0.5 μl Cre 850 and 0.5 μl Cre 403 primers (20 μM; GeneWorks), 0.1 μl MangoTaq DNA Polymerase (Bioline) and 11.8 μl nuclease free water. A negative control in which nuclease free water replaced the DNA template was included in each PCR experiment to ensure that there was no contamination. A positive control using a DNA sample from a previously
72 genotyped Wnt1-cre expressing mouse was also included to ensure the PCR was effective. PCR was performed in a MyCycler™ PCR machine (Bio-Rad). PCR cycler was set to perform 1 cycle of initial denaturation (94°C for 2 min), 35 cycles of denaturing (94°C for 30 s), annealing (62°C for 30 s) and extension (72°C for 30 s), and 1 final cycle of extension (72°C for 10 min). Gel electrophoresis (1.5% 1X Tris-boric acid-EDTA buffer agarose gels with ethidium bromide) was performed to resolve the PCR products (10 μl of each sample) at 100V for 40 min. A 100 bp DNA ladder (Hyperladder IV; Bioline) was used as a reference to approximate PCR product size. Gels were visualized and imaged using a ChemiDoc™ MP System with Image Lab™ 4.1 software (Bio-Rad).
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Figure 2.1. Identifying the presence or absence of Wnt1-cre expression.
PCR performed on DNA extracted from mouse tails (corresponding mice are identified by numbers at the top of the gel). The presence of the Wnt1-cre transgene was identified by a 450 bp band and marked by a + symbol. A positive control using DNA from a previously genotyped sample and negative control where H2O replaced DNA were included. A 100 bp DNA ladder was used as a reference to approximate PCR product size.
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7. Calcium imaging
7.1 Tissue preparation
Male C57BL6 mice and Wnt1-Cre;R26R-GCaMP3 mice of either sex (aged 6 – 12 weeks) were killed by cervical dislocation, a procedure approved by the animal ethics committee of the University of Leuven, Belgium and the University of Melbourne Animal Experimentation Ethics Committee. A segment of jejunum (4 – 5 cm long) or ileal loop (as described in the following section) was collected in 95% O2: 5% CO2 -bubbled physiological saline. The tissue was opened along the mesenteric border and pinned flat with mucosa up in a silicone elastomer-coated dish. The submucosal plexus with the mucosa intact was separated from the underlying muscle layers by microdissection. The circular muscle was then removed to obtain myenteric plexus preparations. The resulting submucosal and myenteric plexus preparations were stabilized by stretching the tissue with the plexus facing up over an inox ring and then clamped over with an O-ring for imaging (Vanden Berghe et al., 2002). Up to 5 ring preparations were obtained from each jejunal segment. Preparations were constantly superfused (1 ml/min) with 95% O2: 5% CO2 - bubbled physiological saline at room temperature throughout the experiment.
7.2 Imaging
Two microscopy setups were used for calcium imaging. In Leuven (BE), a 20× (NA 1.0) water dipping lens was used on an upright Zeiss Examiner microscope (Carl Zeiss, Oberkochen, Germany) equipped with a monochromator (Poly V) and cooled CCD camera (Imago QE) both from TILL Photonics (Gräfelfing, Germany). Images (640 × 512) on this system were acquired at 2 Hz. In Melbourne (AU), preparations were viewed through an Olympus 20× (NA 0.5) water dipping lens on an upright Zeiss Axioscope microscope with a Zeiss AxioCam MRm camera, and images (278 × 278) were acquired at 1 Hz.
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Ganglia of interest were stimulated chemically or electrically. Agonists were applied to the preparations by pressure ejection (9 psi.; Intracel Picospritzer III, Parker Hannifin, Hollis, NH, USA) via a micropipette (tip diameter ~20 µm) positioned immediately adjacent to and above the selected ganglion. Micropipettes were made from borosilicate glass capillaries (Harvard GC1500-15, inner diameter (ID) 1.5 mm x outer diameter (OD) 0.86 mm or Harvard GC100F-10 OD 1.0 mm x ID 0.5 mm) using a micropipette puller (Model 2-9, Narishige Scientific Instrument, Japan, or P-87, Sutter Instrument Co., Novato, CA, USA). The concentration of agonist used to fill the micropipette was at least 10-fold higher than that used when applying by superfusion, to allow for dilution within the bath solution (Liu et al., 1999; Breunig et al., 2007). For electrical stimulation, a single pulse or trains of 300 μs pulses (2 s, 20 Hz; Grass SD9 or S88 stimulator, Grass Instruments, Warwick, RI, USA) were applied by a stimulating electrode (tungsten wire, 50 µm diameter) positioned on an internodal strand leading to the ganglion imaged.
After calcium imaging experiments, some preparations were fixed in 4% formaldehyde/PBS for 45 min at room temperature or overnight at 4°C for post hoc immunohistochemistry.
7.3 Data Analysis
Neurons were identified based on the size of their cell bodies (~20 µm diameter) (Gabella and Trigg, 1984) and lack of fluorescence in their nuclei. Glia were identified based on the size of their cell bodies (< 5 µm diameter) (Gabella and Trigg, 1984), and their morphology and location (either intraganglionic type I cells with irregular branched processes, or elongated type II cells within/at the edge of internodal strands) (Boesmans et al., 2015b). Ca2+-imaging analyses were performed with custom-written routines in Igor Pro (Wavematrics, Lake Oswego, OR, USA). Regions of interest were drawn to calculate the 2+ average [Ca ]i signal intensity. Values were then normalized to the baseline fluorescence
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2+ intensity (Fi/F0). Reponses were considered when the [Ca ]i signal increased above 2+ baseline by at least 5 times the intrinsic noise. [Ca ]i peaks were calculated for each 2+ response, with the peak amplitude taken as the maximum increase in [Ca ]i from baseline
(∆Fi/F0).
8. In vivo incubation of cholera toxin (CT) in a ligated ileal loop
In accordance with procedures approved by the University of Melbourne Animal Experimentation Ethics Committee, male mice on a C57BL6 background, including Wnt1- Cre;R26R-GCaMP3 mice, were removed from food overnight and were first induced under anaesthesia using 4 – 5% isoflurane by inhalation, before gradually lowering the levels to 1.5 – 2% to maintain anaesthesia throughout the surgical procedure. The mouse was placed on a heat pad warmed to 40°C to maintain its body temperature throughout the procedure. The surgery was performed under aseptic conditions. The abdomen of the mouse was first shaved and bupivacaine hydrochloride (0.05%) was administered locally at the site of abdominal incision. A laparotomy was then performed and a 3 – 4 cm ileal loop was constructed from a region of ileum that was selected because it was clear of gut contents, where possible. First, the anal end of the chosen loop was tied off with surgical suture at least 3 cm proximal to the ileocecal junction. The segment was then injected with either sterile physiological saline (control), or CT (12.5 ug/ml) in physiological saline, and tied off at the oral end with surgical suture. The exposed intestine was kept hydrated throughout the procedure with warmed (37°C) sterile physiological saline. The abdomen was closed with stitches and the mice were allowed to recover from the anaesthesia. During the recovery phase, the animal had access to drinking water. After a 3.5 h incubation post- surgery, the mice were killed by cervical dislocation. For immunohistochemistry, the ileal loop was collected in PBS. For Ussing chamber and calcium imaging studies, the ileal loop was collected in physiological saline.
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9. Cryosectioning and anti-CT-B labelling
Segments of control and CT-incubated ileal loops were fixed in 4% formaldehyde/PBS for 1 h 50 min at room temperature and then immersed in 30% sucrose/dH2O overnight at 4°C for cryoprotection. Ileal loop segments were then cut into 2 – 3 mm tubes and placed in OCT (Tissue Tek, Elkhart, IN, USA). Samples were snap frozen in isopentane cooled by liquid nitrogen. Isopentane allows for rapid and homogenous freezing, as compared with directly freezing in liquid nitrogen, where nitrogen gas can form around the warmer sample leading to irregular freezing patterns (Steu et al., 2008). Samples were stored at -80°C prior to sectioning. Sections of tissue (10 μm) were cut using a cryostat (Microm HM 525, Fronine Laboratory Supplies, Riverstone, NSW, Australia) and collected on positively charged SuperFrostPlus slides (Menzel-Glaser, Braunschweig, Germany). Sections allowed to dry for 1 h at room temperature.
Hydrophobic wells encompassing each section on the slide were created using a hydrophobic PAP pen (Abcam, Redfern, NSW, Australia). A primary antiserum against CT-B subunit (goat α CT-B, 1:2500; List Biological Laboratories Inc., Campbell CA, USA; kind gift from Colin Anderson, University of Melbourne, VIC, Australia) diluted in PBS was applied to each section. Slides were placed in humidified containers and left to incubate overnight at 4°C. Excess primary antiserum was cleared by 3 x 10 min PBS washes. Sections were incubated in secondary antiserum (donkey α sheep Alexa 488, 1:200; A11015 Molecular Probes) diluted in PBS for 3 hours at room temperature in a dark humidified container. Excess secondary antiserum was removed with 3 x 10 min PBS washes. Coverslips were applied to the slides with DAKO mounting medium. Sections were then viewed and imaged using a confocal microscope to localize CT-B staining and establish whether CT penetrated the mucosal layer, into the underlying nerve plexuses.
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10. Drugs
Tetrodotoxin (TTX; Alomone Labs, Jerusalem, Israel; Acros, Geel, Belgium), 2-methyl- thio-ADP (2MeSADP), 1,1-dimethyl-4-phenylpiperazinium (DMPP), hexamethonium bromide, and hyoscine hydrobromide (all from Sigma Aldrich), MRS2179 (Abcam; Tocris, Bristol, UK), Vasoactive Intestinal Peptide (VIP; guinea pig; Auspep, Tullamarine, VIC, Australia), and Cholera toxin (List Biologicals, Campbell, CA) were dissolved in distilled water to make stock solutions. Veratridine (Sigma Aldrich) was dissolved in 80% ethanol. Stocks were stored at 4°C.
The VPAC1 receptor antagonist (PG 97-269; Phoenix Pharmaceuticals, Belmont, CA, US; Mimotopes, Clayton, VIC, Australia), VPAC2 receptor antagonist (PG 99-465), VPAC1 agonist [K15, R16, L27]VIP(1-2)/GRF(8-27), and VPAC2 agonist, BAY 55-9837 (all from Mimotopes), VIP (human, mouse, rat; Bachem, Bubendorf, Switzerland), and PPADS (pyridoxal-phosphate-6-azophenyl-2',4'-disulfonic acid; Sigma Aldrich) were prepared as stock solutions using distilled water and stored as aliquots at -20°C.
Drugs were prepared as 100-fold or 1000-fold concentration stock solutions and were diluted in physiological saline to achieve the desired final concentration immediately before use.
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Table 2.4. Drugs and their targets.
Antagonist Selective or specific target/s TTX-sensitive and TTX-insensitive voltage- Bupivacaine gated Na+ channels (Scholz et al., 1998) + Voltage-gated Na channels (Nav1.1, Nav1.2,
Tetrodotoxin Nav1.3, Nav1.6, and Nav1.7) (Bartoo et al., 2005) Muscarinic acetylcholine receptors Hyoscine hydrobromide (Keast et al., 1985b) Nicotinic acetylcholine receptors Hexamethonium bromide (Keast et al., 1985b) PG 97-269 VPAC1 receptors (Gourlet et al., 1997a) PG 99-465 VPAC2 receptors (Moreno et al., 2000) MRS2179 P2Y1 receptors (Burnstock, 2007) PPADS P2 receptors (Burnstock, 2007) Agonist VIP (guinea pig) VPAC1 and VPAC2 receptors VIP (human, mouse, rat) (Dickson and Finlayson, 2009) [K15, R16, L27] VIP (1-2)/GRF(8-27) VPAC1 receptors (Gourlet et al., 1997b) BAY 55-9837 VPAC2 receptors (Tsutsumi et al., 2002) 2MeSADP P2Y1 receptors (Burnstock, 2007) Nicotinic acetylcholine receptors DMPP (Keast et al., 1985b) Voltage-gated Na+ channels Veratridine (Sheldon et al., 1990)
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CHAPTER 3: VPAC1 RECEPTORS REGULATE INTESTINAL SECRETION AND MUSCLE CONTRACILITY BY ACTIVATING CHOLINERGIC NEURONS IN GUINEA PIG JEJUNUM
This chapter has been published in the American Journal of Physiology – Gastrointestinal and Liver Physiology (2014); 306(9), G748-58. It is reproduced here with minimal alterations.
1. ABSTRACT
In the gastrointestinal tract, vasoactive intestinal peptide (VIP) is found exclusively within neurons. VIP regulates intestinal motility via neurally mediated and direct actions on smooth muscle and secretion by a direct mucosal action, and via actions on submucosal neurons. VIP acts via VPAC1 and VPAC2 receptors; however, the subtype involved in its neural actions is unclear. The neural roles of VIP and VPAC1 receptors (VPAC1R) were investigated in intestinal motility and secretion in guinea pig jejunum. Expression of VIP receptors across the jejunal layers was examined using RT-PCR. Submucosal and myenteric neurons expressing VIP receptor subtype VPAC1 and/or various neurochemical markers were identified immunohistochemically. Isotonic muscle contraction was measured in longitudinal muscle-myenteric plexus preparations. Electrogenic secretion across mucosa-submucosa preparations was measured in Ussing chambers by monitoring short-circuit current. Calretinin+ excitatory longitudinal muscle motor neurons expressed VPAC1R. Most cholinergic submucosal neurons, notably NPY+ secretomotor neurons, expressed VPAC1R. VIP (100 nM) induced longitudinal muscle contraction that was
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inhibited by TTX (1 µM), PG 97–269 (VPAC1 antagonist; 1 µM), and hyoscine (10 µM), but not by hexamethonium (200 µM). VIP (50 nM)-evoked secretion was depressed by hyoscine or PG 97–269 and involved a small TTX-sensitive component. PG 97–269 and TTX combined did not further depress the VIP response observed in the presence of PG 97–269 alone. In conclusion, VIP stimulates ACh-mediated longitudinal muscle contraction via VPAC1R on cholinergic motor neurons. VIP induces Cl- secretion directly via epithelial VPAC1R and indirectly via VPAC1R on cholinergic secretomotor neurons. No evidence was obtained for involvement of other neural VIP receptors.
2. INTRODUCTION
Vasoactive intestinal peptide (VIP) is one of the most abundant neuropeptides in the gut. It is found in many different enteric neuronal subtypes, including interneurons, inhibitory motor neurons, and secretomotor neurons (Furness, 2000). Although the presence of VIP in these subtypes is conserved across species (Keast et al., 1985a; Furness et al., 1995b), its functions within the enteric circuitry are unclear. VIP depolarizes many myenteric (Williams and North, 1979; Palmer et al., 1987) and submucosal neurons (Mihara et al., 1985; Reed and Vanner, 2001) and is found in many axon varicosities within the myenteric (Neal and Bornstein, 2008) and submucosal plexuses (Evans et al., 1994; Reed and Vanner, 2001). However, its role in synaptic transmission and the neuronal subtypes that respond to VIP are unknown. VIP is implicated in pathologies such as bacterial toxin-induced diarrhea (Burleigh and Banks, 2007) and irritable bowel syndrome (Palsson et al., 2004), and a better understanding of its actions may prove beneficial for the development of novel therapeutics.
VIP is known to exert its effects by activating two main G protein-coupled receptor subtypes: VPAC1 and VPAC2 (Harmar et al., 1998; Laburthe et al., 2002; Dickson and
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Finlayson, 2009). In addition, a high-affinity VIP-specific receptor subtype has been identified in guinea pig taenia coli (Teng et al., 2001; Zhou et al., 2006). However, the VPAC receptor subtypes expressed by specific enteric neuron subtypes have not been investigated.
Identification of the specific roles of VIP in the gut has been difficult because of the apparent overlapping expression of its receptors on various effector cells and neurons. VIP is clearly important in the control of intestinal motility since VIP knockout mice display disrupted intestinal transit (Lelievre et al., 2007). In the myenteric plexus, VIP is contained within inhibitory motor neurons and descending interneurons (Furness, 2000; Qu et al., 2008). Muscle contractility studies in guinea pig ileum and colon indicate that VIP directly relaxes circular muscle via activation of VPAC2 receptors (VPAC2R) on the smooth muscle (Robberecht et al., 1998; Harmar et al., 2004) but also stimulates neurogenic contraction in longitudinal muscle preparations (Kusunoki et al., 1986; Katsoulis et al., 1992). Motility studies in rat colon demonstrated involvement of VIP and VPAC1 receptors (VPAC1R) (Shi and Sarna, 2008). However, VPAC1R expression on the smooth muscle and/or myenteric neurons has not been examined. VIP is also a major player in the regulation of water and electrolyte secretion. VIP can directly stimulate the mucosa, which has been shown to express VPAC1Rs (Reddix et al., 1994). In addition, secretion studies have shown that VIP also regulates neurogenic secretory tone, but the receptor subtype involved in this is unclear. Overall, understanding the involvement of VIP in physiology and pathophysiology has been challenging because of the multifaceted role of VIP in motility and secretion. A better understanding of the specific neural VPAC receptor that mediates these effects will provide more insight.
Evaluation of VPAC subtypes involved in gut functions has been hampered by lack of reliable antagonists. Nonspecific VIP antagonists have provided some insight, but some inconsistencies in findings between studies have arisen. For instance, the effectiveness of
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the VIP fragment, VIP(10–28), as a VIP antagonist appears to differ between studies (Cox and Cuthbert, 1989; Burleigh and Kirkham, 1993; Reddix et al., 1994; Banks et al., 2005), perhaps because of species differences. Yet, despite using the same animal model, another non-specific VIP antagonist, [4Cl-D-Phe6,Leu17]VIP, was reported to reverse cholera toxin- induced secretion by one group (Mourad and Nassar, 2000) but to be ineffective by another (Banks et al., 2005). However, the selective VPAC1R antagonist PG 97–269 has been used successfully to attenuate cholera toxin-evoked hypersecretion and antagonize VIP effects in monolayers of the human colonic cell line T84 in human ileum and in rat jejunum (Banks et al., 2005).
An antiserum against VPAC1R and the specific VPAC1R antagonist PG 97-269 were employed to examine the roles of this receptor in muscle contractility and secretomotor control in guinea pig jejunum. It was found that VIP acting via VPAC1R on excitatory motor neurons stimulates ACh-mediated longitudinal muscle contraction. Furthermore, VIP was found to activate VPAC1R on cholinergic secretomotor neurons.
3. MATERIALS AND METHODS
3.1 Identification and quantification of Vipr1 and Vipr2 mRNA
Tissue preparation
Guinea pigs of either sex (180–350 g) were killed by stunning and severing the carotid arteries and spinal cord according to protocols approved by the University of Melbourne Animal Experimentation Ethics Committee. Segments of jejunum (4 – 5 cm) were removed from the animal and immediately placed in PBS (4°C).
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RNA extraction
The tissue was segregated into mucosa, submucosa, longitudinal muscle and myenteric plexus, and circular muscle by microdissection. These samples were frozen with liquid nitrogen. Total RNA was extracted for first strand cDNA synthesis as previously described (Foong et al., 2010b) (Chapter 2, sections 2.2 and 2.3).
Reverse transcriptase-PCR
Expression of VIP receptor 1 gene (Vipr1) and VIP receptor 2 gene (Vipr2) transcripts (encode VPAC1R and VPAC2R, respectively) in the layers of guinea pig jejunum was first assessed using RT-PCR (Chapter 2, section 2.4). Each RT-PCR reaction used 1 μl cDNA in a 25 μl reaction containing GoTaq Green Master Mix (Promega, Annandale, SA, Australia), with forward and reverse oligonucleotide primers designed to span an intron (GeneWorks, Hindmarsh, SA, Australia). Guinea pig specific primers for gapdh were included as a positive control. PCR reactions were performed in a MyCycler PCR machine (Bio-Rad, West Ryde, NSW, Australia). PCR products were sequenced by the Applied Genetic Diagnostics laboratory (Department of Pathology, The University of Melbourne, Parkville, VIC, Australia).
Quantitative real-time polymerase chain reaction
Quantification of Vipr1 and Vipr2 gene expression was accomplished using the
ΔΔCT method with ribosomal 18S (Rn18s) as the endogenous control (refer to Chapter 2, section 2.5). Guinea pig specific forward and reverse primers and 6-carboxy fluorescein- labeled TaqMan probes for Vipr1 and Vipr2 were designed using RealTimeDesign (Biosearch Technologies, Novato, CA; Table 2.3). Optimal concentrations for all primers and probes were 10 μM. Quantitative real-time polymerase chain reaction (qPCR)
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experiments were performed on the Opticon 2 PCR machine (Bio-Rad) in triplicate 20 μl volumes containing 1× SensiMix dT (Quantace, Alexandria, NSW, Australia) and 1 – 2 μl cDNA. Mean relative gene expression was plotted with the SEM.
3.2 Immunohistochemical localization of VPAC1R
A jejunal segment (5 cm long) was opened along the mesenteric border, pinned flat, and fixed in 4% paraformaldehyde overnight at 4°C. Tissues were cleared of fixative (3 × 10 min PBS washes) before submucosal plexus and myenteric plexus preparations were dissected and processed for immunohistochemistry as previously described (Gwynne et al., 2009) (Chapter 2, section 3). Briefly, preparations were permeabilized with 1% Triton X- 100 (ProSciTech, Thuringowa, QLD, Australia) for 30 min and then washed (3 × 10 min; PBS). Primary antibodies were applied and incubated overnight at 4°C. Each submucosal preparation was triple-labeled for Hu (pan-neuronal marker), VPAC1R, and one of three markers of cholinergic neurons: choline acetyltransferase (ChAT, all cholinergic neurons), neuropeptide Y (NPY, secretomotor), or calretinin (secretomotor/vasodilator) (Furness, 2000) (Table 3.2). Each myenteric preparation was triple-labeled for Hu, VPAC1R, and one of three markers for different functional subsets of neurons: calbindin (intrinsic sensory neurons), calretinin (excitatory longitudinal muscle motor neurons and ascending interneurons), and nitric oxide synthase (NOS; inhibitory motor neurons and descending interneurons) (Furness, 2000) (Table 3.2). After being washed (3 × 10 min; PBS), preparations were incubated in secondary antibodies (Table 3.2) for 150 min at room temperature. The preparations were washed again (3 × 10 min; PBS) before mounting in Dakocytomation fluorescent mounting medium (Carpentaria).
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3.3 Cell counts
Submucosal plexus and myenteric plexus preparations were imaged using a Zeiss Pascal confocal microscope. The proportion of each neuronal subtype (ChAT+, NPY+, calretinin+, calbindin+, NOS+, and/or VPAC1R+) was obtained by examining co-expression with at least 100 Hu+ neurons in each preparation. The mean proportion of each neuronal subtype was determined by averaging the proportions obtained from three to five animals. Data are expressed as means ± SEM.
3.4 In vitro measurement of longitudinal muscle contractility
Tissue preparation
The abdominal cavity of the animal was opened, and a segment of jejunum (8 – 10 cm in length) was excised and placed in physiological saline (Krebs; composition in mM: 118
NaCl, 25 NaHCO3, 11 D-glucose, 4.8 KCl, 2.5 CaCl2, 1.2 MgSO4, and 1.0 NaH2PO4) bubbled with carbogen.
As described in Chapter 2 (section 5), myenteric plexus and longitudinal muscle preparations were cut into longitudinal strips (1.5 – 2 cm in length) and then suspended in a 6-ml organ bath and attached to isotonic transducers (SDR Technology, Sydney, NSW, Australia) with a resting tension of 0.3 grams. Preparations were continuously bathed with carbogenated physiological saline maintained at 37°C. Contractile responses were recorded using a Biopac M100A (SDR Technology). Data were analyzed using Acqknowledge 3.2.4 software (SDR Technology).
The guinea pig longitudinal strips were equilibrated for 40 min. Time control experiments involved three repeated additions of VIP (100 nM) separated by washout (complete
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changes of the bath solution; 3 × 5 min). In separate experiments, various antagonists were added before the second VIP addition and allowed to incubate for 3, 10, or 15 min.
3.5 In vitro measurement of short-circuit current in Ussing chambers
The gut segment was opened along the mesenteric border and pinned flat on a silicone elastomer-lined dish (Sylgard 184; Dow Corning, North Ryde, NSW, Australia). Mucosal- submucosal preparations were produced by removing the smooth muscle layers by microdissection. Each animal provided up to four mucosa-submucosa preparations. Ussing chambers were set up as described previously (Foong et al., 2010a) (Chapter 2, section 4). Briefly, preparations were mounted in Ussing chambers (CHM2; World Precision Instruments, Sarasota, FL). The mucosal and serosal surfaces were independently bathed in continuously circulating carbogenated physiological saline (10 ml) maintained at 37°C. The transepithelial voltage potential of the preparation was clamped to zero, and short-circuit current (ISC), a measure of epithelial ion transport (Clarke, 2009), was monitored. The tissue was equilibrated for 40 min before data recording. Drugs were added to the serosal half-chamber.
Experimental protocol
Experiments were limited to a duration of 3 h from mounting the preparation to minimize deterioration of the mucosal epithelium. The protocol involved four agonist additions at 20- min intervals. Each addition was separated by a washout (3 changes of fresh physiological saline at 5-min intervals) once the agonist-evoked response reached a plateau. The third and fourth time points were chosen for further pharmacological analysis, since agonist-induced responses at these times did not differ significantly from each other.
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3.6 Data measurement and statistics
The maximum change in muscle length and maximum change in ISC from baseline (ΔISC) evoked by addition of drug was measured. Experiments conducted with tissue from the same animal were matched; control data from adjacent segments of jejunum were not significantly different. Results are presented as mean Δlength ± SEM and mean ΔISC ± SEM. Muscle contractility data were analyzed by a one-way ANOVA, and secretion data were analyzed by a two-way ANOVA, followed by a Bonferroni's post-test where appropriate to determine statistical significance. P < 0.05 was considered significant.
3.7 Drugs used
All drugs were prepared as stock solutions using distilled water. Drugs were directly added to the physiological saline bathing the longitudinal muscle preparations or diluted in the reservoir of physiological saline supplying the serosal side of the Ussing chamber to achieve the desired final concentration. The maximum volume of each drug added did not exceed 100 μl.
Tetrodotoxin (TTX; Alomone Labs, Jerusalem, Israel), hyoscine hydrobromide and hexamethonium bromide (both from Sigma Aldrich, Castle Hill, NSW, Australia), and VIP (guinea pig; Auspep, Tullamarine, VIC, Australia) were stored at 4°C. [Ac-His1,D- Phe2,Lys15,Arg16,Leu27]VIP(1–7)/growth hormone-releasing factor-(8–27) or PG 97–269 (Phoenix Pharmaceuticals, Belmont, CA) was stored as aliquots at −20°C. TTX was incubated in the bathing solution for 15 min, hyoscine and hexamethonium were incubated for 10 min, and the VPAC1R antagonist was incubated for 3 min before agonist addition.
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4. RESULTS
4.1 Differential expression of Vipr1 and Vipr2 across the gut layers
Vipr1 and Vipr2 expression was analyzed in separated layers of gut, including the mucosa, submucosal plexus, longitudinal muscle and myenteric plexus, and circular muscle. Both Vipr1 and Vipr2 gene transcripts were detected by RT-PCR in all layers (Figure 3.1A). The Vipr1 PCR bands appeared strongest in the mucosa and weakest in the circular muscle. Substantial levels of Vipr1 expression were detected in the submucosal plexus and longitudinal muscle plus myenteric plexus layers. The Vipr2 PCR bands were of similar intensity across all layers. Sequenced Vipr1 and Vipr2 PCR products were analyzed using nucleotide BLAST (NCBI), which confirmed that target gene products were amplified. The guinea pig Vipr1 sequence shared 85% homology with that of mice and 84% homology with that of rats. Similarly, the guinea pig Vipr2 sequence shares 89 and 87% homology with mouse and rat Vipr2 sequences, respectively. Sequence analysis showed that the PCR products corresponded to an 84-amino acid fragment of the VPAC1R and a 72-amino acid fragment of the VPAC2R (Expert Protein Analysis System, ExPASy; Swiss Institute of Bioinformatics).
Quantitative analysis of Vipr1 and Vipr2 expression is shown in Figure 3.1 (B, C). Expression of Vipr1 was significantly higher in the mucosa (P < 0.001) than in any other gut layer (Figure 3.1B), although substantial levels were also detected in the submucosal plexus. However, the level of expression detected in the longitudinal muscle and myenteric plexus was relatively low. In contrast, Vipr2 expression was highest in the submucosal plexus, which was notably higher than in the mucosa (P < 0.001) and longitudinal muscle and myenteric plexus (P < 0.05) (Figure 3.1C).
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Adcyap1r1 [which encodes for the PAC1 receptor (PAC1R)] expression was also examined by RT-PCR and was found in all gut layers, whereas quantitative analysis showed that Adcyap1r1 expression was very low in the mucosa relative to the other layers (data not shown).
4.2 Cholinergic excitatory motor neurons and secretomotor neurons express VPAC1R
The expression of VPAC1R within the myenteric and submucosal plexuses was then examined using immunohistochemistry. VPAC1R immunoreactivity was observed in 52 ± 2% of all myenteric neurons (n = 8 animals; Hu+ 5,158 neurons, Figure 3.2) and 42 ± 4% of all submucosal neurons (n = 5 animals; 2,088 Hu+ neurons, Figure 3.3). VPAC1R staining was specific to neurons, particularly localized to the cytoplasm of their cell bodies and nerve fibers (Figures 3.2 and 3.3). Both strongly and weakly staining neurons were seen. In the submucosal plexus, VPAC1R+ neurons were observed to be predominantly located in the periphery of the ganglia (not quantified).
In the myenteric plexus almost all calbindin+ intrinsic sensory neurons (Furness, 2000) were VPAC1R+ (99 ± 0.4%; Figure 3.2A – D; n = 3 animals). A substantial proportion of calretinin+ excitatory motor neurons and ascending interneurons (Furness, 2000) were also VPAC1R-immunoreactive (61 ± 5%; n = 4 animals; Figure 3.2E – H), whereas only 0.88 ± 1% of NOS+ neurons expressed VPAC1R (n = 3 animals; Figure 3.2I – L). Interestingly, in the submucosal plexus, a large proportion of VPAC1R+ submucosal neurons expressed ChAT: 78 ± 4% of VPAC1R+ neurons were ChAT+ and 61 ± 4% of ChAT+ neurons were VPAC1R+ (n = 5 animals; Figure 3.3A – D). Moreover, the majority of NPY+ secretomotor neurons (Furness, 2000) were VPAC1R-immunoreactive (78 ± 4%; n = 4 animals; Figure 3.3E – H), whereas only 14 ± 8% of calretinin+ neurons expressed VPAC1R (n = 4 animals; Figure 3.3I – L).
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Figure 3.1. mRNA expression of vasoactive intestinal peptide receptor 1 gene (Vipr1) and vasoactive intestinal peptide receptor 2 gene (Vipr2) in the gut layers of guinea pig jejunum.
A. Vipr1 mRNA was expressed in the mucosa (M), submucosal plexus (SMP), longitudinal muscle and myenteric plexus (LMMP), and at a low level in circular muscle (CM). The reference gene gapdh was the positive control. A reverse transcriptase negative or RT(−) and a water (H2O) sample served as negative controls. The second band in the RT(−) is likely to be a product amplified from genomic DNA, since these primers did not span an intron. The marker was Hyperladder IV. Quantitative analysis of Vipr1 (B) and Vipr2 (C) expression. Vipr1 expression in the mucosa is significantly higher than in all other tissue layers (B), whereas Vipr2 expression in the mucosa is lower than in both submucosa and circular muscle, and expression is highest in the submucosa (C) (*P < 0.05 and **P < 0.001; One-way ANOVA with a Bonferroni's post-test). Data are presented as means ± SEM.
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Figure 3.2. Localizing VPAC1 receptor (VPAC1R) in the myenteric plexus of guinea pig jejunum.
Images of representative myenteric ganglia, demonstrating neurons stained for Hu (A, E, and I), VPAC1R (B, F, and J), calbindin (C), calretinin (G), and nitric oxide synthase (NOS; K). Merged images of Hu, VPAC1R, and calbindin (D) demonstrate colocalization of VPAC1R with calbindin in all VPAC1R+ neurons. Merged images of Hu, VPAC1R, and calretinin (H) demonstrate colocalization of calretinin with some VPAC1R+ neurons. In contrast, the merged image of Hu, VPAC1R, and NOS (L) shows a lack of colocalization of VPAC1R with NOS. Scale bars = 20 µm. Filled arrows indicate colocalization of VPAC1R with the neuronal subtype marker, whereas open arrows indicate a lack of colocalization between the markers.
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Figure 3.3. Localizing VPAC1R in the submucosal plexus of guinea pig jejunum.
Images of representative submucosal ganglia, demonstrating neurons stained for Hu (A, E, and I), VPAC1R (B, F, and J), choline acetyltransferase (ChAT, C), neuropeptide Y (NPY, G), and calretinin (K). Merged images of Hu, VPAC1R, and ChAT (D) and Hu, VPAC1R, and NPY (H) demonstrate colocalization of VPAC1R with ChAT/NPY in the majority of VPAC1R+ neurons. In contrast, the merged image of Hu, VPAC1R, and calretinin (L) shows a lack of colocalization of VPAC1R with calretinin. Scale bars = 10 µm. Filled arrows indicate colocalization of VPAC1R with the neuronal subtype marker, whereas open arrows indicate a lack of colocalization between the markers.
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4.3 Role of VPAC1R in longitudinal muscle contraction and intestinal secretion
Given the expression of VPAC1R on calretinin excitatory motor neurons and cholinergic secretomotor neurons, the effects of VIP on contractions of longitudinal muscle-myenteric plexus preparations and on electrogenic secretion in submucosa-mucosa preparations were examined. Muscle contractility experiments were conducted on tissues from 20 animals (35 preparations). Ussing chamber experiments were conducted on tissues from 60 animals 2 (112 preparations); basal ISC was 70 ± 4 μA/cm (n = 112).
4.4 VIP evokes longitudinal smooth muscle contractions by activating VPAC1Rs on neurons
Application of VIP (100 nM) evoked reproducible contractions in longitudinal muscle preparations (0.39 ± 0.10 mm; n = 8; Figure 3.4A). The VIP-evoked contractions were reversibly blocked by incubating the tissue with TTX (voltage-gated Na+ channel blocker; 1 μM) for 15 min before VIP application (P < 0.05; n = 7; Figure 3.4B), indicating that VIP was acting on neurons and not directly on the smooth muscle. VIP-evoked contractions were also significantly inhibited in the presence of the VPAC1R antagonist PG 97–269 (1 μM; P < 0.01; n = 6; Figure 3.4C), an effect that persisted after washout. Similarly, the muscarinic antagonist hyoscine (10 μM) significantly inhibited the VIP-evoked contractile response (P < 0.01; n = 8; Figure 3.4D), and its effects did not wash out. In contrast, the VIP response was unaffected by nicotinic receptor blockade (hexamethonium, 200 μM; n = 6; Figure 3.4E).
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Figure 3.4. Effects of various antagonists on vasoactive intestinal peptide (VIP)- induced contraction in longitudinal muscle and myenteric plexus preparations.
Addition of VIP consistently induced longitudinal muscle contraction, as represented by a change in length of the preparation. A. Time controls for VIP (100 nM) additions, where each addition was separated by 3 × 5 min washout with normal Krebs. B. The VIP-evoked contraction was abolished in the presence of tetrodotoxin (TTX, 1 μM; n = 7). C. The VIP response was almost abolished in the presence of PG 97–269 (1 μM; n = 6) and hyoscine (HYO, 10 μM; n = 8) (D) but was unaffected by hexamethonium (200 μM; n = 6) (E).
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4.5 VIP-mediated secretion involves activation of cholinergic secretomotor neurons
Application of VIP (10 – 300 nM) to the serosal side of the preparation induced a concentration-dependent increase in ISC (Figure 3.5A). VIP at 50 nM produced a submaximal response; this concentration was used for pharmacological analyses (Figure 3.5B – D). Time controls involved four repeated VIP (50 nM) additions separated by 3 × 5 min washouts to ensure reproducible responses were achieved. The first and second additions evoked responses that were significantly different from each other and were smaller than the Δ ISC induced by the third and fourth additions (first addition: 64 ± 5 μA/cm2; second addition: 98 ± 12 μA/cm2; P < 0.05; n = 9; Figure 3.5B). The third and fourth additions of VIP produced consistent responses (third addition: 123 ± 15 μA/cm2; fourth addition: 119 ± 19 μA/cm2; n = 9), and thus, in later experiments, the effects of antagonist and washout were examined at these time points, respectively. Because the magnitude of the VIP responses varied between preparations, responses were normalized to the second VIP exposure for each experiment. Data from antagonist experiments were compared with the VIP time controls using two-way ANOVA.
The response to VIP was usually biphasic; an initial phase of the response was present in 95% of the preparations examined. However, because the peak of the initial phase was often subtle, only the second phase was quantified. TTX (1 μM) abolished the initial phase in all preparations examined and reversibly depressed the second phase (P < 0.05; n = 7, Figure 3.5C). The TTX-resistant VIP response was about 80% of the corresponding VIP time control. Hyoscine (10 μM) abolished the first phase in every preparation that displayed a biphasic response and irreversibly depressed the second phase (P < 0.01; n = 5, Figure 3.5D) to about 70% of the corresponding time control. Furthermore, there was no involvement of fast nicotinic synaptic transmission, since the VIP response in the presence of hexamethonium (200 μM) was not significantly different from that of the paired time control (39 ± 8 and 37 ± 8 μA/cm2 respectively; n = 9).
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4.6 VIP evokes a small neurogenic secretory component involving VPAC1R
PG 97–269 (1 μM) was used to examine the role of VPAC1R in the VIP-evoked Δ ISC. This VPAC1R selective antagonist was applied 3 min before the addition of VIP, since this duration of incubation was previously reported to be most effective (Fang et al., 2006). The antagonist blocked the initial phase of the VIP response in 33% of preparations examined and reversibly inhibited the second phase (n = 6; P < 0.001; Figure 3.5C) to about 37% of the corresponding time control.
PG 97–269 and TTX were applied together to determine whether there is a VPAC1R- independent neural response to VIP. The first phase of the response was abolished by the VPAC1R antagonist and TTX combined in 83% of preparations examined. The combination of both antagonists reversibly depressed the second phase of the response (n = 6) compared with time controls (n = 9; P < 0.01, Figure 3.5C). Their combined effect was no greater than that of PG 97–269 alone, but it exceeded that of TTX alone. Hence, no VPAC1R-independent neural effect could be detected.
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Figure 3.5. Effects of various antagonists on the second phase of VIP-induced change in short-circuit current (ΔISC).
A. Addition of VIP (10–300 nM) to the serosal side of the jejunum preparation induced a concentration-dependent increase in ISC, with 50 nM producing a submaximal response. B. Repeated additions of VIP (50 nM), where each addition was separated by 3 × 5 min wash with normal Krebs. The first two additions (solid black bars) evoked responses that were significantly different from each other and were significantly smaller than the 3rd and 4th additions of VIP (striped bars; n = 9). Because the 3rd and 4th VIP additions produced consistent responses, these time points were used to examine the effects of antagonist and washout. C. The VIP response was significantly reduced in the presence of PG 97–269 (1 μM; n = 6), TTX (1 μM; n = 7), and the combination of both antagonists (n = 6), but TTX did not further reduce the response in the presence of PG 97–269. D. The VIP-evoked response was significantly reduced in the presence of hyoscine (10 μM; n = 5) compared with the VIP time control (n = 9).
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4.7 Neurally mediated basal secretion involves VPAC1R
Baseline ISC of the time controls did not change significantly over the course of the experiment. TTX significantly depressed basal secretion by 63% (P < 0.01; paired t- test; n = 11). Hyoscine reduced the baseline by 15% (P < 0.05; n = 10), and PG 97–269 reduced the baseline by 12% (P < 0.05; n = 16). However, hexamethonium did not affect basal secretion (n = 9). Application of PG 97–269 together with hyoscine also significantly depressed basal ISC by 15% (P < 0.05; n = 5) but to a similar extent as hyoscine and PG 97– 269 alone.
5. DISCUSSION
These results demonstrate the involvement of neural VPAC1R in both motility and secretion. VPAC1R were localised to cholinergic motor neurons innervating longitudinal muscle and cholinergic secretomotor neurons. Furthermore, it was demonstrated that this receptor subtype is involved in neurally mediated longitudinal muscle contraction and neurogenic secretion with no evidence for involvement of non-cholinergic neurons in either response. VIP-mediated effects involve activation of cholinergic neurons that is independent of nicotinic neurotransmission. These data also confirm that a major action of VIP is to directly activate secretion via VPAC1R on the mucosal epithelium. Collectively, it appears that VPAC1Rs are involved at the output end of the motor pathways of the myenteric and submucosal plexuses.
5.1 Exogenous VIP activates VPAC1R to evoke longitudinal muscle contraction
VIP was previously associated with smooth muscle relaxation via a direct effect on smooth muscle (Robberecht et al., 1998; Harmar et al., 2004). This study showed that VIP evokes neurogenic longitudinal muscle contraction, since the VIP-evoked contraction was abolished in the presence of TTX. This complements previous reports in guinea pig ileum
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and distal colon (Katsoulis et al., 1992). This is the first demonstration that VPAC1R is the receptor subtype primarily involved in this response, since it was almost abolished by the VPAC1R antagonist PG 97–269. Hence, there is little evidence for the involvement of any other receptor subtypes, namely VPAC2R or the VIP-specific receptor. VPAC1R was immunohistochemically localised to calretinin+ myenteric neurons, many which are excitatory motor neurons innervating the longitudinal smooth muscle (Furness, 2000). VPAC1R immunoreactivity was also localized to calbindin+ neurons (myenteric intrinsic sensory neurons), consistent with VIP depolarizing myenteric afterhyperpolarizing neurons (Palmer et al., 1987), although the role of VPAC1R on these neurons is less clear. Calretinin is also contained within cholinergic ascending interneurons. However, there is little evidence for the involvement of either the ascending interneurons or the intrinsic sensory neurons in VIP-induced contraction, since blocking fast nicotinic transmission with hexamethonium did not alter the response. There was virtually no colocalization of VPAC1R with NOS (which labels inhibitory motor neurons), so this receptor subtype is unlikely to be involved in neurally mediated smooth muscle relaxation. This is also consistent with the notion that VIP directly relaxes circular smooth muscle (Robberecht et al., 1998; Harmar et al., 2004). VIP-evoked contraction appears to be exclusively mediated via ACh, since the response was blocked by hyoscine. These results also suggest that VIP has no direct effect on the longitudinal muscle. If VPAC1R is indeed only localized to myenteric neurons, perhaps a lack of VPAC1R expression in the longitudinal muscle led to the low level of Vipr1 expression detected in the longitudinal muscle and myenteric plexus samples by qPCR. Collectively, the data indicate that VIP directly activates excitatory motor neurons via VPAC1R to evoke ACh release, thereby activating muscarinic acetylcholine receptors on the longitudinal muscle to cause contraction. The most likely source of VIP input is from descending interneurons since it has been previously shown that VIP-immunoreactive varicosities closely oppose the cell bodies of calretinin+ myenteric neurons in guinea pig jejunum (Neal and Bornstein, 2008).
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5.2 Exogenous VIP activates VPAC1R on cholinergic secretomotor neurons
The PCR data showed expression of Vipr1 mRNA in submucosal plexus, and VPAC1R immunoreactivity was identified in cholinergic submucosal neurons, notably by most NPY secretomotor neurons. From the secretion studies, the biphasic VIP response suggests that VIP evokes both cholinergic and non-cholinergic responses (Carey and Cooke, 1986; Carey et al., 1987). The first phase was abolished by hyoscine and TTX, indicating that it results from activation of cholinergic neurons. Furthermore, hyoscine reduced the second phase, and, similarly, TTX also reduced the second phase, albeit only by a small amount. These results are consistent with the finding of VPAC1R on cholinergic neurons and collectively indicate that VIP stimulates neurons in a cholinergic secretomotor pathway. However, like VIP-evoked longitudinal muscle contractions, VIP-evoked secretion was also unaltered by blockade of nicotinic fast synaptic transmission. This suggests that VIP directly excites cholinergic secretomotor neurons. Although Vipr2 expression was highest in the submucosa, there was no evidence for the involvement of VPAC2R or the VIP-specific receptor. Rather, the neural component of the VIP response appears to involve VPAC1R, since TTX in combination with the VPAC1R antagonist did not further reduce the response. Considering that Vipr2 transcripts were expressed in the submucosa, an effective antiserum to localize VPAC2R would be invaluable in providing insight into the role of these receptors. This study and several others have shown that submucosal neurons in guinea pig are divided into two major populations (ChAT+ and VIP+) (Furness et al., 1984; Bornstein and Furness, 1988; Song et al., 1992), and it was also found that submucosal neurons that did not label for ChAT are VIP+ (data not shown). Thus, by inference, because 20% of VPAC1R+ neurons were non-cholinergic, these neurons must also be VIP+. Nonetheless, there was no evidence of VIP acting on VIP secretomotor neurons.
These results also show that TTX and the VPAC1R antagonist separately and in combination reduced basal secretion, supporting previous reports of neurogenic secretory
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tone (Keast et al., 1985b; Carey and Cooke, 1989b; Fang et al., 2006) and that VIP neurons are tonically active (Carey and Cooke, 1989b).
5.3 Signaling pathways involving VIP, PACAP, and NO
VIP and pituitary adenylate cyclase-activating polypeptide (PACAP) are closely related in the VIP family of peptides. PACAP activates PACAP-selective PAC1R and displays similar affinity for the VPAC receptors as VIP (Vaudry et al., 2000). PACAP, like VIP, is involved in regulating motility and secretion (Kuwahara et al., 1993; Katsoulis and Schmidt, 1996). It was shown in the guinea pig ileum that PACAP is expressed by neuronal fibers in the myenteric plexus and smooth muscle (Portbury et al., 1995), and by inhibitory motor neurons innervating circular muscle (Furness, 2000). This is consistent with the finding in guinea pig small intestine that Adcyap1r1 is expressed in the myenteric plexus and smooth muscle layers. Portbury and others (1995) reported that the submucosal neurons did not express PACAP in their cell bodies and that there were PACAP-immunoreactive fibers in submucosal ganglia but not the mucosa of guinea pig small and large intestines (Portbury et al., 1995). In accordance with this, Adcyap1r1 expression was observed in the submucosal plexus, and the level of mRNA detected in the mucosa was the lowest compared with the other gut layers (Portbury et al., 1995). This suggests that PAC1R is not likely to play a major role in secretory function at the level of the mucosal epithelium.
There are strong interactions between VIP, PACAP, and NOS in neuronal reflex pathways. In rat studies, VIP and PACAP release is regulated by nitric oxide (NO) and vice versa in the control of smooth muscle tone, whereas VIP and NO show synergistic effects in regulating secretion (Mourad et al., 2003; Bornstein et al., 2004). In the guinea pig ileum, VIP and PACAP are colocalized with NOS in inhibitory motor neurons, which innervate circular muscle, whereas VIP and NOS are colocalized in inhibitory motor neurons, which supply the longitudinal muscle (Furness, 2000). Furthermore, VIP- and NOS-
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immunoreactive interneurons from the myenteric plexus provide inputs to VIP and cholinergic submucosal neurons (Li et al., 1995). Interestingly, NO suppresses some slow excitatory postsynaptic potentials and enhances inhibitory postsynaptic potentials, yet increases the overall excitability of VIP secretomotor neurons to depolarization. Thus, NO may be acting to fine tune the numerous inputs to VIP neurons (Bornstein et al., 2010). Hence, it is possible that NO may modulate the activity of VIP interneurons, which provide inputs to VPAC1R+ excitatory motor neurons (Furness, 2000; Neal and Bornstein, 2008) and secretomotor neurons (Li et al., 1995; Reed and Vanner, 2001).
5.4 VPAC1R is the major receptor subtype in the mucosa
The Vipr1 transcript was most strongly expressed in the mucosa, with substantial levels in the submucosa and much lower expression in the muscle. Expression of Vipr2, which encodes the other major VIP receptor subtype VPAC2, was significantly lower in the mucosa. This is consistent with VPAC1R being the major receptor subtype mediating VIP- evoked secretion (Banks et al., 2005; Burleigh and Banks, 2007). Accordingly, the VIP response was found to be largely inhibited by blocking VPAC1R. Furthermore, the VIP response was inhibited by the VPAC1R antagonist to a larger extent than by TTX; this together with the PCR data indicates that VIP-evoked secretion is mainly mediated via VPAC1R by a direct action on the mucosa. Although, even in the presence of the VPAC1R antagonist, there was a residual response to VIP suggesting involvement of another VIP receptor subtype, possibly VPAC2 as Vipr2 expression was found in the mucosa. The residual response may also be attributed to the efficacy of the antagonist at the concentration (1 μM) used or the duration of incubation before applying VIP. These were selected because a previous secretion study (Fang et al., 2006) reported this concentration and timing to be most effective. The same study reported that PG 97–269 (1 μM) suppressed or blocked 0.5 μM VIP responses, but, despite the lower concentration of VIP (50 nM) used in this study, PG 97–269 did not block the response. The reason for this
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discrepancy is unclear. Nonetheless, a higher concentration of antagonist or longer incubation might be more effective (Banks et al., 2005).
5.5 Enhanced secretory responses with repeated agonist exposure
Responses to VIP increased with repeated exposures before stabilizing. VIP may potentiate the cholinergic component via intracellular secondary messenger systems. Agents that increase cAMP levels, including VIP, enhance the secretory response to calcium-dependent secretagogues (Yajima et al., 1988), and the calcium-dependent cholinergic component of the secretory response to electrical stimulation (Carey et al., 1987) in guinea pig distal colon and ileum, respectively. There is also evidence of this in human T84 epithelial cell monolayers, where cholera toxin and forskolin (which both raise cAMP levels) significantly increase the response to exogenous ACh (Banks et al., 2004). However, details of the intracellular pathways and how the two systems may interact synergistically remain unknown.
5.6 Conclusion
Collectively, it was shown that cholinergic excitatory motor neurons innervating longitudinal smooth muscle and cholinergic secretomotor neurons express VPAC1R and are involved in mediating neurogenic contractile and secretory responses to exogenous VIP, respectively. However, the neural pathways involved are independent of nicotinic transmission. Identification of the physiological circuitry that leads to release of endogenous VIP on these two classes of cholinergic neurons will be the next step in understanding the place of VPAC1 receptors in the enteric neural circuitry controlling motility and secretion.
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CHAPTER 4: VPAC RECEPTOR SUBTYPES TUNE PURINERGIC NEURON-TO-GLIA COMMUNICATION IN
THE MURINE SUBMUCOSAL PLEXUS
1. ABSTRACT
The enteric nervous system (ENS) situated within the gastrointestinal tract is an intricate network of neurons and glia which together regulate intestinal function. The exact neuro- glial circuitry and the signaling molecules involved, are yet to be fully elucidated. Since vasoactive intestinal peptide (VIP) is one of the main neurotransmitters in the gut, this study aimed to unravel the role of VIP and its VPAC receptors within the circuitry of the submucosal plexus of mouse jejunum using calcium (Ca2+)-imaging. Local VIP application primarily induced neural Ca2+ transients that were inhibited by VPAC1- and VPAC2 antagonists (PG 99-269 and PG 99-465 respectively). Surprisingly, the VPAC2 antagonist revealed glial Ca2+ transients in response to VIP application, while the VPAC1 agonist ([K15, R16, L27]VIP(1-7)/GRF(8-27)) predominantly evoked glial responses. VPAC1- immunoreactivity was found in cholinergic submucosal neurons and nerve fibers, but not in glia. To confirm the presence of an intrinsic VIP/VPAC-initiated neuron-to-glia signaling pathway, trains of electrical stimuli were applied to fiber tracts to induce endogenous VIP- release. This evoked a delayed glial response in the presence of the VPAC2 antagonist. Additionally, VPAC1 agonist-evoked glial responses were inhibited by purinergic antagonists (PPADS and MRS2179). Hence, activating VPAC1 receptors on cholinergic submucosal neurons evokes purinergic signaling to enteric glia, while inhibiting VPAC2 receptors reveals VIP-evoked glial responses. Collectively, these results showed that neurally-released VIP can activate a secondary neuron expressing VPAC1 and/or VPAC2 receptors, to modulate purine-release onto glia. Thus, this study identified a component of
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an enteric neuron-glia circuit, fine-tuned by endogenous VIP via balanced activation of VPAC1- and VPAC2-mediated pathways.
2. INTRODUCTION
The regulation of intestinal functions is finely driven by neurons and glia that interact via neurotransmitters. Though it is now well established that enteric glia are involved in the control of intestinal function, the nature of the signaling mechanisms operating within the neuro-glial circuitry remain largely unknown. Evidence for communication between enteric neurons and glia originates from ultrastructural studies demonstrating that nerve endings terminating on enteric glia have vesicle-containing presynaptic specializations (Gabella, 1981). In addition, active synaptic release sites surrounding enteric glia have been identified (Vanden Berghe and Klingauf, 2007). Enteric glia in turn express a variety of receptors for neurotransmitters (Rühl, 2005), including purinergic receptors that have been established to be important in neuron-glia communication (Gomes et al., 2009; Gulbransen and Sharkey, 2009; Boesmans et al., 2013b).
Vasoactive intestinal peptide (VIP), one of the most abundant neuropeptides in the ENS, has long been considered a putative enteric neurotransmitter (Jessen et al., 1980). VIP- immunoreactive nerve terminals are present in both the submucosal (Foong et al., 2014) and myenteric plexuses (Sang and Young, 1996; Qu et al., 2008), and VIP depolarizes both submucosal (Mihara et al., 1985) and myenteric neurons (Palmer et al., 1987).
Using Ca2+ imaging in Wnt1-Cre;R26R-GCaMP3 mice, it was serendipitously found that selective activation of neural VPAC1 receptors induced purinergic signaling from cholinergic enteric neurons to glia. Components of an enteric neuron-glia circuit which may
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be regulated by VIP via VPAC1 and VPAC2 receptors and purinergic signaling was identified in the submucosal plexus of mouse small intestine.
3. MATERIALS AND METHODS
3.1 Tissue preparation and Ca2+ imaging in submucosal plexus
Wnt1-Cre;R26R-GCaMP3 mice of either sex were used for Ca2+ imaging (see Chapter 2, section 7). Wnt1-Cre;R26R-GCaMP3 mice express the fluorescent Ca2+ indicator GCaMP3 in all enteric neurons and glia (Danielian et al., 1998; Zariwala et al., 2012). A segment of jejunum (approximately 5 cm long) was collected and immersed in 95% oxygen/5% carbon dioxide-bubbled Krebs solution (in mM: 120.9 NaCl, 5.9 KCl, 1.2 MgCl2, 1.2 NaH2PO4,
14.4 NaHCO3, 11.5 glucose, and 2.5 CaCl2). The tissue was opened along the mesenteric border and pinned flat with mucosa up in a silicone elastomer-coated dish. Some tissues were immediately fixed in 4% formaldehyde/PBS for 45 min at room temperature for immunohistochemical staining. In others that were used for imaging, the submucosal plexus with the mucosa intact was separated from the underlying muscle layers by microdissection, and the tissue was stabilized by stretching the tissue with the plexus facing up over an inox ring before clamping it with a rubber O-ring (Vanden Berghe et al., 2002). Up to 5 ring preparations were obtained from each jejunal segment.
Two microscopy setups were used to image the ring preparations as described in Chapter 2 (section 7.2). Briefly, some preparations were imaged via a 20× (NA 1.0) water dipping lens on an upright Zeiss Examiner microscope (Carl Zeiss, Oberkochen, Germany) equipped with a CCD camera (Imago QE) and images (640 × 512) on this system were acquired at 2 Hz. Other preparations were imaged via a 20× (NA 0.5) water dipping lens on an upright Zeiss Axioscope microscope with a Zeiss AxioCam MRm camera, and images
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(278 × 278) were acquired at 1 Hz. Preparations were constantly superfused (1 ml/min) with 95% oxygen/5% carbon dioxide-gassed Krebs at room temperature.
Ganglia of interest were stimulated chemically or electrically. Agonists were applied to the preparations by pressure ejection (2 s duration; 9 psi) via a micropipette (tip diameter ~20 µm) positioned approximately 50 µm from the selected ganglion. For time control experiments, each agonist was applied 3 times separated by 5 minutes to ensure reproducible responses. For electrical stimulation experiments, trains of 300 μs pulses (2 s, 20 Hz) were applied by a focal stimulating electrode (tungsten; 50 µm diameter) on an internodal strand leading to the ganglion imaged. For time control experiments, ganglia were stimulated twice separated by 5 min.
Antagonist experiments consisted of 2 consecutive applications of agonist or 2 consecutive trains of electrical stimulation, with the antagonist superfused prior to the 2nd stimulation. Thus the 1st response is considered the control response. Antagonists were superfused via a local perfusion pipette. Each preparation was only exposed to a single antagonist once and to only one type of agonist. At least 3 animals were examined for each set of experiments.
After calcium imaging experiments, selected preparations were fixed in 4% formaldehyde/PBS for 45 min at room temperature for post hoc immunohistochemistry.
3.2 Drugs
Agonists used included VIP (human, mouse, rat; Bachem, Bubendorf, Switzerland), VPAC1 agonist, [K15, R16, L27] VIP (1-2)/GRF(8-27) and VPAC2 agonist, BAY 55-9837 (both synthesized by Mimotopes, Clayton, Victoria, Australia), and the P2Y1 agonist 2-
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methyl-thio-ADP (2MeSADP) (Sigma Aldrich, Castle Hill, New South Wales, Australia). Superfusion of 1 µM VIP depolarised myenteric neurons (Palmer et al., 1987). Hence, 100 µM VIP was used to fill the micropipette for local application by pressure ejection, as the solution may be diluted up to approximately 100-fold in superfused preparations (Liu et al., 1997). The VPAC1 agonist [K15, R16, L27] VIP (1-2)/GRF(8-27) and the VPAC2 agonist BAY 55-9837 were also used at 100 µM, as VIP and [K15, R16, L27] VIP (1-2)/GRF(8- 27) have similar potencies against the VPAC1 receptor (Gourlet et al., 1997b), while BAY 55-9837 has shown to be effective on hypothalamic neurons when superfused at 1 – 10 µM (Hermes et al., 2009; Pantazopoulos et al., 2010).
Selective VIP antagonists used include the VPAC1 antagonist PG 97–269, and the VPAC2 antagonist, PG 99-465 (both from Mimotopes). PG 97–269 has been previously demonstrated to inhibit VIP-evoked intestinal secretion at 1 µM (Fung et al., 2014), while PG 99-465 inhibits VIP-evoked responses in hippocampal neurons at 100 nM – 10 µM (Pakhotin et al., 2005; Cunha-Reis et al., 2014). Accordingly, both VPAC antagonists were superfused at a concentration of 1 µM. Other antagonists include tetrodotoxin, TTX (1 µM; Acros, Geel, Belgium); hexamethonium bromide (200 µM; Sigma; Bornem, Belgium), pyridoxal-phosphate-6-azophenyl-2',4'-disulfonic acid, PPADS (30 µM; Sigma), and MRS2179 (10 µM; Tocris, Bristol, UK). The VPAC1R and VPAC2R antagonists were applied 5 min prior to agonist application as they have been shown to be most effective with short incubations (Pakhotin et al., 2005; Fang et al., 2006). TTX, hexamethonium, PPADS, and MRS2179 were superfused for 10 min.
All drugs were prepared as stock solutions using water, except for TTX which was prepared in citrate buffer. TTX, 2MeSADP, and MRS2179 were stored at 4°C. Hexamethonium bromide, PPADS, VIP, [K15, R16, L27] VIP (1-2)/GRF(8-27), BAY 55- 9837, PG 97-269, and PG 99-465 were stored as aliquots at −20°C. Agonists and antagonists were diluted in Krebs to achieve the desired final concentration.
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3.3 Analysis
Only ganglia directly adjacent to the micropipette were considered for analysis. Ca2+ imaging analyses were performed with custom-written routines in Igor Pro (Wavematrics, Lake Oswego, Oregon, USA) (refer to Chapter 2, section 7.3). Regions of interest were 2+ 2+ drawn to calculate the average [Ca ]i signal intensity. [Ca ]i peaks were calculated for 2+ each response, with the peak amplitude taken as the maximum increase in [Ca ]i from nd baseline (∆Fi/F0). For time controls, the ΔFi/Fo of the 2 agonist exposure is presented as a st percentage of the 1 (% ΔFi/Fo). Similarly, the ΔFi/Fo evoked in the presence of antagonists is presented as a percentage of the 1st control agonist response. Responses that were observed with the 2nd agonist exposure, but not the 1st were not considered in this analysis. The total number of cells responding to each agonist exposure was also counted. The total number of cells responding to the 2nd agonist application was then similarly normalized to that of the 1st control response and presented as a percentage of the control.
3.4 Immunohistochemistry
Immediately-fixed jejunal segments from mice of a C57Bl6 background including Wnt1- Cre;R26R-GCaMP3 mice, and fixed submucosal ring preparations, were used for immunohistochemical staining. The mucosa and submucosa of immediately-fixed tissues were separated from the underlying muscle layers by microdissection. The mucosa of immediately-fixed preparations and fixed submucosal ring preparations was then removed and the preparations were incubated with a blocking buffer containing 4% donkey serum (Merck Millipore, Overijse, Belgium) and 0.5% triton X-100 (Sigma) in PBS overnight at 4°C. Preparations were subsequently incubated in primary antisera (Table 4.1) for 24 - 48 h at 4°C, washed in PBS (3 x 10 min) and then incubated in secondary antisera (Table 4.2) for 2 h at room temperature. Primary and secondary antisera were diluted in the blocking buffer. Preparations were washed in PBS (3 x 10 min) before mounting on slides with
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Citifluor (Citifluor Ltd., Leicester, UK) or Dakocytomation fluorescent mounting medium (Carpinteria, California, USA).
Fluorescently labelled preparations were viewed under an epifluorescence microscope (BX 41 Olympus, Olympus, Aartselaar, Belgium) equipped with UMNUAUV, U-MWIBA3 and U-MWIY2 filtercubes for visualising blue, green and red probes, respectively. Images were acquired with an XM10 (Olympus) camera using Cell^F software. Pictures were adjusted for contrast and brightness before overlay and quantification. Confocal images were recorded using a Zeiss LSM780 confocal microscope (Cell Imaging Core, KU Leuven, Belgium) or Zeiss Pascal confocal microscope (Biological Optical Microscopy Platform, The University of Melbourne, Australia).
3.5 Data and statistics
Data are presented as mean ± the standard error of the mean (SEM), and ‘n’ number of cells examined unless stated otherwise. One-way analysis of variance (ANOVA) followed by Dunnett’s post hoc test (where appropriate) were conducted to determine statistical significance, unless specified otherwise. P < 0.05 was considered significant. Analyses were performed using GraphPad Prism 5.0 (GraphPad softwares, San Diego, California, USA).
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Table 4.1. Primary antibodies used for immunohistochemistry.
Primary Antibodies Host Dilution Source
VPAC1R Rabbit 1:1000 Pierce Biotechnology
ChAT Goat 1:500 Chemicon
Peripherin (C-19) Goat 1:500 Santa Cruz Biotechnologies
GFAP Chicken 1:5000 Abcam
GFAP Guinea pig 1:1000 Synaptic Systems
Tyrosine Hydroxylase Sheep 1:1000 Millipore
VAChT Guinea pig 1:100 Millipore
CGRP Goat 1:1000 AbD Serotec
Hu Human 1:5000 Gift from Miles Epstein
Abbreviations: VPAC1R, vasoactive intestinal peptide receptor 1; ChAT, choline acetyltransferase; GFAP, glial fibrillary acidic protein; VAChT, vesicular acetylcholine transporter; CGRP, calcitonin gene-related peptide
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Table 4.2. Secondary antibodies used for immunohistochemistry.
Secondary Antibodies Host Dilution Source
Anti-rabbit AMCA Donkey 1:250 Jackson Immuno Labs
Anti-rabbit AF594 Donkey 1:400 Molecular Probes
Anti-goat AF594 Donkey 1:1000 Molecular Probes
Anti-sheep AF647 Donkey 1:500 Molecular Probes
Anti-sheep AF488 Donkey 1:400 Molecular Probes
Anti-chicken AF594 Donkey 1:1000 Molecular Probes
Anti-guinea pig FITC Donkey 1:100 Millipore
Anti-human AF647 Donkey 1:500 Jackson Immuno Labs
Anti-human AF488 Donkey 1:800 Jackson Immuno Labs
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4. RESULTS
2+ 4.1 VIP exclusively evokes neuronal [Ca ]i transients in submucosal plexus
Local VIP (100 µM) application by pressure ejection evoked transient increases in 2+ 2+ intracellular Ca concentration ([Ca ]i) exclusively in neurons (Figure 4.1A); 44 ± 5% of 2+ neurons per ganglion responded with consistent [Ca ]i rises (∆Fi/F0: 0.28 ± 0.02, n = 103; Figure 4.1B). Ganglia imaged in these experiments contained on average 9.3 ± 0.5 neurons. Post hoc immunohistochemistry was performed on some preparations to characterize the types of neurons which responded to VIP (100 µM; Figure 4.1C). Both ChAT+ neurons and - 2+ ChAT neurons were responsive to VIP, but the characteristics of the [Ca ]i transients observed in these two neuronal subtypes were distinctly different (Figure 4.1D). VIP- + 2+ responsive ChAT neurons displayed [Ca ]i transients with a duration (at t50%; 50% of the peak amplitude) of 11.8 ± 0.8 s (n = 24), while ChAT- neurons displayed slower responses with a significantly longer duration of 15.3 ± 1.0 s (n = 42) (unpaired t-test, P = 0.016).
4.2 Inhibiting VPAC2 receptors reveals a VIP-evoked glial response
Next, the relative contributions of VPAC1 and VPAC2 receptors in mediating VIP-induced 2+ [Ca ]i transients using the selective VPAC1 antagonist PG 97-269 (Banks et al., 2005) and the VPAC2 antagonist PG 99-465 (Moreno et al., 2000; Dickson et al., 2006) were examined. The VIP (100 µM)-evoked neuronal response amplitudes were significantly reduced in the presence of the VPAC1 antagonist PG 97-269 (1 µM) (to 22 ± 5% of the control; one-way ANOVA and Dunnett’s test, P < 0.0001; n = 47; Figure 4.2A), but the number of neurons responding did not change (Figure 4.2B). The VPAC2 antagonist PG 99-465 (1 µM) effectively reduced both the amplitude of the VIP-evoked response (to 22 ± 9% of the control; one-way ANOVA and Dunnett’s test, P < 0.0001; n = 30; Figure 4.2A) and the number of responding neurons per ganglion (to 36 ± 18% of the control; one-way
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ANOVA and Dunnett’s test, P < 0.05; Figure 4.2B). These data indicate that the VIP- 2+ evoked [Ca ]i transients in neurons involve both VPAC1 and VPAC2 receptor activation. Interestingly, in 6/7 preparations, glial responses to VIP were uncovered in the presence of 2+ the VPAC2 antagonist, where a [Ca ]i rise was observed in 1.6 ± 0.4 glial cells per ganglion (n = 11; Figure 4.2C).
The VIP-evoked response was also nearly abolished by the Na+-channel blocker TTX (1 µM), which significantly reduced the amplitude of the response (to 15 ± 10% of the control; one-way ANOVA and Dunnett’s test, P < 0.0001; n = 40; Figure 4.2A) and the number of responding neurons per ganglion (to 16 ± 7% of the control; one-way ANOVA 2+ and Dunnett’s test, P < 0.001; Figure 4.2B). This suggests that the majority of [Ca ]i transients observed were a consequence of neuronal activation. Furthermore, reminiscent of the effect of the VPAC2 antagonist, TTX uncovered VIP-evoked glial responses, with 1.9 ± 0.6 glial cells responding per ganglion (n = 15; Figure 4.2C).
Blocking a major form of fast excitatory neurotransmission within the ENS, using the nicotinic receptor antagonist hexamethonium (200 µM), also reduced the amplitude of VIP- evoked responses (to 32 ± 7% of the control; one-way ANOVA and Dunnett’s test, P < 0.001; n = 24; Figure 4.2A). In contrast with TTX and the VPAC2 antagonist, hexamethonium did not reveal VIP-evoked glial responses.
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2+ Figure 4.1. VIP-induced [Ca ]i transients in enteric neurons and glia.
A. Number of submucosal neurons and glia responding per ganglion to 3 repeated local spritz-applications (2 s duration) of VIP (100 µM) separated by 5 min in time control experiments (n = 18 preparations). B. Maximum amplitudes of VIP-evoked neuronal 2+ [Ca ]i transients were reproducible in time control experiments. C. Fluorescence images of + - 2+ ChAT (red arrow) and ChAT submucosal neurons (green arrow) displaying a [Ca ]i rise in response to VIP over time. Scale bar = 20 µm. Corresponding ChAT immunolabelling was used to identify ChAT+ and ChAT- neurons. D. Corresponding traces of VIP-induced 2+ + - [Ca ]i responses in selected ChAT and ChAT neurons. The black arrow in the graph indicates VIP application.
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Figure 4.2. VIP (100 µM)-evoked glial and neuronal responses in the presence of various antagonists.
A. The selective VPAC1 antagonist (PG 97-269; 1 µM; n = 47 neurons), the VPAC2 antagonist (PG 99-465; 1 µM; n = 30 neurons), tetrodotoxin (TTX; 1 µM; n = 40 neurons), and the nicotinic antagonist hexamethonium (Hex.; 200 µM; n = 24 neurons) all 2+ significantly inhibited the peak amplitude of the VIP-evoked neuronal [Ca ]i transients (one-way ANOVA and Dunnett’s test, ***P < 0.001, ****P < 0.0001). B. The VPAC2 antagonist also inhibited the number of neurons responding to VIP compared to control (one-way ANOVA and Dunnett’s test, *P < 0.05; n = 7 preparations). TTX (1 µM) significantly inhibited the number of neurons responding to VIP and near abolished the 2+ neuronal response transients (***P < 0.001; n = 8 preparations).. VIP-induced glial [Ca ]i transients were revealed by antagonists. The number of submucosal glia responding to a local spritz-application of VIP (100 µM) in the presence of C. the VPAC2 antagonist PG 99-465 (1 µM; n = 7 preparations) or tetrodotoxin (TTX; Na+ channel blocker; 1 µM; n = 8 preparations) was significantly increased compared to control conditions (two-way ANOVA and Sidak’s test, *P < 0.05, **P < 0.01). Glial responses were not observed in time control experiments (n = 25 preparations). D. Delayed glial responses to a train of electrical stimuli (300 µs pulses, 2 s, 2 Hz) applied by a stimulating electrode placed on an internodal strand were revealed by PG 99-465 (1 µM; n = 26 ganglia, paired t-test, **P = 0.0043).
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2+ 4.3 Selective VPAC1 activation predominantly evokes [Ca ]i transients in enteric glia
2+ As inhibiting VPAC2 receptors unveiled glial [Ca ]i transients in response to VIP, whether the VIP-induced glial response is mediated via VPAC1 receptors was investigated. Local spritz application of [K15, R16, L27] VIP (1-2)/GRF(8-27) (VPAC1 agonist; 100 µM) 2+ (Gourlet et al., 1997b) to submucosal ganglia predominantly evoked [Ca ]i responses in glial cells (Figure 4.3A). By contrast, BAY 55-9837 (VPAC2-selective agonist; 100 µM) 2+ did not evoke reproducible [Ca ]i responses in neurons or glia. The VPAC1 agonist 2+ evoked [Ca ]i transients in 3.5 ± 0.4 glia per ganglion (Figure 4.3A) with a peak amplitude
(∆Fi/F0) of 0.42 ± 0.03 (n = 108; Figure 4.3B). With post hoc immunohistochemistry for glial fibrillary acidic protein (GFAP), the fibers activated in response to the VPAC1 agonist were confirmed to be glial (Figure 4.3C – E). The types of glial cells responding were classified based on their location and morphology (Boesmans et al., 2015b). Predominantly type I enteric glial cells responded to VPAC1 agonist application: 2.1 ± 0.3 type I vs. 1.4 ± 2+ 0.2 type II cells displayed [Ca ]i transients per ganglion (paired t-test, P = 0.03; n = 31 ganglia).
2+ The VPAC1 agonist also evoked some neuronal [Ca ]i responses (∆Fi/F0: 0.65 ± 0.1; n = 38), where 13 ± 3% of neurons responded per ganglion. In this set of experiments, the mean number of neurons contained per ganglion imaged was 10.6 ± 0.6. Examining the time course of the VPAC1 agonist-evoked responses, some neuronal responses preceded glial activation and, interestingly, some neuronal responses were observed following the onset of 2+ 2+ a [Ca ]i rise in glia. Neurons that responded before glia displayed [Ca ]i transients with a latency of 4.3 ± 1.2 s (n = 10 neurons). Glial responses showed a latency of 14 ± 1.0 s (n = 108 glia) and neurons which responded following glial activation had a latency of 19 ± 2 s (n = 28 neurons).
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2+ Figure 4.3. VPAC1-induced [Ca ]i transients in enteric neurons and glia.
A. Number of submucosal neurons and glia responding to 3 repeated local spritz- applications of VPAC1 agonist (100 µM) separated by 5 min in time control experiments (n 2+ = 22 preparations). B. Maximum amplitudes of VPAC1 agonist-evoked glial [Ca ]i transients were reproducible over time control experiments. C. Glial fibers in a submucosal ganglion responding to VPAC1 agonist over time, as indicated by arrows. Scale bar = 20 2+ µm. D. VPAC1 agonist-induced [Ca ]i responses of selected glial fibers as indicated by colour-coded arrows in C. The black arrow in the graph marks the application of VPAC1- agonist. E. Responding glial fibers identified by GFAP immunofluorescent labelling.
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4.4 The VPAC1 receptor is expressed on cholinergic submucosal neurons and nerve fibers
The localization of the VPAC1 receptor was then examined using immunohistochemistry (Figure 4.4). VPAC1 receptor expression was specifically neural, with the antisera labeling a subset of submucosal neurons and neuronal, but not glial, processes. VPAC1 receptor- immunoreactivity was observed in 12 ± 2% Hu+ cells and these were predominantly ChAT+ (93 ± 4%; n = 284 neurons; 3 animals; Figure 4.4A – D). Some background nuclear labelling was observed in neurons, but this staining was not considered in the analysis. VPAC1 receptor staining also colocalized with peripherin, a label for neuronal fibers (Figure 4.4E – H; n = 3 animals). However, VPAC1 receptors did not colocalize with GFAP+ glial processes (n = 3 animals; Figure 4.4I – L). Since VPAC1 receptors were found to be expressed on some ChAT+ neurons, whether VPAC1 receptors may also be expressed on varicosities containing vesicular acetyltransferase (VAChT) was then examined. However, there was no apparent overlap between VPAC1 and VAChT (n = 2 animals; Figure 4.4M – O).
As it has been reported that enteric glia receive innervation from sympathetic fibers in guinea pig myenteric plexus (Gulbransen et al., 2010), whether VPAC1 receptors may also be expressed on sympathetic adrenergic nerve fibers was addressed. However, no colocalization between tyrosine hydroxylase (TH; labels adrenergic nerves) and VPAC1 receptor expression was observed (n = 2 animals; Figure 4.5A – C). In addition to TH nerve fibers, many submucosal neurons of mouse ileum are surrounded by calcitonin gene-related peptide (CGRP)-immunoreactive varicosities (Mongardi Fantaguzzi et al., 2009). Some colocalization of VPAC1 receptors with CGRP-immunoreactive varicosities was observed (n = 2 animals; Figure 4.5D – F).
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4.5 Inhibiting VPAC2 receptors reveals delayed glial responses to electrical stimulation
To confirm the presence of a VIP/VPAC neuron-to-glia signaling pathway, as suggested by the pharmacological approaches, fiber tracts were stimulated electrically to release endogenously-produced VIP. A similar effect was observed in this set of experiments, 2+ where trains of stimuli evoked neuronal [Ca ]i transients (∆Fi/F0: 0.80 ± 0.08; n = 86) that were unaffected by the VPAC2 antagonist PG 99-465 (1 µM; n = 115). However, in the presence of PG 99-465, delayed glial responses were observed following electrical stimulation in 7/13 preparations (0.4 ± 0.1 glia responded per ganglion) (Figure 4.2D). These delayed glial responses had a latency of 20.5 ± 2.9 s (n = 10) and were not observed in time controls.
4.6 VPAC1 agonist-evoked glial activation involves a TTX-insensitive mechanism
PG 97-269 (VPAC1 antagonist; 1 µM) was used to test the specificity of the VPAC1 agonist (100 µM), and it significantly reduced the number of responding glia per ganglion (to 58 ± 11% of the control; one-way ANOVA and Dunnett’s test, P < 0.05; Figure 4.6B), 2+ as well as the peak amplitude of the glial [Ca ]i transients (to 19 ± 5% of the control; one- way ANOVA and Dunnett’s test, P < 0.01; n = 50; Figure 4.6A). The VPAC1 agonist- induced glial response was not affected by TTX (1 µM; n = 24; Figure 4.6). Given that VPAC1 receptors are not expressed on glial fibers, the VPAC1 agonist-evoked glial response must be mediated by a neural, but TTX-insensitive, mechanism.
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Figure 4.4. Localizing VPAC1 receptor (VPAC1R)-expression in the submucosal plexus of mouse jejunum.
A – D. Confocal micrographs demonstrating that VPAC1R-immunoreactive cell bodies were cholinergic, as marked by B. ChAT and C. Hu staining and indicated by the arrow. E – H. VPAC1R expressing fibers also overlapped with peripherin+ nerve fibers (indicated by filled arrowheads). I – L. However, VPAC1R did not colocalize with the GFAP+ glial fibers (open arrowheads). M – O. VPAC1R- and VAChT-immunoreactivity also did not overlap (open arrowheads). Scale bars = 20 µm.
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Figure 4.5. Confocal images of VPAC1 receptor (VPAC1R)-immunoreactive nerve fibers and varicosities in the submucosal plexus of mouse jejunum taken at a single optical plane.
A – C. VPAC1R did not colocalize with tyrosine hydroxylase (TH), as indicated by open arrowheads. D – F. Confocal images of VPAC1R- and calcitonin related-gene peptide (CGRP)-immunoreactive varicosities in a submucosal ganglion. Some overlap between VPAC1R- and CGRP-labelling was observed (filled arrowhead). However, not all VPAC1R+ varicosities were CGRP+ and vice versa (open arrowhead). Scale bars = 20 µm.
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2+ Figure 4.6. VPAC1 agonist (100 µM)-evoked glial and neuronal [Ca ]i responses in the presence of antagonists.
A. The peak amplitude of the VPAC1 agonist-evoked response was significantly inhibited by the selective VPAC1 antagonist (PG 97-269; 1 µM; one-way ANOVA and Dunnett’s test, **P < 0.01; n = 50 glia), but was not affected by TTX (1 µM; n = 24 glia). The P2 antagonist PPADS (30 µM; n = 31 glia) and the selective P2Y1 antagonist MRS2179 (10 µM; n = 20 glia) were also effective in inhibiting the VPAC1 agonist response amplitude (one-way ANOVA and Dunnett’s test, *P < 0.05, **P < 0.01). B. Only the VPAC1 antagonist significantly reduced the number of glia responding to the VPAC1 agonist (one- way ANOVA and Dunnett’s test, *P < 0.05; n = 13 preparations).
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2+ 4.7 VPAC1 receptor activation stimulates glial [Ca ]i responses via purinergic signaling
As purinergic signaling is known to be important in enteric neuron-glia communication (Gomes et al., 2009; Gulbransen and Sharkey, 2009), the effect of purinergic antagonists on the VPAC1 agonist-stimulated glial responses was examined. The P2 receptor antagonist PPADS (30 µM), significantly reduced the peak amplitude of the VPAC1 agonist-induced 2+ glial [Ca ]i response to 7 ± 3% of the control (one-way ANOVA and Dunnett’s test, P < 0.01; n = 31; Figure 4.6A). The P2Y1 receptor is a P2 receptor associated with enteric glial activation (Gomes et al., 2009; Gulbransen et al., 2012). It was found that the P2Y1- selective antagonist MRS2179 (10 µM) also significantly inhibited the amplitude of the VPAC1 agonist-induced glial response (to 10 ± 5% of the control; one-way ANOVA and Dunnett’s test, P < 0.05; n = 20; Figure 4.6A).
To further investigate the role of P2Y1 receptors, the response to spritz-applied 2MeSADP (100 µM; P2Y1-specific agonist) was examined. Consistent with observations in mouse myenteric plexus (Brown et al., 2016), P2Y1 receptor stimulation with 2MeSADP 2+ primarily evoked [Ca ]i transients in glial cells in the submucosal plexus. 2MeSADP evoked responses in 4.8 ± 0.4 glial cells per ganglion (∆Fi/F0: 0.16 ± 0.02; n = 43). The specificity of 2MeSADP as a P2Y1 agonist was assessed and it was found that the peak amplitude of the response was significantly reduced by MRS2179 (10 µM) (to 21 ± 6% of the control; one-way ANOVA and Dunnett’s test, P < 0.05; n = 29; Figure 4.7A). TTX (1 µM) did not inhibit the 2MeSADP-evoked glial response, rather the number of glia responding was increased (to 122 ± 27% of control; one-way ANOVA and Dunnett’s test, P < 0.05; Figure 4.7B).
2+ The [Ca ]i responses to 2MeSADP (100 µM) and VPAC1 agonist (100 µM) were compared by sequentially applying these agonists to same ganglion, with each application 2+ separated by a 5 min washout. 2MeSADP evoked [Ca ]i responses in significantly more
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glia per ganglion than the VPAC1 agonist (4.6 ± 0.4 vs. 2.8 ± 0.5 respectively; paired t-test, P = 0.027; n = 10 ganglia), and responses were unaffected by the order of agonist application. Notably, some glial cells responded to both agonists (Figure 4.7C, D), but 2+ 2MeSADP induced higher [Ca ]i peak amplitudes (∆Fi/F0: 0.23 ± 0.02 vs. 0.18 ± 0.02 respectively; paired t-test, P = 0.026; n = 24). Moreover, the time course of the responses to the two agonists differed drastically (Figure 4.7E, F). The 2MeSADP-evoked glial response was rapid and short-lived with a latency of 2.8 ± 0.2 s, while the VPAC1 agonist glial response exhibited a significant delay with a latency of 16.3 ± 2.1 s (paired t-test, P < 0.0001; n = 24).
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2+ Figure 4.7. P2Y1 agonist 2MeSADP (100 µM)-evoked glial [Ca ]i responses.
A. The P2Y1 antagonist MRS2179 (10 µM) significantly inhibited the amplitude of glial cells responding to 2MeSADP (one-way ANOVA and Dunnett’s test, *P < 0.05; n = 43 glia). B. The number of glia responding to 2MeSADP was increased in the presence of TTX (1 µM) compared to control (one-way ANOVA and Dunnett’s test, *P < 0.05; n = 6 2+ preparations). C – F. Some glial cells displayed [Ca ]i transients in response to both 2MeSADP (100 µM) and VPAC1 agonist (100 µM), as indicated by arrows. However, the 2+ latency of the 2MeSADP-evoked glial [Ca ]i responses was significantly shorter than that of the VPAC1 agonist. The black arrows in the graphs mark the application of agonist.
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5. DISCUSSION
In this study, a hitherto unknown purinergic neuro-glia interaction initiated by selective VPAC1-activation was identified in the mouse submucosal plexus. VIP and the VPAC1 2+ agonist evoked distinctly different [Ca ]i responses, where VIP exclusively evoked neuronal responses, while the VPAC1 agonist primarily evoked glial responses. These data indicate that both VIP and the VPAC1 agonist initiate responses through neuronal activation. This is in agreement with previous observations showing that VIP depolarizes enteric neurons (Mihara et al., 1985; Palmer et al., 1987). This study is the first to demonstrate that VIP, through neuronal VPAC1 receptors, initiates a sequential activation of neurons and glia, which is in part inhibited by VPAC2 receptor activation.
5.1 VPAC2 receptors inhibit VIP-evoked glial responses
2+ In neurons, both VPAC antagonists inhibited the VIP-evoked increase in [Ca ]i, which indicates that both VPAC receptor subtypes are involved in mediating this response. While 2+ VIP alone did not evoke [Ca ]i transients in glia, glial responses were revealed with VPAC2 receptor antagonism and by blocking TTX-sensitive Na+ channels. The 2+ discrepancy in [Ca ]i responses to VIP and the VPAC1 agonist may be accounted for by a VPAC2 receptor-mediated and TTX-sensitive inhibitory pathway which suppresses VPAC1-mediated glial responses. Interestingly, delayed glial responses were also observed following trains of electrical stimuli when VPAC2 receptors were inhibited, suggesting that VPAC2 receptors are involved in inhibiting glial activation by endogenously released mediators, presumably VIP.
Blocking nicotinic transmission, a major form of excitatory neurotransmission in the ENS (Galligan and North, 2004; Gwynne and Bornstein, 2007b; Foong et al., 2015), with
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hexamethonium also reduced the number of neurons responding to VIP. However, unlike TTX and the VPAC2 antagonist, hexamethonium did not reveal responses in glial cells. Thus, nicotinic neurotransmission and the neurons responding secondarily to VIP do not appear to be involved in this neuron-glia interaction. This may suggest that the neural VPAC2-mediated inhibitory pathway is monosynaptic. Another possibility is that VPAC1 and VPAC2 may be co-expressed on the same neuron, and VPAC2-activation antagonizes the VPAC1-initiated signal.
There are some inconsistencies regarding the role of the VPAC2 receptor as the VPAC2 2+ agonist alone did not evoke any [Ca ]i signals in neurons or glia. This may be because the 2+ VPAC2 agonist does not induce a robust [Ca ]i rise, but preferentially activates the cAMP secondary messenger system (Dickson et al., 2006; Langer, 2012). Alternatively, it is possible that other receptors are activated by VIP to suppress the glial component of the VPAC1 receptor-mediated response, such as VPAC1 and VPAC2 receptor splice variants (Dickson and Finlayson, 2009). VPAC receptor oligomerization and interaction of VPAC receptors with accessory proteins (Couvineau and Laburthe, 2012) have also been reported, although the functional roles of these are unclear.
5.2 Activation of VPAC1 receptors on cholinergic neurons stimulates glial responses
VPAC1 receptor expression on astrocytes has been shown in the central nervous system (CNS) (Joo et al., 2004), but its expression in the ENS was found to be exclusively neural. Yet, the VPAC1 agonist-induced glial response was TTX-insensitive. The glial response may be evoked by activating VPAC1 receptors on nerve fibers in contact with glia. As in the guinea pig, VPAC1 receptor expression in mouse jejunum was predominantly localized to cholinergic submucosal neurons (Fung et al., 2014), but VPAC1-immunoreactive varicosities did not overlap with VAChT in this mouse study. Perhaps this is not surprising, as it has been previously shown that some nerve terminals of ChAT-containing cell bodies
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may not necessarily be ‘cholinergic’ in that they do not contain VAChT (Sharrad et al., 2013). It was also considered whether VPAC1 receptors may be expressed on sympathetic nerve fibers as these innervate glia in the guinea pig myenteric plexus (Gulbransen et al., 2010). However, VPAC1 receptors were not expressed on TH-containing fibers. Instead, partial colocalization between CGRP- and VPAC1 receptors in varicosities was observed. These CGRP+ nerve terminals may either originate from extrinsic primary afferents (De Jonge et al., 2003), myenteric intrinsic sensory neurons (Qu et al., 2008), or submucosal CGRP neurons (Mongardi Fantaguzzi et al., 2009). However, the specific subtype/s expressing VPAC1 receptors, and whether extrinsic inputs are involved in this neuron to glia signaling pathway, require further investigation.
5.3 VPAC1 receptor activation stimulates glial responses via purine release
TTX-insensitive P2X7-mediated enteric neuron-glia signaling has been previously reported in the mouse colon myenteric plexus (Gulbransen et al., 2012), although the mechanism is still unclear. Purines acting on P2 receptors is one of the major signaling pathways involved in neuron-glia transmission (Gulbransen and Sharkey, 2009; Boesmans et al., 2013a) and the most extensively studied. In accordance with these findings, the VPAC1 agonist-evoked glial response was largely inhibited by the P2 antagonist PPADS and also partially inhibited by the P2Y1-selective antagonist MRS2179. In further examining the P2Y1 component of this response, it was observed that glial cells which responded to the VPAC1 agonist could also be activated by the P2Y1 agonist 2MeSADP. Notably, the glial responses to the VPAC1 agonist occurred with a significant delay compared to those evoked by 2MeSADP. This is consistent with the notion that VPAC1 receptor activation first stimulates a neuron or at least a neuronal release site, which then activates glia. Furthermore, based on the timing of responses, it is most likely that P2Y1 receptors are expressed on glial cells and that the 2MeSADP response results from a direct stimulation of these receptors. This is also compatible with the data showing that the 2MeSADP response was not inhibited by TTX. In fact, the number of glia responding to 2MeSADP was
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increased by TTX, suggesting either a 2MeSADP-mediated or an ongoing neural suppression of Ca2+ signaling activity in glia. While studies in guinea pig have shown that functional P2Y1 receptors are expressed in submucosal and myenteric neurons (Hu et al., 2003; Gao et al., 2006), and 2MeSADP (10 µM) has been reported to evoke Ca2+ responses in rat submucosal neurons (Christofi et al., 2004), in these experiments 2MeSADP 2+ predominantly evoked [Ca ]i responses in glia. This may be due to species differences as these findings are in agreement with other studies in mice, demonstrating that the activation of P2Y1 receptors primarily stimulates enteric glia, but not neurons (Gomes et al., 2009; Gulbransen et al., 2012). Taken together, these data are in support of an intermediate neuronal component between VPAC1 receptor activation and a subsequent P2Y1-mediated glial response, although other purinergic receptors may also be involved.
Notably, there were a number of neurons which responded following glial activation. It is 2+ tempting to speculate that the activated glia can induce [Ca ]i transients in neurons to modulate activity within the enteric circuitry as shown in the CNS (Hansson and Ronnnback, 2003). While there is no evidence for a physiological equivalent of the VPAC1 2+ agonist, and VIP under control conditions does not stimulate glial [Ca ]i transients, it is conceivable that imbalances in the network may lead to the VPAC1 receptor-mediated pathway being preferentially activated. Alternatively, selective VPAC1-activation may be possible in a physiological setting with localized VIP release, if VPAC1 and VPAC2 receptors are differentially expressed. Further, the observation of delayed glial responses to electrical stimulation revealed by inhibiting VPAC2 receptors suggests that these pathways may be activated by endogenous VIP to modulate signals from neurons to glia. Nonetheless, how these pathways may be functionally involved in physiology and disease remains to be elucidated.
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5.4 Conclusion
These results showed that activating VPAC1 receptors on cholinergic submucosal neurons 2+ stimulates purine release via a TTX-resistant mechanism, to then evoke [Ca ]i transients in glia. Additionally, inhibiting VPAC2 receptors uncovered glial responses. The presence of an intrinsic VIP/VPAC neuron-to-glia signaling pathway was confirmed by electrically stimulating fiber tracts to evoke endogenous VIP release. Collectively, it was demonstrated that stimulating neurally-released VIP can activate a secondary neuron, which may express VPAC1 and/or VPAC2 receptors, to modulate purine release onto glial cells. Thus, this study identified a component of the enteric neuron-glia circuit that is regulated by VIP via a balanced activation of excitatory VPAC1- and inhibitory VPAC2-mediated pathways.
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CHAPTER 5: CHOLERA TOXIN INDUCES SUSTAINED EXCITABILITY OF CHOLINERGIC SUBMUCOSAL NEURONS IN MOUSE ILEUM
1. ABSTRACT
Cholera-induced hypersecretion causes dehydration and death if untreated. Cholera toxin (CT) partly acts via the enteric nervous system (ENS), but the circuitry involved is unclear. To examine this, I incubated mouse ileal loops with CT (12.5 µg/ml) or saline (control) for 3.5 h in vivo, then studied the tissue in vitro. Incubated ileal segments were used to measure electrogenic ion secretion in Ussing chambers and assessed for c-Fos and pCREB (activity- dependent markers) expression. Ileal loops from Wnt1-Cre;R26R-GCaMP3 mice which express a Ca2+ indicator in the ENS were used to examine circuit excitability with Ca2+ imaging. Recorded neurons were identified by post-hoc immunohistochemistry. CT-treated preparations showed a higher basal secretion than controls. Neurogenic responses to DMPP (nicotinic agonist) and veratridine (activates voltage-gated Na+ channels) were unchanged. In the submucosal plexus, neuronal (marked by Hu) pCREB expression was significantly higher in CT-treated tissues than controls. More spontaneously active neurons were observed in CT-incubated tissues (31/160 neurons) than controls (4/149 neurons). Furthermore, spontaneously active neurons were cholinergic (ChAT+). DMPP- (10 µM) and electrically evoked (single pulse and train of 20 pulses) responses were also increased in cholinergic neurons. In the myenteric plexus, CT increased neuronal c-Fos expression compared to controls, but spontaneous activity was depressed. Overall, DMPP- and electrically evoked Ca2+ responses were also reduced. Thus, CT-incubation induced hypersecretion by acting directly on the mucosa and by increasing cholinergic secretomotor neuron excitability.
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2. INTRODUCTION
Cholera toxin (CT), produced by Vibrio cholera, causes severe diarrhoea and cholera remains a major health issue in developing countries. It is well established that CT induces hypersecretion in various animal models in vivo (De and Chatterje, 1953; Basu and Pickett, 1969) and in vitro (Field et al., 1972; Carey and Cooke, 1986; Burleigh and Borman, 1997). Most in vitro studies suggest that CT primarily stimulates hypersecretion by direct effects on the mucosal epithelium (Field et al., 1972; Burleigh and Borman, 1997). Yet, in vivo studies indicate the involvement of the enteric nervous system (ENS) (Cassuto et al., 1982a; Cassuto et al., 1983) and it has been proposed that the activation of neural secretomotor pathways is the major mechanism of action. The discrepancies between these studies are yet to be resolved.
Evidence for ENS involvement in CT-induced hypersecretion mainly comes from studies in rats (Cassuto et al., 1982a; Cassuto et al., 1983; Mourad et al., 1995; Turvill et al., 1998; Turvill et al., 2000a, b; Kordasti et al., 2006) and guinea pigs (Carey and Cooke, 1986; Kirchgessner et al., 1992; Gwynne et al., 2009). The ENS incorporates two ganglionated plexuses: the submucosal and the myenteric plexus plus several plexuses of axons, situated within the walls of the gastrointestinal tract. The submucosal plexus contains efferent secretomotor neurons and regulates intestinal secretion, and hence has been the focus of most CT studies. CT is thought to indirectly activate enteric pathways by stimulating serotonin (5-HT) release from enteroendocrine cells (Beubler et al., 1989a), which in turn excites intrinsic sensory nerve endings in the mucosa via 5-HT3 receptors (Turvill and Farthing, 1997; Bertrand et al., 2000a; Kordasti et al., 2006). Gwynne et al (2009) showed that luminal incubation of CT in segments of guinea pig jejunum in vitro induced sustained hyperexcitability of the two major subsets of secretomotor neurons: cholinergic (acetylcholine, ACh-containing) and non-cholinergic (vasoactive intestinal peptide, VIP- containing) neurons. Thus, it was postulated that the increased excitability of secretomotor
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neurons may be involved in the mechanism underlying CT-evoked hypersecretion (Gwynne et al., 2009). However, the myenteric plexus also contributes significantly to CT-induced hypersecretion as chemically ablating this plexus can inhibit the secretory response (Jodal et al., 1993). Moreover, CT has also been shown to affect intestinal motility (Fung et al., 2010; Balasuriya et al., 2016). This highlights a key limitation in many in vitro studies examining CT-effects, as muscle-stripped preparations lacking the myenteric plexus were commonly used (Field et al., 1972; Carey and Cooke, 1986; Burleigh and Borman, 1997). Thus, the neural contribution to CT-mediated effects may have been underestimated using such models.
Studies in vivo suggest that CT-induced secretion in rats and human tissues is predominantly mediated by VIP, as various VIP antagonists attenuate cholera hypersecretion (Mourad and Nassar, 2000; Banks et al., 2005; Kordasti et al., 2006). However, there are considerable discrepancies between these studies, as the effectiveness of specific VIP antagonists in attenuating CT-induced secretion was inconsistent even with similar animal models. Further, in vitro studies in guinea pig small intestine and human epithelial cells implicate the involvement of ACh (Carey and Cooke, 1986; Banks et al., 2004; Gwynne et al., 2009), although muscarinic antagonists do not have any significant effects on CT-evoked hypersecretion (Cassuto et al., 1982a; Kobayashi et al., 2001; Kordasti et al., 2006; Gwynne et al., 2009). Thus, the relative contributions of cholinergic and non-cholinergic secretomotor neurons to CT-induced hypersecretion remain unclear.
This study aimed to examine the relative contributions of submucosal and myenteric neurons to the sustained hypersecretion evoked by CT. A combination of in vivo and in vitro techniques was used. CT was incubated in vivo to more closely mimic conditions in the whole animal and then in vitro analyses were performed to provide the spatial resolution required to examine the circuitry involved. Mice in which the ENS expresses a genetically encoded fluorescent Ca2+ indicator GCaMP3 were used to examine the
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excitability of the enteric circuitry following CT exposure with Ca2+ imaging. It was found that CT induced a sustained hypersecretion by a direct action on the mucosa, and in part by specifically increasing the excitability of cholinergic submucosal neurons.
3. MATERIALS AND METHODS
3.1 Mice
8 – 12 week old male mice on a C57BL6 background, including Wnt1-Cre;R26R-GCaMP3 mice (Danielian et al., 1998; Zariwala et al., 2012) were used (Chapter 2).
3.2 In vivo incubation of CT in a ligated ileal loop
An ileal loop was constructed as described in Chapter 2 (section 8), and was incubated with sterile physiological saline (Krebs’ composition in mM: NaCl 118, NaHCO3 25, D-glucose
11, KCl 4.8, CaCl2 2.5, MgSO4 1.2, NaH2PO4 1.0) as a control, or CT (12.5 ug/ml; List Biologicals, Campbell, CA) in physiological saline, for 3.5 h. The ileal loop was removed from abdomen and flushed clean of the luminal contents prior to in vitro analyses. In preliminary experiments, 1 – 2 cm segments of ileum approximately 2 cm proximal to the ileal loop, the jejunum, and the proximal colon were also collected to assess potential off- target effects.
3.3 In vitro measurement of short-circuit current in Ussing chambers
As previous studies have suggested that the myenteric plexus is important in the CT-evoked hypersecretion (Jodal et al., 1993), full thickness preparations in which all intestinal layers
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are intact were used rather than standard mucosa-submucosal plexus preparations. Each ileal loop provided up to 2 Ussing chamber preparations (refer to Chapter 2, section 4).
Short-circuit current (ISC) was measured throughout the experiment. Preparations were equilibrated for 30 min prior to collecting data to allow for the basal ISC of the tissue to stabilize.
Data collection and analysis were performed using AcqKnowledge 3.9.0 software (BIOPAC Systems, Inc., SDR Clinical Technology, Middle Cove, NSW, Australia). The maximum change in ISC from baseline (ΔISC) induced by drug addition was calculated. Data obtained from tissue of the same animal were matched; control data from adjacent segments of mouse ileum were not significantly different. Drugs were added to the serosal side of the preparation.
3.4 Ca2+ imaging and analysis
Ileal loops of Wnt1-Cre;R26R-GCaMP3 mice were used for Ca2+ imaging (refer to Chapter 2, section 7). Up to 2 ring preparations of each plexus (submucosal and myenteric) were obtained from each ileal loop. Preparations were constantly superfused (1 ml/min) with
95% O2: 5% CO2 physiological saline at room temperature throughout the experiment. Ring preparations were imaged through an Olympus 20× (NA 0.5) water dipping lens on an upright Zeiss Axioskope microscope with a Zeiss AxioCam MRm camera and images (278 × 278) were acquired at 1Hz.
Preparations were stimulated either chemically using the nicotinic agonist 1,1-dimethyl-4- phenylpiperazinium (DMPP) or electrically (Foong et al., 2015). DMPP (10 µM) was applied to the preparations by pressure ejection (500 ms duration; 9 psi). Electrical stimuli (single pulse or train of stimuli: 300 μs, 20 pulses, 20 Hz) were applied via a stimulating
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electrode (50 µm non-insulated tungsten wire) on an internodal strand leading to the ganglion imaged. Some preparations were also stimulated chemically with the VPAC1 agonist [K15, R16, L27] VIP (1-2)/GRF(8-27) (refer to Chapter 4).
Ca2+ imaging analyses were performed using Igor Pro (Wavematrics, Lake Oswego, 2+ Oregon, USA) (Boesmans et al., 2013a). The amplitude of each [Ca ]i peak was calculated for each response and was measured as the maximum increase in fluorescence from baseline (∆Fi/F0) as described previously (Chapter 2). At least 3 animals were examined for each condition.
3.5 Drugs
CT, 1,1-dimethyl-4-phenylpiperazinium (DMPP; Sigma Aldrich), and tetrodotoxin (TTX; Alomone Labs, Jerusalem, Israel) were dissolved in distilled water to make stock solutions. Veratridine (Sigma Aldrich) was dissolved in 80% ethanol. Stocks were stored at 4°C. Drugs were then diluted in physiological saline to working concentrations when required.
3.6 Immunohistochemistry
Fixed wholemount preparations, Ca2+ imaging preparations (see Chapter 4), and cryosections were used for immunohistochemical labelling (also refer to Chapter 2, sections 3 and 9).
Briefly, wholemount preparations of submucosal plexus and myenteric plexus were double or triple-labelled by incubating with various combinations of primary antisera (Table 5.1) including rabbit α pCREB or rabbit α c-Fos for 48 - 72 h at 4°C. Preparations were then
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washed in PBS (3 x 10 min), and incubated with biotinylated donkey α rabbit IgG (1:100; Jackson Immuno Labs, West Grove, Pennsylvania, USA) for 2 h at room temperature. Following PBS washes (3 x 10 min), preparations were incubated with strepavadin AF594 (1:200; Molecular Probes, Eugene, Oregon, USA) and secondary antisera (Table 5.1) for 2 h at room temperature.
Cryosections (10 µm) were incubated in a goat α CT-B (Table 5.1) overnight at 4°C. Excess antisera were washed away with 3 x 10 min PBS washes. Sections were then incubated with a secondary antiserum for 3 h at room temperature.
3.7 Confocal imaging and cell counts
Submucosal plexus and myenteric plexus preparations, and cryosections were imaged with 40x and 20x objectives respectively, using a Zeiss Pascal confocal microscope. The proportion of each neuronal subtype (ChAT+, nNOS+, calretinin+, pCREB+, and/or c-Fos+) was obtained by examining co-expression with at least 100 Hu+ submucosal neurons or 200 Hu+ myenteric neurons per preparation. The mean proportion of each neuronal subtype was determined by averaging the proportions obtained from three to five animals.
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Table 5.1. Primary and secondary antibodies used for immunohistochemistry.
Primary Antibodies Host Dilution Source
pCREB Rabbit 1:1000 Millipore
c-Fos Rabbit 1:5000 Oncogene
ChAT Goat 1:500 Chemicon
nNOS Sheep 1:1000 Gift from P. Emson
calretinin Goat 1:1000 SWANT
Hu Human 1:5000 Gift from Miles Epstein
CT-B Goat 1:1000 List Biologicals; gift from Colin Anderson
Secondary Antibodies Host Dilution Source
Anti-goat AF594 Donkey 1:1000 Molecular Probes
Anti-sheep AF647 Donkey 1:500 Molecular Probes
Anti-sheep AF488 Donkey 1:400 Molecular Probes
Anti-human AF647 Donkey 1:500 Jackson Immuno Labs
Anti-human AF488 Donkey 1:800 Jackson Immuno Labs
Abbreviations: ChAT, choline acetyltransferase; nNOS, neuronal nitric oxide synthase
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3.8 Data and statistics
Data are presented as mean ± the standard error of the mean (SEM), and ‘n’ number of cells examined unless stated otherwise. Two-tailed unpaired t-tests were conducted to determine statistical significance, unless specified otherwise. P < 0.05 was considered significant. Analyses were performed using GraphPad Prism 5.0 (GraphPad softwares, San Diego, California, USA).
4. RESULTS
Following 3.5 h incubation of CT (12.5 µg/ml), all ligated ileal loops displayed an increased accumulation of fluid in the lumen compared to the saline controls (Figure 5.1A, B). As previously reported in the guinea pig jejunum (Gwynne et al., 2009), anti-CT-B labelling was only observed in CT-treated tissues and was confined to the mucosal epithelium (Figure 5.1C, D). Notably, CT-B-immunoreactivity was observed predominantly in the villi, but also close to the crypts (Figure 5.1 D).
4.1 Acute CT exposure induces a sustained increase in basal secretion
First, it was confirmed that this model of CT-incubation induced a sustained hypersecretion in full thickness preparations by monitoring basal short-circuit current (ISC) in Ussing 2 chambers. The basal ISC in CT-exposed tissues was 105 ± 8 µA/cm (n = 11 mice) and was significantly higher than in control tissues (51 ± 8 µA/cm2; n = 10 mice; P < 0.0001; Figure 5.1G).
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A neuronal component underlying basal secretion was identified using tetrodotoxin (TTX, 1 µM; voltage-dependent Na+ channel blocker) applied to the serosal side of the preparation.
The reduction in basal ISC by TTX did not differ significantly between control (∆ISC: -6.8 ± 2 2 3.0 µA/cm ; n = 6) and CT-treated preparations (∆ISC: -9.9 ± 1.9 µA/cm ; n = 4) (Figure 5.1H). Moreover, TTX did not return the increased basal secretion of CT-treated preparations to control levels. There were no differences observed between control and CT- treated preparations in responses to neural stimulation with DMPP (nicotinic agonist) or veratridine (activates voltage-dependent Na+ channels) applied serosally over a range of concentrations (1, 3, 10, 30 µM) (Figure 5.1E, F).
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Figure 5.1. CT-treated vs. control ileal loops following 3.5 h incubation in vivo.
Images of ileal loops after a 3.5 h incubation in vivo with saline (control, A), or CT (12.5 µg/ml, B) in saline. CT-treated loops displayed substantial accumulation of fluid that was absent in controls. Confocal micrographs of cryosections from control (C) and CT-treated ileal loops (D) fluorescently labelled for CT-B. CT-B staining was specifically observed in CT-treated tissues and was confined to the mucosal epithelium. CT-B-immunoreactivity was observed predominantly in the villi, but also close to the crypts, as indicated by arrowheads. Scale bar = 100 µm. Concentration-effect curves (1, 3, 10, 30 µM) for E. DMPP (nicotinic agonist) and F. veratridine (voltage-sensitive Na+ channel activator) applied serosally to control and CT-treated tissues. No significant differences in neurally- evoked secretion was observed (DMPP: n = 10 per condition; veratridine: n = 8 per condition). G. Basal ISC measured in full thickness ileal tissues following CT-incubation (n = 11) were significantly higher than that of saline controls (n = 10; ****P < 0.0001). H.
Representative traces of tetrodotoxin (TTX)-evoked depression of basal ISC in control (blue) and CT-treated (red) preparations. TTX application is marked by the arrow.
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4.2 CT-incubation induced pCREB expression in submucosal neurons
To examine submucosal neurons that may have been active during CT-incubation, the nuclear expression of two activity-dependent markers, pCREB and c-Fos (Sheng and Greenberg, 1990; Ginty et al., 1992), was examined in the submucosal plexus of CT- incubated and control preparations (Figure 5.2). Higher levels of pCREB expression were observed in CT-incubated preparations where 66 ± 6% of all submucosal neurons were pCREB+ (n = 3 mice, 404 Hu+ neurons examined), while only 11 ± 5% were pCREB+ in controls (n = 3 mice; 390 Hu+ neurons examined; P = 0.002) (Figure 5.2E). No c-Fos expression was observed in the submucosal plexus, although the conditions under which c- Fos expression is induced in submucosal neurons of mouse are unclear as this has not been previously examined.
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Figure 5.2. pCREB labelling of submucosal neurons in control vs CT-treated ileal loops.
A – D. Confocal images of submucosal ganglia from a control (A, C) and a CT-treated ileal loop (B, D), labelled for pCREB (red) and the neuronal marker Hu (blue). Scale bar = 20 µm. E. A significantly higher proportion of submucosal neurons (Hu+) displayed nuclear pCREB staining (as indicated by arrows) after CT-treatment compared to controls (n = 3 in each condition; **P < 0.01).
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4.3 CT-exposure increases sustained excitability in cholinergic submucosal neurons
Using Ca2+ imaging, submucosal plexus preparations from ileal loops of Wnt1-Cre;R26R- GCaMP3 mice were first imaged without stimulation to examine spontaneous activity. 2+ Submucosal neurons rarely displayed spontaneous [Ca ]i transients in control preparations (4/149 neurons were spontaneously active). However, a significantly larger proportion of 2+ submucosal neurons (31/160 neurons; Figure 5.3A) displayed spontaneous [Ca ]i transients in CT-incubated preparations (Fisher’s exact test; P < 0.0001). These spontaneously active neurons were found to be exclusively cholinergic using post hoc immunostaining for ChAT (choline acetyltransferase, the synthesizing enzyme for ACh; Figure 5.3B – D).
To then assess any changes to the excitability of the neural network, submucosal ganglia were stimulated chemically and electrically (Figure 5.4). DMPP (10 µM; nicotinic agonist) 2+ application by pressure ejection evoked [Ca ]i transients with a significantly higher response amplitude (∆Fi/F0) after CT-exposure (CT: ∆Fi/F0 = 0.61 ± 0.05, n = 61; control: ∆Fi/F0 = 0.51 ± 0.02, n = 90; P = 0.044; Figure 5.4D). Using post hoc labelling of imaged ganglia, cholinergic ChAT+ neurons specifically were found to display significantly larger responses to DMPP in CT-exposed tissues compared to controls (P = 0.017). The transients seen in ChAT- neurons did not differ significantly between CT and saline- incubated preparations.
Electrical stimulation (single pulse, and a train of stimuli) also evoked significantly larger 2+ [Ca ]i responses in CT-incubated submucosal plexus preparations compared to controls. A single pulse stimulated a larger response amplitude in submucosal neurons of CT-treated preparations (CT: ∆Fi/F0 = 0.15 ± 0.01, n = 39; controls: ∆Fi/F0 = 0.11 ± 0.01, n = 46; P = 0.004; Figure 5.4E). Again, it was specifically cholinergic ChAT+ neurons that displayed larger responses (CT: ∆Fi/F0 = 0.16 ± 0.02, n = 22; control: ∆Fi/F0 = 0.10 ± 0.01, n = 36; P
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= 0.0006). Similarly, a train of stimuli evoked larger responses after CT-incubation (CT: ∆Fi/F0 = 0.51 ± 0.02, n = 64 vs. control: ∆Fi/F0 = 0.39 ± 0.02, n = 92; P = 0.0003; Figure 5.4F) and this was also specific to cholinergic neurons (CT: ∆Fi/F0 = 0.61 ± 0.04, n = 24; control: ∆Fi/F0 = 0.44 ± 0.02, n = 58; P = 0.0002; Figure 5.4F). Responses to electrical stimulation, whether single pulse or trains, did not differ between saline and CT-incubated ChAT- neurons.
Figure 5.3. Spontaneous activity in submucosal neurons following CT-incubation.
2+ A. The frequency of spontaneous [Ca ]i transients in submucosal neurons of control vs. CT-treated tissues. Submucosal neurons of CT-treated preparations displayed significant levels of spontaneous activity, while this was rarely observed in control preparations. B. A submucosal ganglion which showed spontaneous activity following CT-exposure. C. These neurons were identified as cholinergic neurons using post hoc staining for choline acetyltransferase (ChAT, red). Scale bar = 20 µm. D. Traces from spontaneously active neurons marked in B and C.
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2+ Figure 5.4. DMPP and electrically-evoked [Ca ]i responses in submucosal neurons are increased following CT-exposure.
A – C. A submucosal ganglion from a CT-incubated preparation responding to a train of stimuli (20 pulse). Cholinergic neurons were identified with post hoc labelling for choline acetyltransferase (ChAT, red, C). Arrows indicate cholinergic ChAT+ neurons. Scale bar = 20 µm. D. DMPP (10 µM) response amplitudes (∆Fi/F0) in control (blue) vs. CT-treated (red) preparations. In each histogram (D – F) the amplitude of ‘Total’ neurons that responded was presented (left) then divided into those of ChAT- (middle) and ChAT+ (right) groups. Overall responses to DMPP were significantly higher after CT-treatment (*P < 0.05), and in particular, responses in cholinergic (ChAT+) submucosal neurons were significantly increased (*P < 0.05). Electrically-evoked responses (E. single pulse and F. 20 pulse) of submucosal neurons after CT-incubation had a higher amplitude compared to that of controls (Single pulse: **P < 0.01, 20 pulse: ***P < 0.001), and this was specifically observed in cholinergic neurons (***P < 0.001). G. Averaged traces of responses to 20 pulse stimulation (applied at 10 s) in ChAT+ (black trace) vs ChAT- neurons (grey trace; mean ± SEM) in a CT-treated preparation, as depicted in A – C.
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2+ 4.4 CT-exposure did not alter VPAC1 agonist-evoked glial [Ca ]i responses
Since a number of Hu- cells within the submucosal ganglia – presumably glial cells – were observed to express pCREB following CT-exposure (Figure 5.2B, D), whether CT might also affect enteric glial activity was also examined. Given the findings from Chapter 4, local spritz application of the VPAC1 agonist ([K15, R16, L27]VIP(1-7)/GRF(8-27)) (100 µM) was used to evoke glial responses in submucosal plexus preparations. However, no significant differences in glial response amplitudes were observed between CT-treated and control preparations (CT: ∆Fi/F0 = 0.23 ± 0.04, n = 19; control: ∆Fi/F0 = 0.30 ± 0.02, n = 45; P = 0.16). The number of responding glial cells also did not significantly differ (CT: ∆Fi/F0 = 2.7 ± 0.5; control: ∆Fi/F0 = 4.5 ± 0.7; P = 0.08).
4.5 CT-exposure induces c-Fos expression in myenteric plexus
The expression of pCREB and c-Fos was also examined in the myenteric plexus of CT- incubated and control preparations. There was a significant increase in c-Fos expression in the myenteric plexus after CT-incubation, where 28 ± 3% of myenteric neurons displayed c-Fos-immunoreactivity (n = 5 mice; 2644 Hu+ neurons examined), while only 6 ± 2% of myenteric neurons were c-Fos+ in controls (n = 5 mice; 2438 Hu+ neurons examined) (Figure 5.5). Given that c-Fos expression was observed in some myenteric neurons, various neurochemical markers were used to determine if specific functional subtypes were activated by CT-treatment. c-Fos expression was mainly observed in nNOS+ neurons (63 ± 5% of c-Fos+ neurons were nNOS+ and 56 ± 6% of nNOS+ neurons were c-Fos+; n = 4 mice; Figure 5.5B, E). The remaining c-Fos+ neurons were mostly calretinin+ neurons, where 30 ± 1% of c-Fos+ neurons were calretinin+ (19 ± 2% of calretinin+ neurons were c- Fos+; n = 3 mice; Figure 5.5C, F). This suggests that motor neurons and interneurons were activated (Furness, 2000; Qu et al., 2008). Notably, there was a lack of c-Fos labelling in large calretinin+ myenteric neurons, which are typically sensory AH neurons (Qu et al.,
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2008). pCREB expression in the myenteric plexus was profuse even in control preparations and thus was not a useful marker.
Figure 5.5. c-Fos labelling in myenteric neurons in control vs. CT-treated ileal loops.
A – F. Confocal images of myenteric ganglia from a control (A, D) and a CT-treated ileal loop (B, C, E, F), labelled for c-Fos (red) and the neuronal marker Hu (blue). Some CT- treated preparations were co-labelled with neuronal nitric oxide synthase (nNOS, E) and some were co-labelled with calretinin (calr, F). Arrows indicate Hu+ neurons displaying nuclear c-Fos labelling and nNOS or calretinin expression. Scale bar = 20 µm. G. A significantly higher proportion of myenteric neurons (Hu+) displayed nuclear c-Fos labelling after CT-treatment compared to controls (n = 5 in each condition; ****P < 0.0001).
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4.5 The myenteric plexus displays an overall reduction in neuronal excitability following
CT-exposure
In contrast to the submucosal plexus, a smaller proportion of myenteric neurons displayed 2+ spontaneous [Ca ]i transients in CT-treated preparations (1/251 neurons were spontaneously active) than controls (14/279 neurons; Fisher’s exact test P < 0.01).
2+ Further, a reduction in the amplitude of [Ca ]i responses to DMPP (10 µM) was observed in CT-treated preparations compared to controls (CT: ∆Fi/F0 = 0.77 ± 0.03, n = 152; control: ∆Fi/F0 = 0.88 ± 0.04, n = 146; P = 0.028; Figure 5.6D). Given that a large proportion of c-Fos+ neurons were nNOS+, the responding neurons were divided into + - - nNOS and nNOS groups using post hoc immunostaining. nNOS neurons were found to display smaller responses after CT-treatment (CT: ∆Fi/F0 = 0.79 ± 0.04; n = 105; control: ∆Fi/F0 = 0.91 ± 0.04; n = 119), while responses in nNOS+ neurons were unchanged. Stimulating electrically with a single pulse similarly evoked smaller responses in myenteric neurons of CT-treated preparations (CT: ∆Fi/F0 = 0.20 ± 0.01, n = 84; control: ∆Fi/F0 = 0.28 ± 0.03, n = 62; P = 0.002; Figure 5.6E). This effect was observed in nNOS+ neurons (CT: ∆Fi/F0 = 0.19 ± 0.02, n = 8; control: ∆Fi/F0 = 0.41 ± 0.07, n = 9; P = 0.016), rather than nNOS- neurons. In contrast, nNOS- neurons appear to show a significantly increased response to the train of stimuli after CT-exposure (CT: ∆Fi/F0 = 0.92 ± 0.05, n = 101; control: ∆Fi/F0 = 0.76 ± 0.04, n = 81; P = 0.012; Figure 5.6F), while responses of nNOS+ neurons were not different. This difference was not apparent when comparing the overall response amplitude of all neurons responding (CT: ∆Fi/F0 = 0.87 ± 0.04, n = 131; control: ∆Fi/F0 = 0.80 ± 0.03, n = 112; P = 0.17).
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2+ Figure 5.6. DMPP- and electrically-evoked [Ca ]i responses in myenteric neurons are decreased following CT-exposure.
A – C. A myenteric ganglion from a CT-incubated preparation responding to a train of stimuli (20 pulse). Cholinergic neurons were identified with post hoc labelling for nNOS (neuronal nitric oxide synthase; red, C). Arrows indicate the nNOS+ neurons that responded. Scale bar = 20 µm. D. DMPP (10 µM) response amplitudes (∆Fi/F0) in control (blue) vs. CT-treated (red) preparations. In each histogram (D – F) the amplitude of ‘Total’ neurons that responded was presented (left) then divided into those of nNOS- (middle) and nNOS+ (right) groups. Overall DMPP response amplitudes were significantly smaller after CT- treatment, and in particular, nNOS- myenteric neurons were significantly decreased (*P < 0.05). E. Single pulse stimuli also evoked smaller amplitude responses after CT-incubation (**P < 0.01) and this was specifically observed in nNOS+ neurons (*P < 0.05). F. Overall responses to 20 pulse stimulation were not different after CT-exposure, however a slight but significant increase (*P < 0.05) was observed in nNOS- neurons. G. Averaged traces of responses to 20 pulse stimulation (applied at 10 s) in nNOS+ (black trace) vs nNOS- neurons (grey trace; mean ± SEM) from a CT-treated preparation, as depicted in A – C.
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4.6 Effects of CT were confined to the ileal loop
Some bacterial toxins such as Clostridium difficile toxin A have been shown to activate nerve reflexes involving extrinsic neurons to induce widespread inflammatory and secretory responses (Pothoulakis and Lamont, 2001; Spiller, 2002). The possibility that CT may spread its hypersecretory effects through extrinsic networks was considered in preliminary studies and potential off-target effects were assessed in intestinal segments outside the ileal loop. Ileal segments proximal to the CT-treated loop were examined in 2 Ussing chambers. Basal ISC in these tissues was 43 ± 9 µA/cm (n = 9 mice) and was not different from that of control ileal loops (51 ± 8 µA/cm2; n = 10 mice) or ileal segments proximal to the control ileal loop (42 ± 15 µA/cm2; n = 4 mice).
Additionally, c-Fos labelling was observed to be confined to the CT-incubated ileal loop, with no labelling in the jejunum, the region just proximal to the ileal loop, or the region distal to the ileal loop (proximal colon of these mice (n = 4; Figure 5.7), suggesting that CT does not have any off-target effects.
Electrical stimulation (single pulse, and a train of stimuli) of submucosal and myenteric plexus preparations of ileal segments proximal to the CT-treated loop (n = 3 mice) and 2+ proximal to control loops (n = 2 mice) evoked neuronal [Ca ]i responses that did not differ from those of control loops (Figure 5.8).
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Figure 5.7. c-Fos labelling of myenteric neurons were confined to the CT-treated ileal loop.
A – D. Confocal images of myenteric ganglia from jejunum (A, A’), a segment proximal to the ileal loop (B, B’), proximal colon (D, D’), and the CT-treated ileal loop (C, C’), labelled for c-Fos (red), nNOS (green) and the neuronal marker Hu (blue). Arrows indicate neurons (Hu+) displaying colocalization of c-Fos- and nNOS-immunoreactivity. Nuclear c- Fos labelling was confined to the CT-treated ileal loop and was not observed in other intestinal regions. Scale bars = 20 µm.
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Figure 5.8. Changes in excitability of enteric neurons following CT-exposure were confined to the ileal loop.
2+ A. [Ca ]i responses of submucosal neurons to electrical stimulation (single pulse and 20 pulse) in segments of ileum proximal to the control loop or proximal to the CT-treated loop did not differ from controls (*P < 0.05, ***P < 0.0001 vs. control). B. Responses amplitudes (∆Fi/F0) of myenteric neurons to electrical stimulation (single pulse and 20 pulse) in segments of ileum proximal to the control loop or proximal to the CT-treated loop also did not differ from controls (*P < 0.05 vs. control). Number of neurons examined are displayed within the bars.
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5. DISCUSSION
Previous studies in rats and humans have shown that CT-evoked hypersecretion is predominantly associated with non-cholinergic VIP-containing secretomotor neurons (Cassuto et al., 1982a; Mourad and Nassar, 2000; Banks et al., 2005; Kordasti et al., 2006). Although Gwynne et al. (2009) demonstrated that CT-incubation in vitro induced a sustained hyperexcitability in both VIP- and cholinergic NPY-containing secretomotor neurons in guinea pig, findings from this present study suggest that any neural contribution to the CT secretory response primarily involves cholinergic neurons in the mouse. Indeed, the main action of CT in this model appears to be a direct effect on the mucosal epithelium.
5.1 CT primarily acts on the mucosa epithelium to induce hypersecretion
There are two possible sites which CT may target either directly or indirectly (i.e. mucosa and/or ENS) and whether a particular site or both are involved in the effects of CT continues to be disputed in the literature. In this present study, I found that CT exerts its hypersecretory effects mainly at the mucosal level, with only a small contribution from the ENS. Addition of the neural activity-blocker TTX to the serosal half of Ussing chambers did not return the basal ISC of CT-treated preparations to control levels. Additionally, responses to chemically-evoked secretion using the nicotinic agonist DMPP and Na+ channel activator veratridine were also similar between controls and CT-incubated preparations. Yet, in concordance to the previous study in the guinea-pig jejunum (Gwynne et al., 2009), I also demonstrated CT-induced sustained hyperexcitability in secretomotor neurons. Interestingly, the evidence for CT primarily exerting its hypersecretory effects by directly acting on the mucosa was largely obtained from in vitro Ussing chamber studies (Field et al., 1971; Field et al., 1972; Carey & Cooke, 1986a; Moriarty et al., 1989), whereas data suggesting a substantial neural contribution from the ENS comes from in vivo models (Cassuto et al., 1982a; Cassuto et al., 1983; Mourad et al., 1995; Turvill et al.,
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2000b, a; Kordasti et al., 2006). However, it is important to note that while CT was incubated in vivo in this present study, analysis of its effects was conducted in vitro. Moreover, it was the sustained changes in the enteric circuitry following CT-exposure that were assessed in the present study, rather than effects in the presence of CT.
The reason for the discrepancies between in vivo and in vitro studies remain unclear. Field and Semrad (1993) proposed that, as in vitro Ussing chamber preparations are well stretched, CT does in fact reach the crypts to directly stimulate secretion and thus the direct mucosal effect is more prominent. They also suggested that this model more closely mimics cholera infection where V. cholerae grow on the surface of both villi and crypts (Millet et al., 2014), whereas in vivo models may not allow adequate mixing or luminal distension for CT to reach the crypts. CT was incubated in vivo in this study, and CT-B labelling was found to be largely on the mucosal villi as previously reported (Weiser and Quill, 1975; Gwynne et al., 2009), yet a major mucosal effect was observed. One possibility is that CT does contact the secretory region in the mouse ileum as the CT-B labelling appears to reach the start of the crypts.
5.2 CT induced sustained hyperexcitability in cholinergic submucosal neurons
I found increased neuronal excitability in the submucosal plexus following CT-incubation. Specifically, it was the cholinergic submucosal neurons that displayed an increase in spontaneous activity and increased responses to chemical and electrical stimuli. This was surprising given that much of the previous literature highlights a role for VIP, a potent secretagogue, and VIP-containing secretomotor neurons (Mourad et al., 1995; Banks et al., 2005; Kordasti et al., 2006; Gwynne et al., 2009). Increased levels of VIP have been found in the stool of patients with cholera (Bloom et al., 1976), and VIP release is also increased in an in vivo model of CT-induced hypersecretion in cats (Cassuto et al., 1981b). There is considerable evidence showing that VIP antagonists can attenuate CT-evoked
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hypersecretion, although the involvement of VIP has not been completely clear as there have been discrepancies between studies in their reports of the efficacy of different VIP antagonists (Mourad and Nassar, 2000; Banks et al., 2005; Kordasti et al., 2006). Nonetheless, my finding that cholinergic submucosal neurons are involved in the sustained effects of CT is consistent with observations by Carey & Cooke (1986), which suggests that cholinergic secretomotor neurons contribute to the CT-induced hypersecretion. It has also been shown that the nicotinic antagonist hexamethonium attenuates CT-evoked hypersecretion once it had established, but did not affect the induction of the response (Kordasti et al., 2006). This further supports my observation of the involvement of cholinergic neurons in the sustained secretory effects of CT. Additionally, Banks and colleagues (2004) have reported a cholinergic contribution to CT-evoked hypersecretion whereby the secretory responses to acetylcholine are potentiated by the presence of CT at the level of the enterocytes. If CT evokes a sustained hyperexcitability in cholinergic submucosal neurons and cholinergic output is enhanced, this additional mechanism at epithelium may further amplify the secretory response.
Interestingly, the nicotinic antagonist hexamethonium, but not the muscarinic antagonist atropine, is effective in inhibiting CT-evoked hypersecretion (Cassuto et al., 1982a; Kobayashi et al., 2001; Kordasti et al., 2006). The cholinergic pathways involved are yet to be elucidated. One possibility is that cholinergic submucosal neurons form recurrent networks to reinforce activity within the circuitry via nAChR-mediated fast EPSPs (Chambers et al., 2005). Hence, CT increases the excitability of interconnected cholinergic submucosal neurons, which signal via nAChRs, to increase the output secretory response. However, whether submucosal neurons make connections with other submucosal neurons in the mouse small intestine as in the guinea pig remains to be determined (Reed and Vanner, 2001). Further, computer modelling data suggests that nAChR activation and increased fast EPSP transmission only evokes transient effects, whereas slow EPSP input from sensory and/or VIP neuron networks is necessary for sustaining hyperactivity in the circuitry (Chambers et al., 2005). Thus, an increase in cholinergic drive, without increasing
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slow EPSP input, perhaps accounts for the lack of a substantial neural contribution to CT- evoked hypersecretion observed in the mouse.
The expression of activity dependent markers pCREB and c-Fos indicate an increase in neuronal activity during CT-incubation. In particular, the majority of submucosal neurons, cholinergic and non-cholinergic, were active during CT-exposure as marked by pCREB and this is consistent with the increased fluid secretion observed. While CT predominantly evoked sustained changes in the properties of cholinergic submucosal neurons, it is possible that VIP secretomotor neurons are involved in the induction of the hypersecretion, rather than maintaining hypersecretion. This is compatible with work by Kordasti et al. (2006) showing that VIP antagonism was only effective in preventing CT-induced hypersecretion, but was ineffective in attenuating an already established hypersecretion. While pCREB expression may have also been induced in glial cells by CT-exposure, no apparent changes in glial activity were found using the VPAC1 agonist (Chapter 4). Thus, there is no evidence for any glial involvement in the sustained effects of CT.
The other neuron activity marker, c-Fos expression was not observed in submucosal neurons in either the control condition or following CT-exposure. While pCREB is generally associated with the activation of G-protein coupled receptors and increased cAMP, and c-Fos expression is associated with Ca2+-influx through voltage-gated Ca2+ channels, there is a degree of overlap in these secondary messenger pathways (Sheng and Greenberg, 1990; Sheng et al., 1991). In some cells, phosphorylation of CREB can also be stimulated by Ca2+ (Sheng and Greenberg, 1990; Sheng et al., 1991). Furthermore, pCREB can activate c-Fos expression by binding a Ca2+ response element (CRE) located upstream of the initiation site for c-fos transcription (Ginty et al., 1992). Hence, the absence of c-Fos labelling was surprising as pCREB acts upstream of c-Fos. However, the circumstances under which c-Fos expression is activated in submucosal neurons in the mouse intestine remain unclear. Only a limited number of other studies have used c-Fos as an activity
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dependent marker in the mouse submucosal plexus (Bjerknes and Cheng, 2001; Hitch et al., 2012). Of these, either a lack of expression in submucosal neurons was reported (Hitch et al., 2012) or c-Fos expression in submucosal neurons was not convincingly demonstrated (Bjerknes and Cheng, 2001). The temporal dynamics of pCREB and c-Fos expression may differ as pCREB expression is stimulated more rapidly than the transcription and translation of c-Fos, since it simply requires phosphorylation by cAMP activated protein kinase A (Fields et al., 1997; Ji and Rupp, 1997; Gammie and Nelson, 2001). However, considering that the duration of CT-incubation was 3.5 h, it seems unlikely that the absence of c-Fos in submucosal neurons is simply due to a difference in the timing required for its induction. Alternatively, it may be that the intensity of the stimulus is simply not sufficient to evoke c-Fos expression in submucosal neurons, whereas pCREB expression is observed due to its lower activation threshold (Fields et al., 1997).
5.3 Some myenteric neurons may be hyperactive during but not following CT incubation
CT-exposure did not evoke significant changes in the excitability of myenteric neurons. Despite the observed increase in c-Fos expression in myenteric neurons, this was seen in a relatively small proportion of neurons, including NOS+- and calretinin+- motor neurons and interneurons. Thus, it is perhaps predictable that only subtle changes to the activity of this plexus were found. An overall depression of spontaneous activity and stimulated responses were seen with Ca2+ imaging, despite a slight increase in responses to trains of electrical stimuli in NOS- neurons. It may be that myenteric neurons contribute to the induction of CT-evoked hypersecretion, but not its maintenance, as it has been proposed that these are separate processes involving distinct neural pathways (Kordasti et al., 2006). Another possibility is that the interneurons and motor neurons which expressed c-Fos were activated secondarily to the distension caused by fluid accumulation, rather than as a direct effect of the toxin. In contrast with the hypersecretory effects which may take hours to develop, CT can evoke rapid changes in motility within 15 min of exposure (Fung et al., 2010; Balasuriya et al., 2016). Thus, it is also feasible that c-Fos expression may result from an
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activation of neurons involved in these early motility effects. pCREB was not a useful marker in the myenteric plexus as its expression was abundant even in control tissues. This is not surprising given that spontaneous activity within the myenteric plexus was observed in control preparations, and suggests a significant level of basal neural activity in the myenteric plexus.
It has been shown in the submucosal plexus that hyperexcitability is only observed in neurons in close proximity to the mucosa, but not in regions where the mucosa was removed (Gwynne et al., 2009). It may be that changes to neuronal excitability in the myenteric plexus also require the ongoing release of mucosal mediators. As the myenteric plexus preparations used for Ca2+ imaging are stripped of the mucosa, this may account for the lack of effects in these neurons. Further, a potential limitation is that Ca2+ imaging was used to examine network excitability. As Ca2+ signals are only secondary to electrical events, rather than a direct measure, it may be that this technique is not sensitive enough to detect subtle changes to neuronal excitability. However, given that Ussing chamber experiments showed a lack of a prominent neural component to basal and stimulated secretion in CT-treated full thickness preparations, there is little evidence for any substantial increases in neural activity. Collectively, the data suggests that the myenteric plexus does not play any significant role in the sustained effects following acute CT- exposure. Nevertheless, as with VIP secretomotor neurons, the potential involvement of the myenteric plexus in the induction of CT-induced hypersecretion cannot be ruled out.
5.4 Conclusion and future directions
This is the first study that combined in vivo and in vitro techniques to examine CT-evoked secretion and the role of both the submucosal and myenteric plexuses. It was demonstrated that CT induced hypersecretion by mainly acting directly on the mucosa, and also partly by increasing the excitability of cholinergic submucosal neurons specifically.
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While CT-exposure increased expression of activity dependent markers pCREB and c-Fos in submucosal and myenteric neurons respectively, whether these neurons are directly involved in the hypersecretion requires further investigation. Further, no apparent roles for VIP or myenteric neurons were found in the sustained effects of acute CT-treatment, but whether they are involved in the induction process remains to be determined. Additional experiments involving the co-incubation of CT with specific neural blockers will be important in addressing some of these questions.
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CHAPTER 6: DISCUSSION
Secretomotor circuits within the wall of the gastrointestinal tract are essential for controlling the balance of water and electrolytes in the body. These pathways are tightly regulated under physiological conditions. However, they are susceptible to invading pathogens which can release enterotoxins to induce hypersecretion, diarrhea, and dehydration. In severe cases, this hypersecretion can be fatal. Moreover, diarrhoeal disease remains a significant global health issue, affecting over a billion people annually (WHO, 2013). The two major subtypes of efferent secretomotor neurons are cholinergic (contain acetylcholine, ACh) and non-cholinergic (contain vasoactive intestinal peptide, VIP) neurons. VIP and its receptors have been of particular interest as VIP is implicated in CT- evoked hypersecretion, a representative model of bacterial-toxin induced diarrhoea, yet their roles within the neural circuitry have been unclear. Previous studies of VIP involvement in both physiological secretion (Cox and Cuthbert, 1989; Burleigh and Kirkham, 1993; Reddix et al., 1994; Banks et al., 2005) and pathophysiological secretion (Mourad and Nassar, 2000; Banks et al., 2005; Kordasti et al., 2006) have yielded conflicting results. Many of these discrepancies can be accounted for by the lack of selective pharmacological tools (Burleigh and Kirkham, 1993; Banks et al., 2005) and progress has also been hindered by technical limitations. Further, the relative contributions of cholinergic and VIP secretomotor neurons in regulating secretion in physiology and pathophysiology remain unclear. In order to develop more effective treatments for secretory diseases, these need to be clarified. Thus, my thesis aimed to address these questions. First, I examined the expression and function of VPAC receptors in the ENS by performing a combination of molecular, immunohistochemical, and pharmacological studies. Secondly, I investigated the role of VIP and VPAC receptors within the ENS using selective agonists and antagonists that are now available, as well as utilizing transgenic Wnt1-Cre;R26R- GCaMP3 mice, in which their enteric neurons and glia are encoded with a fluorescent Ca2+ indicator, for calcium imaging. Finally, I studied the effects of CT using an in vivo ileal loop model and again exploited Wnt1-Cre;R26R-GCaMP3 mice to assess the excitability of
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the enteric circuitry. In addressing these aims, a number of novel findings resulted and these contribute significantly to the understanding of physiological and pathophysiological secretion. A number of surprising findings also raised additional questions that need be tackled in future studies to further our understanding of intestinal function, but are beyond the scope of this thesis. Thus, this chapter will discuss the implications of the novel findings of this thesis and also highlight some future directions.
1.1 Neural VPAC1 receptors regulate intestinal secretion and muscle contractility
A key novel finding of this thesis is that functional VPAC1 receptors are expressed on cholinergic NPY+ secretomotor neurons and cholinergic excitatory longitudinal muscle motor neurons (Chapter 3) (Fung et al., 2014). While it is well known that VIP is a potent secretagogue and is released by a subset of secretomotor neurons to stimulate secretion through VPAC1 receptors on the mucosa, this is the first demonstration of a role of VIP in neurogenic secretion and that enteric neurons also express VPAC1 receptors. Further, VIP has been associated with inducing smooth muscle relaxation, and this is thought to be mediated by VPAC2 receptors (Laburthe et al., 2002; Harmar et al., 2004). However, the present data identified an additional role for VIP in stimulating longitudinal muscle contraction via VPAC1 receptors (Chapter 3) (Fung et al., 2014). Hence, these data demonstrate a dual role for VIP in the regulation of secretion and motility and suggest that VIP is an important mediator in the co-ordination of secretomotor and motility reflexes.
As VIP and VPAC1 receptors have been implicated in CT-induced hypersecretion (Banks et al., 2005), the possibility of their involvement not only at the level of the mucosa, but also within the enteric circuitry, was interesting. Modelling studies by Chambers et al. (2005) suggested that VIP submucosal neurons may become hyperexcitable by signaling via slow EPSPs in recurrent networks and may have a role in hypersecretion. However, the notion that VIP submucosal neurons innervate other VIP submucosal neurons, or even
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other submucosal neurons, has been a matter of contention. While the idea is supported by electrophysiological data (Mihara et al., 1985; Reed and Vanner, 2001), it is contested by immunohistochemical data (Costa and Furness, 1983; Furness et al., 2003). The findings that VPAC1 receptors are predominantly expressed on cholinergic neurons in guinea pigs and mice further refute the notion that VIP neurons innervate other VIP neurons (Chambers et al., 2005). Rather, VIP neurons likely provide inputs to cholinergic submucosal neurons. VPAC2 receptor expression on VIP neurons remains a possibility, but is yet to be demonstrated. Furthermore, there was no functional evidence to suggest that VPAC2 receptors are involved in VIP-evoked neurogenic secretion.
1.2 VIP and VPAC receptors regulate neuron-to-glia signaling
The major result of Chapter 4 was identifying a novel role for VIP and VPAC receptors in the modulation of enteric neuron-to-glia signaling. The study of the enteric neuro-glial circuitry was facilitated by the use of Wnt1-Cre;R26R-GCaMP3 mice. Local VIP application to submucosal ganglia stimulated neuronal Ca2+ responses, however selective inhibition of VPAC2 receptors surprisingly revealed Ca2+ responses in glia. Electrically stimulating endogenous VIP-release also evoked delayed glial Ca2+ responses when VPAC2 receptors were inhibited. Furthermore, activation of VPAC1 receptors on cholinergic submucosal neurons stimulated Ca2+ transients in enteric glia via purinergic signaling. The finding of purinergic involvement is consistent with the literature showing that ATP is a key signaling molecule in neuron-to-glia communication (Ruhl et al., 2004; Gulbransen and Sharkey, 2009). However, this is the first demonstration that VIP and VPAC receptors are involved in modulating the communication between neurons and glia in the ENS. These data also indicate that glial activity is tightly regulated by specific neuronal pathways in the ENS. In particular, it suggests that VIP acting on VPAC2 receptors may be involved in the tonic inhibition of glial activity.
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Notably, a number of neurons were seen to display Ca2+ transients following glial activation. Since glia contain neuroactive substances (Zhang et al., 2003; Ochoa-Cortes et al., 2016), the activated glia may be able to induce Ca2+ transients in neurons to modulate activity within the enteric circuitry as in the CNS (Hansson and Ronnnback, 2003). This is an area that has been relatively unexplored as selective activation of glia is challenging. However, recently McClain et al. (2015) applied a Designer Receptors Exclusively Activated by Designer Drugs (DREADDs) approach to show that selectively activating enteric glia can evoke neurogenic contractions and alter intestinal motility, although glia to neuron signaling was not specifically examined within the nerve plexus. Hence, it will be interesting to apply this model to investigate the possible mechanisms by which enteric glia may signal to or modulate the activity of neurons at the network level.
Enteric glia and neuron-glia signaling are currently of great interest given their multifaceted role in intestinal function and disease (Cabarrocas et al., 2003; Gulbransen and Sharkey, 2012; Ochoa-Cortes et al., 2016; Sharkey, 2016). This is the first demonstration that VIP is involved in the communication between enteric neurons and glia, and it will be interesting to further extend these findings. Information on the role of enteric glia in intestinal secretion is lacking. Studies examining the consequences of disrupting glial activity have predominantly focused on myenteric plexus, and highlight changes in motility but with minimal alterations to secretion (Nasser et al., 2006b; McClain et al., 2014; MacEachern et al., 2015; Brown et al., 2016). Although, MacEachern and colleagues (2015) have reported a VIP-evoked neurogenic secretory response which was only observed when glial activity was perturbed in colitis. Whether a similar neuron-glia interaction involving VPAC1 receptors as described in the small intestine is involved in pathological conditions remains to be elucidated. Further it was reported that neurally-mediated electrogenic ion transport was depressed in colitis, but was partially restored by inhibiting glial activity (MacEachern et al., 2015). Thus, future studies examining enteric glia in the submucosal plexus are crucial for a better understanding of their role in the regulation of intestinal secretion, particularly in inflammatory bowel disease.
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While the studies in the thesis have added to the understanding of VIP and VPAC1 receptors within the enteric circuitry, questions regarding VPAC2 receptors remain. In chapter 4, VIP evoked neuronal Ca2+ transients were shown to involve both VPAC1 and VPAC2 receptors using selective antagonists, thus suggesting that both receptor subtypes are expressed on neurons. This is also compatible with the qPCR data on guinea pig tissues demonstrating a high level of Vipr2 expression in the submucosal plexus relative to all other intestinal layers (Chapter 3). However, the functional studies in guinea pig did not suggest any neural roles for VPAC2 in secretion or muscle contractility. Furthermore, the VPAC2 agonist did not evoke any responses in the submucosal plexus in Ca2+ imaging studies. Admittedly, neuronal activity was inferred from Ca2+ responses and it is possible that examining electrophysiological responses may have provided an answer. Localization of VPAC2 receptors was also attempted using immunohistochemistry, but the antisera applied did not provide any specific labelling. Thus, in order to better understand the VIP/VPAC neuron-to-glia signaling pathway, it will be important to establish whether VPAC2 receptors are indeed expressed on enteric neurons and, if so, how their pattern of expression relates to that of VPAC1 receptors.
1.3 ENS involvement in the sustained effects of acute cholera toxin exposure
The findings in Chapter 5 showed that the sustained CT-evoked hypersecretory effects in mouse ileum mainly involve a direct effect on the mucosal epithelium, and partly involve an increased excitability of cholinergic submucosal neurons. This was done using an in vivo ileal loop model of CT-incubation and I also used Wnt1-Cre;R26R-GCaMP3 mice to assess the excitability of the enteric circuitry. This was the first study to examine both the submucosal plexus and myenteric plexus in CT-evoked hypersecretion. Although an increased excitability of cholinergic secretomotor neurons was observed following CT- incubation, the myenteric plexus appears to play a minor role, at least in the sustained hypersecretory effects. Further, electrogenic secretory responses to agonist (DMPP and veratridine) stimulation were unaltered. These findings were unexpected given the
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extensive body of work conducted on cats and rats in vivo demonstrating that there is a significant neural contribution to CT-induced hypersecretion (Cassuto et al., 1982a; Cassuto et al., 1983; Jodal et al., 1993; Sjöqvist et al., 1993; Lundgren and Jodal, 1997; Mourad and Nassar, 2000; Kordasti et al., 2006). Yet, the majority of studies on rabbits and guinea pigs where secretion was measured in vitro suggest that CT induces hypersecretion mainly through direct effects on the mucosa (Field et al., 1971; Field et al., 1972; Carey and Cooke, 1986; Moriarty et al., 1989). While the studies conducted in this thesis involved CT-incubation in vivo, secretion was ultimately measured in vitro. Whether the differences observed between these studies are due to technical or species differences remain to be determined. Thus, in future studies, it may be necessary to measure secretion in vivo and to examine the neural contribution using local anesthetics during CT-incubation to verify that there is indeed a major direct mucosal effect in the mouse.
The current understanding of neurogenic secretion of mouse small intestine is limited and generalizations are often made based on knowledge of the guinea pig small intestine. However, it is important to consider potential species differences. Ussing chamber studies of mouse jejunum using full thickness preparations showed that the nicotinic agonist DMPP evoked a secretory response that was unaffected by the muscarinic antagonist atropine (Sheldon et al., 1989). This suggests that nicotinic activation preferentially excites non-cholinergic secretomotor neurons to stimulate secretion. Similarly, veratrine (a mixture of Veratrum alkaloids including veratridine), which activates Na+ channels, also evoked a secretory response that was insensitive to muscarinic blockade (Sheldon et al., 1990; Hirota and McKay, 2006). By contrast, DMPP and veratridine have been shown to stimulate neurogenic secretory responses that do involve muscarinic receptors in guinea pig small intestine (Keast et al., 1985b; Foong et al., 2010a). The reason for these apparently conflicting findings is unclear as both agonists are expected to activate both cholinergic and non-cholinergic secretomotor neurons either directly, or indirectly. It is possible that the wiring of the mouse small intestine is different, such that cholinergic secretomotor neurons receive more inhibition and thus the summation of excitatory and inhibitory inputs results
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in a net depression of their activity. Indeed, there is evidence of differences between the regulation of neural secretion of mice and guinea pigs, where muscarinic blockade does not reduce basal ISC in the mouse small intestine (Carey and Cooke, 1989a; Sheldon et al., 1989), but is effective in the guinea pig small intestine (Fang et al., 2006; Fung et al., 2014). Yet, potential species differences do not explain why electrically-stimulated secretory responses in mouse jejunum involve muscarinic receptors (Carey and Cooke, 1989a; Hirota and McKay, 2006). Nonetheless, these findings may help explain the lack of differences observed between the DMPP- and veratridine-evoked secretory responses in CT-treated and control preparations despite a change in the excitability of cholinergic submucosal neurons. Ultimately, this highlights the need to better characterize the mouse enteric circuitry, particularly as they are animal model of choice for a range of diseases.
CT was found to exert its sustained effects partly via cholinergic submucosal neurons in the mouse ileum. This contrasts with previous work on CT-evoked hypersecretion, which largely implicates the involvement of VIP and non-cholinergic secretomotor neurons (Cassuto et al., 1982a; Mourad and Nassar, 2000; Banks et al., 2005; Kordasti et al., 2006; Gwynne et al., 2009). However, it has been proposed that the induction and maintenance of CT-evoked hypersecretion involve different neural mechanisms (Kordasti et al., 2006). Thus, it is possible that the maintenance of CT-evoked hypersecretion involves cholinergic submucosal neurons, while VIP secretomotor neurons may have a more prominent role in its induction. Further investigations examining the effects of co-incubating CT with various neural blockers, including specific antagonists for muscarinic and VPAC receptors, will be necessary to better understand the neural involvement in the induction of CT-induced hypersecretion. However, taking into account that VPAC antagonists are typically peptides with a short half-life (Couvineau and Laburthe, 2012), the 3.5 h CT incubation in a ligated ileal loop may not be the ideal model for these studies. Rather, constant luminal perfusion with the antagonist may be necessary.
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1.4 The role of cholinergic and non-cholinergic secretomotor neurons in the regulation of intestinal secretion
The data presented in Chapter 5 highlighted a role for cholinergic submucosal neurons in the sustained hypersecretory effects of CT-exposure. This was interesting since muscarinic antagonists are ineffective in inhibiting CT-evoked hypersecretion (Cassuto et al., 1982a; Kobayashi et al., 2001; Kordasti et al., 2006; Gwynne et al., 2009). However, a recent and extensive study on human biopsies has demonstrated that cholinergic, rather than non- cholinergic secretomotor neurons, largely contribute to electrically-stimulated neurogenic secretion in the small intestine (Krueger et al., 2016). Yet, human studies of CT suggests a major VIP involvement (Bloom et al., 1976; Banks et al., 2005). Whether CT-exposure shifts the balance to VIP-mediated secretion remains to be determined. Nevertheless, it is apparent from the collective findings of this thesis that there is a degree of interaction between cholinergic and VIP signaling within the ENS; especially considering VPAC receptors are predominantly expressed on cholinergic neurons in both the guinea pig and mouse. This also raises the possibility that some of the reported effects of VIP- or VPAC1 antagonists in attenuating CT-evoked hypersecretion may in fact involve cholinergic neurons.
1.5 Conclusion
This thesis presented the first evidence that enteric neurons express VPAC1 receptors, and VIP acting via VPAC1 receptors regulate neurogenic secretion and longitudinal muscle contractility. These findings suggest that VIP is probably involved in the co-ordination of secretomotor and motility reflexes within the ENS. Further, I identified an additional role of VIP as a transmitter involved in enteric neuron-glia communication. The novel VIP/VPAC neuron-to-glia signaling pathway identified provided some insights into the complexity of the enteric circuitry. However, these findings also raised questions regarding the functional role of glia in intestinal secretion and the possibility of glia-to-neuron
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signaling. These will be interesting avenues to pursue in future studies of the involvement of glia in gastrointestinal diseases. An interesting finding was that VPAC1 receptors are predominantly localized to cholinergic neurons in guinea pigs and mice, which suggests that there is a degree of interaction between VIP and acetylcholine within the ENS and in the regulation of intestinal secretion. Further, despite the focus on VIP secretomotor neurons, it is clear from the data presented in this thesis that cholinergic submucosal neurons have a more prominent role in the regulation of pathophysiological intestinal secretion than previously thought. To date, studies of intestinal secretion have considered cholinergic and VIP secretomotor pathways as being distinctly separate. Thus, a consideration for future investigation is the possibility that these separate populations of secretomotor neurons may be linked using VIP as a neurotransmitter. Finally, it will be important to better characterize cholinergic secretomotor pathways to gain a more complete understanding of the regulation of intestinal secretion.
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Minerva Access is the Institutional Repository of The University of Melbourne
Author/s: Fung, Candice
Title: The role of submucosal neurons in physiological and pathophysiological intestinal secretion
Date: 2016
Persistent Link: http://hdl.handle.net/11343/123274
File Description: The Role of Submucosal Neurons in Physiological and Pathophysiological Intestinal Secretion
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