University of Nevada, Reno

Mechanisms of release, metabolism and action of purines in the enteric and central nervous systems

A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Cellular and Molecular Pharmacology and Physiology

by

Leonie Durnin

Violeta N. Mutafova-Yambolieva, M.D, Ph.D/Dissertation Advisor August 2013

© by Leonie Durnin 2013 All Rights Reserved

THE GRADUATE SCHOOL

We recommend that the dissertation prepared under our supervision by

LEONIE DURNIN

entitled

Mechanisms of release, metabolism and action of purines in the enteric and central nervous systems

be accepted in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Violeta Mutafova-Yambolieva, MD,PhD, Advisor

Kenton M. Sanders, Ph.D, Committee Member

Sean M. Ward, Ph.D, Committee Member

Brian A. Perrino, Ph.D, Committee Member

Thomas Kidd, Ph.D, Graduate School Representative

Marsha H. Read, Ph. D., Dean, Graduate School

August, 2013

i

Abstract

It has been fifty years since the first descriptions of non-adrenergic non- (NANC) were made in the gastrointestinal (GI) tract and more than forty years since it was discovered that NANC neurotransmission was mediated by a purine nucleotide. These influential advances in understanding neuroeffector mechanisms in led Dr. Geoffrey Burnstock to coin the terms ‘purinergic co-transmission’ and ‘purinergic ’. This led to an explosion of interest in purinergic mechanisms controlling the activity of various smooth muscle organs.

While adenosine 5’-triphosphate (ATP) has traditionally been considered to be the purine nucleotide responsible for NANC responses, the last decade has uncovered important additional purinergic mechanisms controlling smooth muscle function. Based on the results obtained using extremely sensitive high-pressure liquid chromatography

(HPLC) methodologies, nicotinamide adenine dinucleotide (NAD+) has emerged as a novel extracellular signaling factor released during stimulation of peripheral nerves and has a putative or neuromodulator role. These original findings have significantly advanced our understanding in the field of purinergic signaling.

There is substantial evidence that stimulation-evoked release of NAD+ in smooth muscles originates from neural sources, however the complex organization of smooth muscle organs makes verification of vesicular NAD+ release a challenging task.

Thus in Chapter 2 of this dissertation we utilized a single cell model to examine storage and release of NAD+ from vesicles in nerve-growth factor (NGF)-differentiated rat ii pheochromocytoma PC12 cells which phenotypically resemble sympathetic . In this study we verified the presence of NAD+ in vesicles along with ATP and catecholamines (dopamine). Interestingly, we revealed differential mechanisms of release of these three substances from vesicles: release of NAD+ and dopamine required intact

SNAP-25-mediated whereas ATP was released largely via SNAP-25- independent mechanisms. These observations in conjunction with a previous finding demonstrating ω-conotoxin GVIA-insensitive ATP release in blood vessels led us to question the true identity of the NANC neurotransmitter in GI muscles where purinergic neurotransmission was first described.

In the GI tract, purines released from inhibitory motor neurons elicit postsynaptic hyperpolarization transients (inhibitory junction potentials, IJPs) in circular smooth muscles causing relaxation. In the attempt to clarify which purines are involved in mediating gut relaxation we carried out a series of experiments in murine and primate colonic muscles comparing mechanisms of release, metabolism and action of extracellular purines. In Chapter 3 we demonstrated that electrical field stimulation (EFS) evoked release of NAD+ that was dependent on the level of nerve stimulation and was significantly attenuated by blockers of neural activity. We also demonstrated that postsynaptic hyperpolarizations to exogenous NAD+ were abolished by factors inhibiting the endogenous purine-mediated IJP. On the other hand, release and postsynaptic effects of ATP remained largely intact by inhibitors of enteric purine neurotransmission.

Therefore our evidence suggests that NAD+ is a better candidate than ATP as the purinergic inhibitory motor transmitter in colons from humans and non-human primates. iii

Extracellular nucleotidases in the gut degrade purines in the extracellular compartment. Rapid metabolism of ATP once released might explain the discrepancies between exogenous ATP and the endogenous enteric purine transmitter. In Chapter 4 we examined postjunctional effects of direct metabolites of ATP and NAD+, adenosine 5’- diphosphate (ADP) and ADP-ribose (ADPR), respectively, in colonic muscles. First, we demonstrated that these metabolites are produced very rapidly in murine and primate colons (within 1 sec). Next, we found that membrane hyperpolarizations to ADPR, but not to ADP, mimicked the pharmacology of endogenous purine response; this is the first study demonstrating a bioactive role of ADPR in enteric smooth muscles. Our evidence indicates that rapid metabolism cannot explain the failure of ATP to match the endogenous transmitter in colon. Moreover our evidence suggests that multiple purines might contribute to enteric inhibitory responses produced during NANC neurotransmission. Thus purinergic inhibitory regulation of enteric smooth muscle is more complex than originally believed.

EFS is a common approach for stimulating neural activity however it often fails to differentiate the precise sources of released molecules. For example, during EFS substances could be released from neuronal cell bodies or or from non-neuronal sources such as . In Chapter 5 we attempted to clarify the sites of release of ATP and

NAD+ in GI smooth muscles by utilizing an alternative approach to stimulate purine release. Here we chemically activated the neuronal ligand-gated receptors, nicotinic acetylcholine receptors and serotonin 5-HT3 receptors, which are localized on cell bodies and of inhibitory motor neurons. We demonstrated that the release of ATP and NAD+ upon activation of these receptors originated from different sites iv within neurons and via different mechanisms. The release of NAD+ appeared to originate exclusively from nerve terminals and was abolished by neural inhibitors. However the release of ATP remained intact in the presence of neural inhibitors suggesting that ATP release may have originated primarily from the nerve cell bodies. Therefore in agreement with our previous studies release of NAD+ in the gut occurs by mechanisms consistent for a neurotransmitter.

With the strong evidence supporting NAD+ as an enteric inhibitory neurotransmitter we also wanted to determine if NAD+ fulfills neurotransmitter criteria in the (CNS). In Chapter 6 we examined release, metabolism and postjunctional effects of NAD+ in the rat brain. We demonstrated that in isolated rat forebrain synaptosomes NAD+ is released by mechanisms requiring intact vesicle exocytosis machinery. We also found that localized application of NAD+ (and ADPR) elicited Ca2+ transients in cultured cortical neurons suggesting that endogenous NAD+ could participate in neuronal-neuronal communication in the brain. Finally, we demonstrated that mechanisms involved in terminating the extracellular action of NAD+ exist in the brain. This is the first study suggesting that NAD+ qualifies as a putative neurotransmitter in the CNS.

In summary, the work described in this dissertation provides novel information regarding the role of NAD+/ADPR in the enteric and central nervous systems.

NAD+/ADPR qualify as enteric inhibitory motor regulating colonic smooth muscle contractility. In addition, NAD+/ADPR may participate in neurotransmission in the CNS. Our findings are significant not only to further our understanding of complex purinergic mechanisms regulating central and enteric nervous v system functions but also afford the opportunity for selectively targeting the

NAD+/ADPR system for the therapeutic treatment of pathological conditions resulting from altered purinergic signaling.

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Acknowledgements

First, I would like to express my sincere thanks to my advisor Dr. Violeta

Mutafova-Yambolieva for her wonderful guidance, insight and patience throughout my graduate studies at the University of Nevada. She has been a fantastic supervisor who has taught me so much throughout my time in her lab. I feel extremely lucky to have been afforded this opportunity to work in her laboratory and I am grateful for everything she has helped me with to get where I am today. Her enthuasism for science has truly made this journey a very enjoyable experience. Thank you!

I would also like express my thanks to Dr. Sean Ward not only for his help and guidance over the years, but also for giving me the chance to come to Reno as an undergraduate student to gain experience in the research environment. It was this opportunity that inspired me to climb higher and pursue this Ph.D.

Sincere thanks to the other members of my advisory committee, Dr. Kenton

Sanders, Dr. Brian Perrino and Dr. Thomas Kidd. They have always been supportive and have always found time in their busy schedules to give me advice, for which I am extremely grateful.

I would also like to thank everyone in the department of Physiology and the

Cellular and Molecular Pharmacology and Physiology Graduate Program. I would like to acknowledge Debi Russell, Sung Jin Hwang, Ilia Yamboliev, Yanping Dai, Sang Don

Koh, Laura Dwyer, Yulia Bayguinov and Lauren Peri for their contributions to various projects throughout my studies. Thanks also to Charles River Preclinical Services and

Renown and St. Mary’s hospitals for generous donations of primate tissues. Special vii thanks to Nancy Horowitz for collecting primate tissues and maintaining our colonies of mice.

I would like to thank the many friends I have made in Reno who have made this experience all the more worthwhile. I feel very lucky to have met so many kind and supportive friends along the way.

I am especially grateful to my parents, Leonard and Kathleen. They have always been supportive of everything I have done. Thank you for your help, encouragement and love throughout.

Finally I would like to offer a special thanks to my fiancé Robert Corrigan. I cannot express with words how lucky I am to have had your support, encouragement and love through every step of this Ph.D. Thank you!

viii

Publications in this dissertation

* denotes joint first author

Yamboliev IA, Smyth LM, Durnin L, Dai Y, & Mutafova-Yambolieva VN (2009).

Storage and secretion of beta-NAD, ATP and dopamine in NGF-differentiated rat pheochromocytoma PC12 cells. Eur J Neurosci 30, 756-768.

Hwang SJ*, Durnin L*, Dwyer L, Rhee PL, Ward SM, Koh SD, Sanders KM, &

Mutafova- Yambolieva VN (2011). [beta]-Nicotinamide Adenine Dinucleotide Is an

Enteric Inhibitory Neurotransmitter in Human and Nonhuman Primate Colons.

Gastroenterology 140, 608-617.

Durnin L*, Hwang SJ*, Ward SM, Sanders KM, & Mutafova-Yambolieva VN (2012b).

Adenosine 5-diphosphate-ribose is a neural regulator in primate and murine large intestine along with beta-NAD(+). J Physiol 590, 1921-1941.

Durnin L, Dai Y, Aiba I, Shuttleworth CW, Yamboliev IA, & Mutafova-Yambolieva

VN (2012a). Release, neuronal effects and removal of extracellular beta-nicotinamide adenine dinucleotide (beta-NAD) in the rat brain. Eur J Neurosci 35, 423-435.

Durnin L, Sanders KM, & Mutafova-Yambolieva VN (2013). Differential release of beta-NAD(+) and ATP upon activation of enteric motor neurons in primate and murine colons. Neurogastroenterol Motil 25, e194-e204. ix

Other publications related to purinergic neurotransmission in smooth muscles

Durnin L & Mutafova-Yambolieva VN (2011). Cyclic ADP-ribose requires CD38 to regulate the release of ATP in visceral smooth muscle. FEBS J 278, 3095-3108.

Hwang SJ, Blair PJ, Durnin L, Mutafova-Yambolieva V, Sanders KM, & Ward SM

(2012). P2Y1 purinoreceptors are fundamental to inhibitory motor control of murine colonic excitability and transit. J Physiol 590, 1957-1972.

Roberts JA, Durnin L, Sharkey KA, Mutafova-Yambolieva VN, & Mawe GM (2013).

Oxidative Stress Disrupts Purinergic Neuromuscular Transmission in the Inflamed Colon.

J Physiol 591, 3725-3737.

x

Table of Contents

CHAPTER 1: Introduction ...... 1

1.1 Chemical synaptic transmission and criteria to define a neurotransmitter ...... 2

1.2 Purinergic non-adrenergic non-cholinergic (NANC) neurotransmission in smooth muscle ...... 3

1.3 Difficulties associated with the study of purinergic neurotransmission in smooth muscles ...... 8

1.3.1 Variable purine release mechanisms ...... 9

1.3.2 Complex metabolic pathways of extracellular purines ...... 9

1.3.3 Abundant purine receptor expression ...... 11

1.3.4 Inadequate methodologies for purine detection ...... 12

1.4 Evidence for NAD+ as a NANC neurotransmitter in peripheral smooth muscles ...... 13

1.4.1 High-pressure liquid chromatography (HPLC) techniques for sensitive

detection of purines ...... 13

1.4.2 Extracellular actions of NAD+ ...... 14

1.4.3 Emergence of NAD+ as a putative neurotransmitter or neuromodulator in

peripheral smooth muscles ...... 14

1.5 A single-cell model to characterize storage and release of NAD+ ...... 18

1.6 Purinergic neuromuscular transmission in the ...... 20

1.6.1 Brief overview of the ...... 20

1.6.2 Purines in the enteric nervous system ...... 25

1.6.2.1 Regulated release from neuronal sources ...... 26

1.6.2.2 Regulated release from non-neuronal sources ...... 28 xi

1.6.2.3 Non-regulated release from damaged and inflamed tissues ...... 28

1.7 Road to identifying the enteric purine motor transmitter...... 29

1.7.1 Characterization of the enteric purine neurotransmitter ...... 31

1.7.2 The role of purine metabolites in the neuroeffector junction ...... 33

1.7.3 Differential sites of release of ATP and NAD+ in the .. 35

1.7.4 Differential sites for degradation of ATP and NAD+ in the colon ...... 36

1.7.5 Effects of purines in the colon of P2Y1R-/- mice ...... 37

1.8 Targets of neurogenic purines in the GI tract ...... 38

1.9 Non-synaptic transmission ...... 40

1.10 Purinergic neurotransmission in the central nervous system ...... 44

1.10.1 Overview of purinergic signaling in the central nervous system ...... 44

1.10.2 NAD+ as a putative neurotransmitter in the central nervous system ...... 45

1.11 Summary ...... 48

1.12 References ...... 51

CHAPTER 2: Storage and secretion of β-NAD, ATP and dopamine in NGF-

differentiated rat pheochromocytoma PC12 cells ...... 71

2.1 Abstract ...... 72

2.2 Introduction ...... 73

2.3 Materials and Methods ...... 75

2.3.1 Cell culture ...... 75

2.3.2 Overflow experiments ...... 75

2.3.3 Treatment with BoNT/A ...... 76 xii

2.3.4 Fractionation of synaptic vesicles by glycerol and sucrose gradient

centrifugation ...... 76

2.3.5 Preparation of samples for purine detection ...... 77

2.3.6 HPLC assay of etheno-purines ...... 78

2.3.7 Sample concentration and HPLC fraction analysis ...... 79

2.3.8 Preparation of samples for catecholamine detection ...... 80

2.3.9 HPLC assay of catecholamines ...... 80

2.3.10 Western immunoblot analysis of SNAP-25 in total PC12 cell extracts.... 81

2.3.11 Western immunoblot analysis of vesicular protein markers and SNAP-25

in glycerol and sucrose gradient fractions ...... 82

2.3.12 Statistics ...... 82

2.4 Results ...... 83

2.4.1 Spontaneous secretion of catecholamines and purines in NGF-treated

PC12 cells ...... 83

2.4.2 Secretion of catecholamines and purines evoked by 60 mM KCl and

effects of BoNTA ...... 83

2.4.3 Secretion evoked by nicotine 100 μM ...... 86

2.4.4 Distribution of β-NAD, ADPR, cADPR in cell superfusates determined by

HPLC fraction analysis ...... 87

2.4.5 Fractionation of synaptic vesicles by glycerol gradient: neural markers,

content of DA and purines, HPLC fraction analysis of β-NAD, ADPR,

and cADPR ...... 88 xiii

2.4.6 Fractionation of synaptic vesicles by sucrose gradient: neural markers,

content of DA and purines, HPLC fraction analysis of β-NAD, ADPR,

and cADPR ...... 89

2.4.7 Western immunoblot analysis of SNAP-25 in glycerol and sucrose

gradient centrifugation fractions ...... 90

2.5 Discussion...... 104

2.6 References ...... 111

CHAPTER 3: β-nicotinamide adenine dinucleotide (β-NAD) is an enteric inhibitory

neurotransmitter in human and non-human primate colons ...... 116

3.1 Abstract ...... 117

3.2 Introduction ...... 118

3.3 Materials and Methods ...... 119

3.3.1 Tissue Preparation ...... 119

3.3.2 Purine overflow ...... 120

3.3.3 HPLC Assay of Etheno-purines ...... 120

3.3.4 HPLC Fraction Analysis ...... 121

3.3.5 Electrophysiology and Contractions ...... 121

3.3.6 Expression of P2Y receptors ...... 122

3.3.7 NADPH Diaphorase Histochemistry ...... 122

3.3.8 Data Analysis ...... 123

3.3.9 Drugs ...... 123

3.4 Results ...... 124 xiv

3.4.1 Neural release of purines ...... 124

3.4.2 Purinergic component of inhibitory junction potentials (IJPs) ...... 125

3.4.3 Effects of β-NAD and ATP on SMC conductance ...... 127

3.5 Discussion...... 138

3.6 Supplementary material ...... 144

3.6.1 Supplemental Experimental Procedures ...... 144

3.6.2 Supplemental Results ...... 145

3.7 References ...... 154

CHAPTER 4: Adenosine 5’-diphosphate-ribose is a neural regulator in primate and murine large intestine along with β-NAD+ ...... 159

4.1 Abstract ...... 160

4.2 Introduction ...... 161

4.3 Materials and Methods ...... 163

4.3.1 Ethical procedures ...... 163

4.3.2 Tissue preparation ...... 164

4.3.3 Degradation of purine nucleotides in mouse and monkey colonic smooth

muscle ...... 164

4.3.4 HPLC assay of etheno-nucleotides/nucleosides and cGDPR ...... 167

4.3.5 HPLC fraction analysis ...... 167

4.3.6 Preparation of etheno-substrates ...... 168

4.3.7 Preparation of cGDPR ...... 168

4.3.8 Intracellular electrical activity and force measurements ...... 169 xv

4.3.9 RNA isolation and RT-PCR...... 170

4.3.10 Western immunoblot analysis ...... 170

4.3.11 Statistics ...... 171

4.3.12 Drugs and chemicals ...... 172

4.4 Results ...... 172

4.4.1 β-NAD+ forms ADPR in primate colonic smooth muscles ...... 172

4.4.2 β-NAD+ is likely to be degraded by multiple enzymes in murine colon .175

4.4.3 Extracellular ATP is degraded in the colon ...... 176

4.4.4 EFS-induced IJPs are sensitive to apamin and P2Y1 receptor inhibition in

the monkey circular smooth muscle cells ...... 178

4.4.5 Hyperpolarization responses to localized application of exogenous ADPR

and ADP ...... 179

4.4.6 Electrophysiological evidence that degradation of β-NAD+ does not

exclusively require CD38 ...... 180

4.4.7 Mechanisms for ADPR degradation are present in the colon ...... 181

4.5 Discussion...... 203

4.6 Supplementary material ...... 211

4.7 References ...... 212

CHAPTER 5: Differential release of β-NAD+ and ATP upon activation of enteric motor neurons in primate and murine colons ...... 217

5.1 Abstract ...... 218

5.2 Introduction ...... 219 xvi

5.3 Materials and Methods ...... 220

5.3.1 Tissue preparation ...... 220

5.3.2 Purine overflow ...... 221

5.3.3 HPLC assay of purines in tissue superfusates...... 222

5.3.4 HPLC fraction analysis ...... 222

5.3.5 Statistics ...... 223

5.3.6 Drugs ...... 223

5.4 Results ...... 224

5.4.1 Release of ATP and β-NAD+ elicited by activation of nAChR ...... 224

+ 5.4.2 Release of ATP and β-NAD elicited by activation of 5-HT3R ...... 226

5.5 Discussion...... 238

5.6 References ...... 243

CHAPTER 6: Release, neuronal effects and removal of extracellular β-nicotinamide

adenine dinucleotide (β-NAD+) in the rat brain ...... 248

6.1 Abstract ...... 249

6.2 Introduction ...... 250

6.3 Materials and Methods ...... 252

6.3.1 Animals ...... 252

6.3.2 Preparation of synaptosomes ...... 252

6.3.3 HPLC assay of etheno-purines ...... 253

6.3.4 Intrasynaptosomal content of purines ...... 254

6.3.5 Spontaneous secretion of purines by synaptosomes ...... 255 xvii

6.3.6 High K+-evoked overflow ...... 255

6.3.7 HPLC fraction analysis ...... 256

6.3.8 Degradation of β-NAD+ and ATP ...... 256

6.3.9 Uptake of NAD+ ...... 257

6.3.10 Western immunoblot analysis ...... 258

6.3.11 Cell culture ...... 258

6.3.12 Imaging of intracellular Ca2+ transients ...... 259

6.3.13 Statistics ...... 260

6.3.14 Drugs ...... 260

6.4 Results ...... 261

6.4.1 Intra-synaptosomal content of purines and spontaneous release of

endogenous purines from rat brain synaptosomes ...... 261

6.4.2 High-K+-evoked release of purines ...... 262

6.4.3 Degradation of ATP and NAD+ ...... 264

6.4.4 Expression of CD38 and -associated proteins ...... 265

6.4.5 Uptake of NAD+ ...... 265

6.4.6 Neuronal Ca2+ dynamics ...... 266

6.5 Discussion...... 285

6.6 Supplementary material ...... 291

6.7 References ...... 293

CHAPTER 7: Summary and Conclusions ...... 299 xviii

List of Tables

Table S3.1 Oligonucleotide Primers for P2Y Receptors ...... 151

Table S3.2 Human Proximal Colon ...... 152

Table S3.3 Monkey Proximal Colon ...... 153

Table 4.1 Degradation of eADO (0.05 μM) by monkey WM, monkey CM and

murine colon under control conditions or in the presence of either EHNA

(10 μM), NBMPR (10 μM), or EHNA + NBMPR combined ...... 200

Table 4.2 Effects of antagonists to neural responses and exogenous ADPR and ADP

in colons isolated from wild type CD38+/+ mice ...... 201

Table 4.3 Effects of apamin and MRS2500 on neural responses and exogenous

purines in colons isolated from CD38-/- mice ...... 202

Table 6.1 Intra-synaptosomal content and spontaneous overflow of purines in rat

brain synaptosomes (pmol/mg protein) ...... 284

xix

List of Figures

Figure 1.1 Sucrose gap recording of membrane potential changes in smooth muscle

of guinea pig taenia coli ...... 6

Figure 1.2 Biotransformation pathways for extracellular purines ...... 10

Figure 1.3 HPLC-fluorescence detection analysis of purine release upon EFS ...... 17

Figure 1.4 The organization of the ENS of human and medium-large mammals ....23

Figure 1.5 Simplified neural circuits responsible for the peristaltic reflex ...... 24

Figure 1.6 Ganglionic stimulation of motor neurons causes release of β-NAD+ at

nerve varicosities ...... 36

Figure 1.7 A simplified model of purine-mediated neurotransmission in the colon

smooth muscle neuroeffector junction ...... 43

Figure 1.8 The known signaling functions of extracellular NAD(P) ...... 50

Figure 2.1 Protocol for isolation of SSVs and LDCVs from cultured NGF-

differentiated rat PC12 cells...... 91

Figure 2.2 Spontaneous and high-K+-evoked secretion of DA in NGF-treated PC12

cells ...... 92

Figure 2.3 Spontaneous and high-K+-evoked release of adenine purines in NGF-

treated PC12 cells ...... 94

Figure 2.4 Spontaneous and nicotine (100 µM)-evoked release of adenine purines in

NGF-treated PC12 cells ...... 96

Figure 2.5 HPLC fraction analysis of the mixture of β-NAD, ADPR and cADPR ..97

Figure 2.6 Fraction separation of small synaptic vesicles (SSV) by glycerol gradient

centrifugation ...... 99 xx

Figure 2.7 Fraction separation of large dense-core vesicles (LDCVs) by sucrose

gradient centrifugation ...... 101

Figure 2.8 Western immunoblot analysis of SNAP-25 in fractions isolated by

glycerol and sucrose gradient centrifugation ...... 103

Figure 3.1 Overflow of ATP and β-NAD in human colonic muscle ...... 129

Figure 3.2 ATP and β-NAD are released in monkey whole and circular muscle .....131

Figure 3.3 Purinergic component of IJPs ...... 133

Figure 3.4 Membrane responses to exogenous purines ...... 134

Figure 3.5 Expression of P2Y receptors in monkey and human tissues ...... 136

Figure 3.6 β-NAD and ATP activate inward currents in SMCs of monkey and

human ...... 137

Figure S3.1 Inhibitory neural regulation of colonic muscles ...... 147

Figure S3.2 Concentration-dependent inhibition of monkey and human colonic

muscles by β−nicotinamide adenine dinucleotide (β-NAD) ...... 149

Figure 4.1 Degradation of eNAD in monkey whole muscle (WM) and circular muscle

(CM) colon preparations ...... 183

Figure 4.2 Degradation of eNAD in colon preparations isolated from wild-type and

CD38-/- mice ...... 184

Figure 4.3 Degradation of eNAD after brief contacts with murine colon ...... 185

Figure 4.4 Degradation of NGD in colon preparations isolated from wild-type and

CD38-/- mice ...... 186

Figure 4.5 Degradation of eATP in colon preparations isolated from monkey and

murine large intestine ...... 187 xxi

Figure 4.6 Inhibitory junction potentials and effects of ADPR and ADP on membrane

potential of monkey colonic muscles ...... 189

Figure 4.7 Concentration-response relationship for ADPR and ADP on membrane

hyperpolarizations in murine colon ...... 191

Figure 4.8 Effects of ADPR and ADP on membrane potential and inhibitory junction

potentials of colonic muscle cells from wild-type CD38+/+ mice ...... 193

Figure 4.9 Effects of β-NAD+, ATP, ADPR and ADP on membrane potential and

inhibitory junction potentials of colonic circular muscles from CD38-/-

mice ...... 195

Figure 4.10 Degradation of eADPR in colon preparations isolated from monkey and

murine large intestine ...... 197

Figure 4.11 Superposition of inhibitory junction potential (IJP) and hyperpolarization

responses to exogenous ATP and β-NAD spritzed near the site of

recording in a murine colonic preparation ...... 199

Figure S4.1 ADPR inhibited spontaneous contractions of colonic muscles cut parallel

to the circular muscle fibers ...... 211

Figure 5.1 Stimulation of nicotinic acetylcholine receptors (nAChRs) causes release

of purines in monkey colon whole muscle preparations ...... 229

Figure 5.2 Release of ATP and β-NAD+ during nicotinic acetylcholine receptor

(nAChR) stimulation in whole muscle (WM) and circular muscle (CM)

preparations of monkey colon ...... 231

Figure 5.3 ATP and -NAD+ release during stimulation of nicotinic acetylcholine

receptors (nAChRs) and 5-HT3Rs in mouse colon ...... 232 xxii

Figure 5.4 Purine release by stimulation of 5-HT3 receptors in monkey colon whole

muscle (WM) preparations ...... 233

+ Figure 5.5 Release of ATP and -NAD during 5-HT3 receptor stimulation in whole

muscle (WM) and circular muscle (CM) preparations of monkey

colon ...... 235

Figure 5.6 Sites of release of ATP and -NAD+ in response to stimulation of

nicotinic acetylcholine receptor (nAChR) and 5-HT3R in colonic

muscles ...... 236

Figure 6.1 Protocol for isolation of synaptosomes from rat forebrain ...... 269

Figure 6.2 Diagram of major enzymatic pathways for the degradation of β-NAD+

and ATP ...... 270

Figure 6.3 Spontaneous release and high-K+-evoked release of adenine purines .....271

Figure 6.4 Effects of botulinum neurotoxin A (BoNT/A), bafilomycin A1 and ω-

conotoxin GVIA (ω-Ctx GVIA) on spontaneous and high-K+-evoked

release of adenine purines ...... 272

Figure 6.5 Degradation of exogenous purine substrates during superfusion of

synaptosomes ...... 274

Figure 6.6 Degradation of eNAD and expression of CD38 ...... 276

Figure 6.7 Uptake of eNAD ...... 278

Figure 6.8 Intracellular Ca2+ increases in cultured neurons following localized

application of β-NAD+ ...... 280

2+ Figure 6.9 Similar neuronal [Ca ]i transients with ADPR and ATP ...... 282 xxiii

Figure S6.1 Control showing lack of effect of pressure ejection delivery method on

Ca2+ dynamics in cultured neurons ...... 291

Figure S6.2 Control showing lack of effect of localized application of acidic solutions

on Ca2+ dynamics in cultured neurons ...... 292

xxiv

Abbreviations

5-HT 5-hydroxytryptamine 5-HT3R 5-HT3 receptor ACh acetylcholine ADO adenosine ADP adenosine 5′-diphosphate ADPR adenosine 5′-diphosphate ribose α,βMeATP α,β-methylene-ATP AMP adenosine 5′-monophosphate ATP adenosine 5′-triphosphate β-NAD+ beta-nicotinamide adenine dinucleotide BoNT/A botulinum neurotoxin A CNS central nervous system CM circular muscle CO carbon monoxide ω-Ctx GVIA ω-conotoxin GVIA cADPR cyclic ADPR cAMP cyclic AMP cGDPR cyclic guanosine diphosphate-ribose DA dopamine DMPP dimethylphenylpiperazinium eADO 1,N6-etheno-ADO eADP 1,N6-etheno-ADP eADPR 1,N6-etheno-ADPR eAMP 1,N6-etheno-AMP eATP 1,N6-etheno-ATP eNAD 1,N6-etheno-NAD EFS electrical field stimulation EHNA erythro-9-(2-hydroxy-3-nonyl)-adenine E-NPP ecto-nucleotide pyrophosphatases/phosphodiesterase E-NTPDase ecto-nucleotide triphosphate diphosphohydrolase ENS enteric nervous system EPSP excitatory postsynaptic potential fEPSP fast excitatory postsynaptic potential fIJP fast inhibitory junction potential GI gastrointestinal HEPES N-2-hydroxyethyl piperazine-N’-ethanesulphonic acid HPLC high-pressure liquid chromatography HPLC-FLD high-pressure liquid chromatography with fluorescence detection H2S hydrogen sulfide ICC interstitial cells of Cajal IJP inhibitory junction potential IPSP inhibitory postsynaptic potential KBH Krebs-bicarbonate-HEPES xxv

LDCV large dense-core-like vesicle LM longitudinal muscle L-NNA NG-nitro-L-arginine MG myenteric ganglia MRS2179 2’-deoxy-N6-methyladenosine 3’,5’-bisphosphate tetrasodium salt MRS2500 (1R,2S,4S,5S)-4-[2-Iodo-6-(methylamino)-9H-purin-9-yl]- 2- (phosphonooxy)bicyclo[3.1.0]hexane-1-methanol dihydrogen phosphate ester tetraammonium salt NA/NE noradrenaline/ NANC non-adrenergic non-cholinergic nAChR nicotinic acetylcholine receptor NGD nicotinamide guanine dinucleotide NGF nerve growth factor NMN nicotinamide mononucleotide NO NPP nucleotide pyrophosphatase PACAP pituitary adenylate cyclase-activating peptide PDGFRα platelet-derived growth factor receptor α PPADS pyridoxal-phosphate-6-azophenyl-2’,4’-disulfonate sIJP slow inhibitory junction potential SK channels small-conductance Ca2 + -activated K+ channels SLMV synaptic-like microvesicle SNAP-25 25-kDa synaptosomal associated protein SNARE soluble N-ethylmaleimide-sensitive factor attachment protein receptor SSV small synaptic-like vesicle TTX tetrodotoxin Tris tris hydroxymethyl aminomethane VDCC voltage-dependent Ca2 + channels WM whole muscle VIP vasoactive inhibitory peptide 1

Chapter 1

Introduction

2

1.1 Chemical synaptic transmission and criteria to define a neurotransmitter

Chemical signaling constitutes a major mechanism in neuroeffector transmission in the central and peripheral nervous systems. Chemical synaptic transmission occurs when an influx of calcium ions (Ca2+) during a presynaptic nerve impulse triggers exocytosis of neurotransmitter substances from synaptic vesicles. Released neurotransmitters diffuse across the synaptic cleft and occupy receptors localized on the membrane of postsynaptic effector cells. Either directly or via activation of second messengers, this interaction activates ion channels and, depending on the conductance activated, the physiological response is an excitatory or inhibitory postsynaptic potential

(EPSP or IPSP, respectively) (Pitman, 1984). We are now well beyond the simplistic notion that a synthesized and released a single neurotransmitter and with the realization that many pharmacologically active signaling molecules can exist in nerves

(Burnstock, 1976; Sneddon & Westfall, 1984; Lundberg, 1996) it has become progressively more difficult to identify which substances can be considered genuine neurotransmitters. To aid in the identification of novel neurotransmitters and to discriminate between neuromodulators, which control the efficacy of synaptic transmission, and other signaling molecules, several criteria were established: 1) the substance must be synthesized in the neuron and stored within the synaptic vesicles; 2) release must occur by Ca2+-dependent vesicle exocytosis and is proportionate to the level of nerve stimulation; 3) exogenous application of the candidate neurotransmitter must mimic the endogenous response; 4) specific receptors for the candidate are present on the 3 post-synaptic target cell; and 5) a mechanism to terminate the extracellular action of the substance must exist (reviewed in Burnstock, 2012).

These ‘classical’ criteria have been broadened with the discoveries that membrane permeable substances such as nitric oxide (NO), carbon monoxide (CO) and hydrogen sulfide (H2S) can function as neurotransmitters without strictly conforming to established criteria (Baranano et al., 2001). The mode of synthesis, storage and release of these transmitters differs markedly from those of other neurotransmitters in that they are not stored in synaptic vesicles, thus not released by exocytosis, and instead synthesized when required. Additionally, instead of binding to receptors on postjunctional membranes, membrane permeable transmitters simply diffuse into target cells immediately affecting excitability by interacting with their intracellular targets. NO, for example, diffuses into the postsynaptic cell and directly activates the intracellular second messenger soluble guanylyl cyclase which facilitates activation of signaling pathways leading to smooth muscle relaxation (Mustafa et al., 2009). Thus classical criteria for neurotransmitter status are continually developed as we learn more about intricacies of neurotransmission.

Still, the non-membrane permeable neurotransmitters are expected to be stored in and released from synaptic vesicles and to act on receptor proteins in the cell membrane of target cells.

1.2 Purinergic non-adrenergic non-cholinergic (NANC) neurotransmission in smooth muscle

Smooth muscle organs, such as urinary bladder, airways, uterus, blood vessels and gastrointestinal (GI) tract, are multicellular organs with an intricate organization 4 consisting of myocytes, nerves, glia, interstitial cells and epithelial cells.

Neurotransmitters have a critical influence on the contractile state of smooth muscles and coordinate muscle contractions and relaxations that underlie the normal motility in these organs. Chemical synaptic transmission in smooth muscle organs is complex and involves multiple molecules with potent excitatory or inhibitory actions in the neuroeffector junction. In the early 1960s it became clear that communication between nerves and smooth muscle is mediated by neurotransmitters in addition to the classical transmitters, acetylcholine (ACh) and norepinephrine (NE), which gave rise to the concept of non-adrenergic non-cholinergic (NANC) neurotransmission (Bennett et al.,

1966; Burnstock et al., 1966). This was first demonstrated in guinea-pig taenia coli where vagal nerve stimulation in the presence of cholinergic and adrenergic blockers elicited a tetrodotoxin (TTX)-sensitive membrane hyperpolarization and smooth muscle relaxation

(Burnstock et al., 1963; Bennett et al., 1966) (Fig 1.1A). The hyperpolarizations were identified as inhibitory junction potentials (IJPs) in response to NANC neurotransmission. By the end of the 1960s NANC nerves in the respiratory, cardiovascular, urinogenital and gastrointestinal tract had been described (Burnstock,

1969).

Efforts directed at identifying the NANC neurotransmitter ensued and in 1970 it was proposed that adenosine 5’-triphosphate (ATP) was released by neurons and appeared to satisfy criteria needed to establish it as a NANC neurotransmitter in the gut

(Burnstock et al., 1970; Bennett, 2013). This proposal, however, was indirectly derived from evidence demonstrating efflux of adenosine (ADO) and inosine, assumed to originate from ATP, during vagal nerve stimulation in guinea pig stomach (Fig 1.1B) 5

(Burnstock et al., 1970). However, we now know that ATP is not the sole source of extracellular adenosine: for example, ADO can be derived from any chemical substance that can produce AMP, including nicotinamide adenine dinucleotide (NAD+), adenosine

5’-diphosphate ribose (ADP-ribose, ADPR) and cyclic ADPR (discussed below)

(Zimmermann et al., 2012). Moreover, recent works have elegantly demonstrated direct neuronal release of ADO (Wall & Dale, 2007; Lovatt et al., 2012). In the above referenced initials studies in the guinea-pig taenia coli the presence of ADO and inosine also provided indirect evidence for the existence of extracellular nucleotidases which would provide an inactivation mechanism for released ATP. Finally, similar postjunctional effects to nerve stimulation and exogenous ATP on muscle contractility in guinea pig taenia coli indirectly suggested that ATP might be involved in NANC neurotransmission (Burnstock et al., 1970) (Fig 1.1C). With the evidence described emerged the concept of ‘purinergic neurotransmission’ (Burnstock, 1972).

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Figure 1.1. (A) Sucrose gap recording of membrane potential changes in smooth muscle of guinea-pig taenia coli in the presence of atropine (0.3 μM) and guanethidine (4 μM). Transmural field stimulation (0.5 ms, 0.033 Hz, 8 V) evoked transient hyperpolarizations, which were followed by rebound . Tetrodotoxin (TTX, 3 μM) added to the superfusing Kreb’s solution (applied at arrow) rapidly abolished the response to transmural field stimulation establishing these as inhibitory junction potentials in response to NANC neurotransmission. Image adapted from Burnstock, 1986. (B) Chromatographic evidence for purine release from stomach stimulated via the vagosympathetic nerve. The figure shows an ultraviolet photoprint of a chromatogram spotted with (from the left) AMP, ATP, perfusate from unstimulated stomach, perfusate from stimulated stomach, adenosine. Vagosympathetic nerve trunks were stimulated for 45 s in every 2 min for a total of 30 min with pulses of 2 ms duration, at 30 Hz; the voltage was progressively increased from 5 to 40 V. Note that stimulation increases the amounts of adenosine and inosine released from the stomach. Image from Burnstock et al., 1970. (C) Mechanical responses of the guinea pig taenia coli to intramural nerve stimulation (NS: 1 Hz, 0.5-ms pulse duration, for 10 s at supramaximal voltage) and ATP (2 x 10-6 M). The responses consist of a relaxation followed by a “rebound contraction.” Atropine (1.5 x 10-7 M) and guanethidine (5 x 10- 6 M) were present. Image adapted from Burnstock & Wong, 1978.

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The proposal of purine-mediated NANC neurotransmission was received with skepticism stemming from the fact that ATP has well-established intracellular signaling roles and it seemed unlikely that such a ubiquitous molecule would also function extracellularly. However hints in the literature for extracellular roles of purines originally surfaced more than 40 years earlier when powerful extracellular inhibitory actions of

ADO on heart, blood vessels and intestine were demonstrated (Drury & Szent-Gyorgyi,

1929). Then in 1959 Pamela Holton provided the first evidence using the relatively selective and sensitive firefly chemiluminescence assay that ATP was released during stimulation of sensory nerves supplying the rabbit ear artery (Holton, 1959). Although commonly overlooked, this was the first study demonstrating a possible neural source of

ATP as suggested by significantly reduced ATP release after mechanical denervation.

However whether ATP was released as a neurotransmitter or rather as a result of the release of other neurotransmitters during nerve stimulation deserves consideration. Some investigators point out that ATP is important for acidifying vesicle lumens, for creating a proton gradient for vesicular neurotransmitter uptake and for steps in exocytosis

(Sperlagh & Vizi, 1996) thus release of ATP from vesicles is not conclusive evidence for its role as a neurotransmitter.

Although the physiological relevance of extracellular purines has grown and is now very well appreciated, with roles extending beyond those of neurotransmitters and neuromodulators to paracrine and autocrine mediators released from non-excitable cells

(Schenk et al., 2008; D'hondt et al., 2011; Junger, 2011), considerable controversy surrounds the identity of the purine neurotransmitter in some smooth muscle organs. For example, in the GI tract of several species ATP does not appear to mediate NANC 8 neurotransmission (Serio et al., 1990; Serio et al., 1992; Serio et al., 1996; Ohno et al.,

1996; Ivancheva et al., 2000; Mutafova-Yambolieva et al., 2007; Hwang et al., 2011;

Hwang et al., 2012; Durnin et al., 2013). In recent years work in Mutafova-Yambolieva’s laboratory has shown that other purines, such as NAD+ and its metabolite ADPR, may have important extracellular roles in the central and peripheral nervous systems, and in the enteric nervous system NAD+ and/or ADPR better fulfill criteria as neurotransmitter(s) than ATP (Mutafova-Yambolieva et al., 2007; Hwang et al., 2011;

Durnin et al., 2012b; Durnin et al., 2013). These studies also demonstrated that during nerve stimulation the presence of extracellular ADO in the colon originates primarily from metabolism of NAD+, rather than ATP, casting doubt on the claims for ATP release due to the presence of ADO (Burnstock et al., 1970). Knowing the true identity of the purinergic neurotransmitter is important for understanding critical mechanisms of cell-to- cell communication and may also have important translational aspects. Thus, identifying the purine neurotransmitter in smooth muscles could lead to identifying selective and specific targets including mechanisms of release, metabolism and/or action of the transmitter for therapeutically treating conditions associated with altered purinergic signaling. Thus the journey to identifying the purinergic neurotransmitter continues.

1.3 Difficulties associated with the study of purinergic neurotransmission in smooth muscles

The contribution of purines to the regulation of smooth muscle contractility is a notoriously difficult area to study attributable to a number of factors, outlined below.

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1.3.1 Variable purine release mechanisms

Purine nucleotides such as ATP and NAD+ are ubiquitous intracellular constituents that can be released from essentially every cell type via numerous mechanisms. ATP, for instance, can be released through and pannexin hemichannels, purinergic P2X7 receptor pores, maxi ion channels, volume-regulated ion channels and via vesicle exocytosis (Arcuino et al., 2002; Stout et al., 2002; Darby et al.,

2003; Suadicani et al., 2006; Pankratov et al., 2006; Lazarowski, 2012; Gulbransen et al.,

2012). In fact, ATP is sometimes used as a universal tracer of all cellular secretion events

+ (Aspinwall & Yeung, 2005). NAD can also pass through P2X7 receptor pores and connexin hemichannels (Bruzzone et al., 2001; Lu et al., 2007). In recent years continuous work in Mutafova-Yambolieva’s laboratory demonstrated regulated release of

NAD+ including release via vesicle exocytosis in blood vessels, urinary bladder and GI muscles (Smyth et al., 2004; Breen et al., 2006; Smyth et al., 2006b; Mutafova-

Yambolieva et al., 2007; Hwang et al., 2011), but it is generally recognized that identifying the purine(s) that are released as neurotransmitter(s) in various smooth muscle organs is challenging (Mutafova-Yambolieva, 2012).

1.3.2 Complex metabolic pathways of extracellular purines

Following their release into the extracellular compartment, purines are quickly degraded to metabolites that might be bioactive in the neuroeffector junction. As depicted in Fig 1.2 ATP is degraded sequentially to adenosine 5’-diphosphate (ADP), adenosine

5’-monophosphate (AMP) and adenosine (ADO) by ectonucleoside 5’-triphosphate diphosphohydrolases (E-NTPDases, CD39 family), nucleotide pyrophosphatases (NPPs) 10 and 5’-nucleotidase (CD73), respectively (Zimmermann, 2000). NAD+ is degraded to

ADP-ribose (ADPR) and cyclic ADPR (cADPR) by NAD glycohydrolase and ADP- ribosyl cyclase, respectively, and cADPR is hydrolyzed to ADPR by ADP-ribosyl hydrolase (Zocchi et al., 1993; Lee, 2001; De Flora et al., 2004). These enzyme activities are mainly associated with type II integral membrane glycoprotein, CD38 (Graeff et al.,

1998; Lee, 2001), although other proteins such as CD157 (Ortolan et al., 2002) or CD73

(Garavaglia et al., 2012) may also play roles in NAD+ metabolism. After its formation from NAD+, ADPR is further degraded to AMP and ADO by NPPs and 5’-nucleotidase, respectively (Zimmermann, 2000). In addition NPPs can produce nicotinamide mononucleotide (NMN) and AMP from NAD+ which are further degraded to ADO

(Zimmermann, 2000). While the presence of ADO in the neuroeffector junction originates primarily from ATP or NAD+ metabolism, there may also be some populations of neurons that release ADO directly (Wall & Dale, 2007; Lovatt et al., 2012). Many purines can therefore exist in the neuroeffector junction with potential roles in regulating smooth muscle contractility.

Figure 1.2. Biotransformation pathways for extracellular purines. Image from Peri et al., 2013. 11

1.3.3 Abundant purine receptor expression

There is widespread expression of purine receptors on multiple cell types within smooth muscles. In fact, purine receptors may be the most abundant of all mammalian receptors (Burnstock & Knight, 2004). Their discovery in 1978 was highly significant, reinforcing the ‘purinergic hypothesis’ (Burnstock, 1978). Purine receptors are divided into P1 and P2 receptor families. P1 receptors are G-protein coupled receptors which mediate responses to ADO and are comprised of A1, A2A, A2B and A3 receptor subtypes

(Abbracchio et al., 2009; Burnstock, 2012). Typically A1 and A3 couple to the Gi/o family of G proteins inhibiting intracellular cyclic AMP (cAMP) production whereas A2A and

A2B stimulate cAMP production via Gs. P2 receptors are categorized as ionotropic P2X or metabotropic P2Y receptor families. Seven P2X (P2X1-7) and eight P2Y (P2Y1,2,4,6,11-14) receptors are expressed in the mammalian nervous system (Abbracchio et al., 2009) and mediate responses to various purines. P2X receptors are classical ligand-gated cation channels permeable to Na+, K+ and Ca2+ ions upon activation by purines (North, 2002); in the central nervous system these receptors modulate fast neurotransmission by regulating the release of neurotransmitters. G-protein coupled P2Y receptors are activated by adenine, pyridine and pyrimidine nucleotides (Harden et al., 2010). P2Y1, 2, 4, 6 and 11 primarily use Gq/G11 to activate the phospholipase C/inositol triphosphate intracellular

2+ Ca -release pathway (Verkhratsky, 2005) whereas P2Y12, 13 and 14 receptors almost exclusively couple to Gi/o which inhibits adenylyl cyclase (Abbracchio et al., 2006). In general smooth muscle contractile responses to purines are mediated primarily by P2X receptors whereas relaxation responses are commonly mediated by P2Y receptors. 12

However with the diversity of receptor subtypes, expression of multiple subtypes by many cells, and the formation of homo- and hetero-multimers (Burnstock, 2008b;

Abbracchio et al., 2009), it becomes increasingly challenging to identify the purinergic mechanisms regulating contractility in different smooth muscles.

1.3.4 Inadequate methodologies for purine detection

Examination of purines released during nerve stimulation is difficult using conventional detection methods due to rapid degradation and low final concentrations that could remain below detection thresholds (Bobalova et al., 2002). Spectrophotometry, fluorescence, chemiluminescence and bioluminescence assays and electrochemical biosensors have been used for the detection of ATP in various systems, however these methodologies are limited to the detection of a single purine nucleotide, do not take into consideration the production of metabolites, and thus are not sufficient for identifying which purine(s) are released during nerve stimulation. Furthermore, the lowest detection limits for these methods are relatively high, ranging from detection of low micromolar concentrations of ATP using fluorescence assays (Wang et al., 2005) to high nanomolar concentrations using chemiluminescence assays (Wang et al., 2005; Zhang et al., 2009) and biosensors (Kueng et al., 2004). As a result, most of the literature on purinergic neuromuscular transmission is based on studies examining postsynaptic responses to purines without evaluating presynaptic release mechanisms. This indirect evidence cannot determine the exact nature of the purine that is released from the presynaptic vesicle causing some investigators to question the identity of the purine transmitter in some smooth muscles (Serio et al., 2003). 13

Together these factors demonstrate the difficulties associated with understanding complex purinergic mechanisms regulating smooth muscle function which makes identification of novel purine mediators challenging.

1.4 Evidence for NAD+ as a NANC neurotransmitter in peripheral smooth muscles

1.4.1 High-pressure liquid chromatography (HPLC) techniques for sensitive detection of purines

Perhaps the most effective method for purine detection is the reversed-phase gradient high-pressure liquid chromatography assay with fluorescence detection (HPLC-

FLD), which allows highly sensitive detection of multiple purines that might be released during nerve stimulation in smooth muscle preparations (Mutafova-Yambolieva et al.,

1997). To increase detection sensitivity, chloroacetaldehyde added to samples forms fluorescent 1,N6-etheno-purine analogs, effectively reducing the detection limits to low femtomolar concentrations of purines released (Levitt et al., 1984; Bobalova et al., 2002).

HPLC detection of purines is a technique central to our laboratory that gives us the major advantage over conventional detection methods of simultaneously measuring the release of principle purines (i.e. ATP and NAD+) plus their metabolites (ADP, AMP, ADO, cADPR and ADPR*; see p14) in a single biological sample (see Fig 1.3). Thus direct comparisons in the mechanisms of release of different purines can be accurately achieved. Of special importance, using this methodology our laboratory has made several significant contributions to our understanding of purinergic regulation of smooth muscles including novel extracellular signaling roles of NAD+ in the peripheral nervous system. 14

*Note (Fig 1.3): NAD+ and cADPR are unstable during the high temperatures of the etheno- derivatization process and form ADPR; therefore NAD+, ADPR and cADPR elute as a single peak in HPLC chromatograms. HPLC fraction analysis of the NAD++ADPR+cADPR peak was developed in our laboratory and is routinely carried out to determine the contribution of each nucleotide to the mixture in different smooth muscle preparations, which varies. For example in human urinary bladder NAD+ represents ~90-95% of the mixture, ADPR ~5-10% and cADPR is absent (Breen et al., 2006) whereas in canine mesenteric vessels NAD+ represents ~60%, ADPR ~30% and cADPR ~10% (Smyth et al., 2004). In primate colons NAD+ contributes ~64-82%, ADPR 18-33% and cADPR 0.6-3% of the NAD++ADPR+cADPR mixture (Hwang et al., 2011); novel extracellular functions of ADPR in the gut are discussed below.

1.4.2 Extracellular actions of NAD+

NAD+, like ATP, is an important intracellular regulator of energy transfer, serving as a coenzyme for cellular oxidation-reduction reactions, a donor of ADPR in the posttranslational modification of proteins, and a precursor to cADPR and other intracellular second messengers with Ca2+-releasing activity (Lee, 2001; Berger et al.,

2004). Besides these key roles in energy metabolism, extracellular functions of NAD+ have also emerged (Ziegler & Niere, 2004; Billington et al., 2006). For example, extracellular NAD+ induces Ca2+ signaling and apoptosis in human osteoclastic cells

(Romanello et al., 2001), activates purinergic P2Y11 receptors in human granulocytes leading to increased cAMP production (Moreschi et al., 2006), and together with ADPR, activates Ca2+ influx in human monocytes (Gerth et al., 2004).

1.4.3 Emergence of NAD+ as a putative neurotransmitter or neuromodulator in peripheral smooth muscles

In the last decade evidence has been accumulating that NAD+ is released during nerve stimulation in vascular and visceral smooth muscles and has a putative neurotransmitter role, extending the complexity in understanding purinergic mechanisms 15 regulating smooth muscle function (Smyth et al., 2004; Breen et al., 2006; Smyth et al.,

2006b; Mutafova-Yambolieva et al., 2007; Hwang et al., 2011). In 2004, it was elegantly demonstrated using HPLC methodologies that electrical field stimulation (EFS) of canine mesenteric vasculature evoked release of NAD+ along with ATP and the classical sympathetic neurotransmitter, NE, in a manner consistent for a neurotransmitter (Smyth et al., 2004). Thus, EFS evoked release of NAD+ that was dependent on the level of neural activity and was significantly attenuated by the neural inhibitors, TTX and ω- conotoxin GVIA, and by cleavage of the soluble N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE), SNAP-25, with botulinum neurotoxin A

(BoNT/A) (Smyth et al., 2004; Smyth et al., 2006b). In addition, sympathetic denervation in mesenteric vessels abolished NAD+ release (Smyth et al., 2006b) establishing a neural origin of NAD+ in vascular smooth muscle.

Not limited to the vasculature, neural release of NAD+ has been identified in numerous smooth muscles from many species including rat and mouse tail artery, canine, rabbit, guinea pig and mouse urinary bladder, rat , canine and guinea pig mesenteric vasculature (Smyth et al., 2004), human urinary bladder (Breen et al., 2006), and human, monkey and mouse GI tract (Mutafova-Yambolieva et al., 2007; Hwang et al., 2011), suggesting NAD+ might have important extracellular roles in multiple smooth muscle organs. The classical sympathetic neurotransmitters, NE and ATP, and the neuropeptides, calcitonin gene-related peptide, vasoactive intestinal peptide (VIP), substance P and neuropeptide Y did not evoke the release of NAD+, nor did contraction of the smooth muscle per se (Smyth et al., 2006b), suggesting an independent mechanism of release during nerve stimulation. Interestingly, the release of ATP in these same 16 experiments could not be blocked with the neural N-type Ca2+ channel blocker, ω- conotoxin GVIA (Smyth et al., 2009) (Fig 1.3), suggesting mechanisms other than Ca2+- dependent vesicle exocytosis may be involved in its release. As a result, it is fundamental that we shift our thinking beyond that of ATP and consider other purine nucleotides that may also be involved in purine-mediated neurotransmission to smooth muscle organs.

While these studies did not aim to determine whether NAD+ mimics the postjunctional effects of the endogenous neurotransmitter in smooth muscles, they did demonstrate physiological activity of extracellular NAD+ on smooth muscle tone. For example, exogenously applied NAD+ was reported to relax blood vessels (Smyth et al.,

2009) and reduce the frequency and amplitude of spontaneous contractions in urinary bladder (Breen et al., 2006; Smyth et al., 2006b).

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Figure 1.3. HPLC-fluorescence detection analysis of purine release upon EFS (16 Hz, 0.3 ms, 120 s) in canine isolated mesenteric artery (left column) and vein (right column). Original chromatograms are shown from tissue superfusate samples collected before (pre- stimulation, PS, A and B) and during EFS (stimulation, ST) in the absence (control – C, D) and presence of ω-CTX GVIA (5 nM – E, F). Image adapted from Smyth et al., 2009.

For neurotransmission to be effective, neurotransmitters must be degraded and removed after their release into the neuroeffector junction. A mechanism for terminating the extracellular action of NAD+ must therefore exist in nerve-smooth muscle preparations to qualify as a neurotransmitter substance. In blood vessels, a functional role of CD38 in the metabolism of neurally released NAD+ was suggested by: 1) the presence of NAD+ metabolites, ADPR, cADPR, AMP and ADO, in overflow samples collected during nerve stimulation (Smyth et al., 2004; Smyth et al., 2006b); 2) the degradation of 18 exogenous NAD+ after contacting smooth muscles; and 3) the presence of CD38 exclusively at nerve terminals in canine vasculature, demonstrated by co-localization with the vesicle proteins and SV2 (Smyth et al., 2006a). Importantly, the catalytic site of CD38 faces the extracellular space (Lee et al., 1993; Munshi et al., 2000) thus providing an effective mechanism for degradation and removal of extracellular

NAD+ released upon nerve stimulation.

Although these earlier studies demonstrated unequivocally that NAD+ can be released upon nerve stimulation by Ca2+-dependent exocytosis, the quest for additional evidence for a neurotransmitter role of NAD+ and other purines continues.

1.5 A single-cell model to characterize storage and release of NAD+

An important aspect in the identification of new neurotransmitter candidates is verification that the substance is stored in synaptic vesicles. NAD+ is a ubiquitous intracelluar molecule therefore its presence in nerve terminals cannot be disputed.

Furthermore, release of NAD+ from smooth muscle by mechanisms requiring intact exocytotic machinery is suggestive of storage in vesicles (Smyth et al., 2004; Breen et al., 2006; Smyth et al., 2006b). Nonetheless, as described in Section 1.3, there are many difficulties associated with the study of neurotransmission in multi-cellular smooth muscle organs and only somewhat indirect information about co-storage and co-release of NAD+ with other neurotransmitters is possible.

To overcome some of the challenges associated with smooth muscles, single cell models are often used to verify storage of molecules in vesicles and to aid in the 19 identification of putative new neurotransmitter substances. Nerve growth factor (NGF)- differentiated rat pheochromocytoma PC12 cells are one such model that phenotypically resemble sympathetic neurons. They contain large dense core granules and small synaptic vesicles and synthesize and store the sympathetic catecholamine neurotransmitters (NE and dopamine) (Greene & Tischler, 1976) and ATP (Wagner, 1985). We reported that

NAD+ is also present in vesicles in PC12 cells and is released upon membrane along with dopamine and ATP (Yamboliev et al., 2009; Chapter 2). This is an important piece of information as it validates that NAD+ can be stored in and released from vesicles.

It should be highlighted again, however, that the presence of purines in vesicles is not conclusive evidence for their roles as neurotransmitters; for example, as mentioned earlier, ATP might be present in vesicles for regulating steps in the synthesis, uptake and release of other neurotransmitters (Sperlagh & Vizi, 1996). Thus the mechanisms mediating release of purines from vesicles should also be considered. Interestingly, in

PC12 cells the release of ATP, NAD+ and dopamine is differentially regulated, NAD+ and dopamine, but not ATP, requiring intact SNAP-25-mediated vesicle exocytosis

(Yamboliev et al., 2009). Thus, similar to what was reported in complex vascular smooth muscles (Smyth et al., 2009), there appears to be different preferential mechanisms or sites of release of ATP and NAD+ upon membrane depolarization in PC12 cells. Parallel release with dopamine may suggest a role of NAD+, rather than ATP, as a co-transmitter substance.

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1.6 Purinergic neuromuscular transmission in the gastrointestinal tract

Highly sensitive HPLC assays in conjunction with functional studies in blood vessels and urinary bladder have identified novel mechanisms in purinergic regulation of smooth muscle function. Specifically, a novel role of extracellular NAD+ as a neuromodulator or neurotransmitter in smooth muscle has been suggested. In some systems it is conceivable that ATP is not the purine responsible for NANC neurotransmission due to its release via non-exocytotic mechanisms (e.g. Smyth et al.,

2009; Yamboliev et al., 2009). While the initial discoveries of purine-mediated NANC neurotransmission were made in gastrointestinal smooth muscles (Burnstock et al.,

1963), presynaptic release mechanisms of purines (i.e. ATP) were largely overlooked in these studies. An important question therefore arises: taking into account the mechanisms of release, metabolism and action of extracellular purines, does ATP truly qualify as the purinergic neurotransmitter in the enteric nervous system? This question has been addressed in detail in Chapters 3, 4 and 5 of this dissertation and the major findings are summarized below (see Section 1.7).

1.6.1 Brief overview of the enteric nervous system

The GI tract is composed of anatomically and functionally distinct regions, beginning at the esophagus, proceeding to the stomach, small intestine, large intestine and ending at the rectum. GI smooth muscles are multifaceted tissues composed of many cell types including myocytes, nerve cells, glial cells and several types of interstitial cells.

Neural control of enteric smooth muscle is considerably complex, constituting an 21 extrinsic system, the parasympathetic and sympathetic nervous systems, and an intrinsic system, the enteric nervous system (ENS).

The ENS controls GI function independent of direct connections with the central nervous system (CNS) by neurons located in the myenteric and submucosal nerve plexuses that regulate GI motility and secretion, respectively. The tunica muscularis of the GI tract is composed of an inner circular and an outer longitudinal muscle layer innervated by excitatory and inhibitory motor neurons of the myenteric plexus between the muscle layers (Kunze & Furness, 1999; Bornstein et al., 2004) (Fig 1.4). These motor neurons release multiple neurotransmitter substances that coordinate circular muscle contractions that underlie effective propulsion of luminal contents (Bertrand, 2003).

Enteric excitatory neuromuscular transmission is mediated predominantly by ACh and neurokinins such as substance P and neurokinin A (Brookes et al., 1992; Lomax &

Furness, 2000). Inhibitory neurotransmission primarily involves nitric oxide (NO) and purines (Spencer & Smith, 2001; Gil et al., 2010), although the relative roles of each appear to vary considerably between gut regions and between species. For example in the rat, NO-mediated neurotransmission appears to be more prominent in proximal colon than distal colon (Suthamnatpong et al., 1993; Takahashi & Owyang, 1998). In mice, purinergic signaling may be more prominent in the distal colon, suggested by delayed pellet transit in the distal region of a mouse with disrupted purinergic motor transmission

(Hwang et al., 2012). In human colon, inhibitory neurotransmission does not appear to have a prominent nitrergic component (Gallego et al., 2008a; Hwang et al., 2011; see

Chapter 3). VIP and pituitary adenylate cyclase-activating peptide (PACAP) may also play roles in enteric inhibition (Lomax & Furness, 2000). After their release from nerve 22 terminals, motor neurotransmitters influence gut motility either by directly targeting receptors on the smooth muscle or by targeting receptors on different classes of interstitial cells that transduce signals to the smooth muscle via gap junctions (discussed below).

Upstream of motor neurons, orally-directed ascending and aborally- directed descending interneurons form sophisticated synaptic networks in the myenteric plexus and release multiple transmitter substances that influence the activity of interneurons and muscle motor neurons (Bornstein et al., 2004). In addition, intrinsic sensory neurons (intrinsic primary afferent neurons, IPANS) are activated in response to mechanical and chemical stimuli and transduce information from the gut lumen to influence activity of enteric interneurons and motor neurons (Furness et al., 1998). Thus the ENS regulates intestinal motility by the coordinated activity of intrinsic sensory neurons, several types of interneurons and enteric motor neurons (depicted in Fig 1.5 from Rodriguez-Tapia & Galligan, 2011). Understanding the precise signaling events leading to contraction or relaxation in GI smooth muscles is appreciably complex.

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Myenteric plexus Circular muscle

Deep muscular plexus Inner SMP Outer SMP

Longitudinal muscle

Mucosa Muscularis mucosae Submucosal artery

Figure 1.4. The organization of the ENS of human and medium-large mammals. The ENS has ganglionated plexuses, the myenteric plexus between the longitudinal and circular layers of the external musculature and the SMP that has outer and inner components. Nerve fiber bundles connect the ganglia and also form plexuses that innervate the longitudinal muscle, circular muscle, muscularis mucosae, intrinsic arteries and the mucosa. Innervation of gastroenteropancreatic endocrine cells and gut-associated lymphoid tissue is also present, which is not illustrated here. Abbreviations: ENS, enteric nervous system; SMP, submucosal plexus. Image from Furness, 2012.

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Figure 1.5. Simplified neural circuits responsible for the peristaltic reflex. A bolus of gastrointestinal content stretches the gut wall resulting in contraction above and relaxation below the point of stimulation. (left panel) IPANs are found in the submucosal and myenteric plexuses. Mechanical or chemical stimulation of the mucosa causes enterochromaffin cells to release serotonin (5-HT). 5-HT acts to excite the mucosal terminals causing action potentials to propagate retrogradely back to the cell body. IPANs connect with orally and anally directed interneurons which connect with excitatory and inhibitory motoneurons. (right panel) There are also multifunctional neurons in the myenteric plexus which respond to mechanical stimulation. These multifunctional neurons connect with interneurons to initiate polarized contractions and relaxations of gut smooth muscle. Image from Rodriguez-Tapia & Galligan, 2011.

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1.6.2 Purines in the enteric nervous system

Purinergic signaling is essential to normal GI function; in particular, purines are involved in coordinating contractions and relaxations in GI smooth muscle therefore are important mediators of propulsive motility. However a comprehensive understanding of purinergic mechanisms involved in the physiological control of digestive functions is exceptionally complex. In the ENS purines can act as inhibitory neurotransmitters from motor neurons to the circular smooth muscle (via P2Y receptors), excitatory neurotransmitters between enteric interneurons and from interneurons to motor neurons

(via P2X and P2Y receptors) and sensory mediators from epithelial sources to intrinsic sensory nerve terminals (Bertrand, 2003). Thus purines participate in essentially all aspects of intestinal function, from the sensory transduction of stimuli from the gut lumen to the subsequent initiation and propagation of enteric reflexes. In addition, purines are involved in inflammatory processes in response to tissue damage (see below).

There is widespread distribution of purine receptors throughout the gut that are involved in regulating intestinal motility both by modulating enteric neurotransmitter release and by directly controlling smooth muscle contractility (Antonioli et al., 2013).

Additionally, the activity of enzymes and transporters modulates the magnitude and duration of purinergic signaling by controlling levels of bioactive nucleotides and nucleosides in the extracellular compartment (Antonioli et al., 2013). The term ‘enteric purinome’ has recently been used to holistically describe purine nucleotides, catabolic enzymes, transporters and receptors involved in triggering, maintaining and terminating purinergic signaling in the gut (Antonioli et al., 2011); much efforts are currently channeled at gaining more detailed information on the involvement of the ‘enteric 26 purinome’ in regulating intestinal motility. Three major mechanisms underlie purine release in the GI tract: 1) regulated release from neuronal sources (i.e. neurotransmitters);

2) regulated release from non-neuronal sources; and 3) non-regulated release from damaged or inflamed tissues.

1.6.2.1 Regulated release from neuronal sources

Purines released during stimulation of myenteric motor nerve terminals mediate relaxation of circular smooth muscle which is necessary for functions such as sphincter relaxation, gastric accommodation and the peristaltic reflex (Bennett, 1997). In addition spontaneous purine release from motor neurons mediates constant tonic inhibition of the circular muscle at rest (Dickson et al., 2010b) which keeps the muscle in a hyperpolarized state for normal transit and stool formation (Dickson et al., 2010a). Tonic inhibition may be an extremely important feature of colonic behavior and serve to suppress uncoordinated contractile activity or development of tone between propulsive contractions (Koh et al., 2012). In NANC conditions, nerve stimulation elicits biphasic

IJPs causing relaxation of gut smooth muscles. The IJPs consist of a fast hyperpolarization (fIJP) followed by a slower hyperpolarization (sIJP) (Shuttleworth et al., 1997; Serio et al., 2003). The slow, longer-lasting hyperpolarization is nitrergic while the fast initial hyperpolarization is mediated by purine(s) (Hirst et al., 2004; Vetri et al.,

2004; Gallego et al., 2006). Given the diversity of purine receptor subtypes it is well

2+ + accepted that the P2Y1 receptor, linked to openings of apamin-sensitive Ca activated K channels (SK channels) on the postsynaptic cell, is the predominant mediator of purinergic fIJPs and smooth muscle relaxation (Banks et al., 1979; Hirst et al., 2004; 27

Gallego et al., 2006; Mutafova-Yambolieva et al., 2007; Hwang et al., 2011). Disrupted colonic transit in a mouse model with genetic deactivation of P2Y1 receptors highlights the importance of intact purinergic mechanisms to normal GI function (Hwang et al.,

2012). Currently, it is not clear whether purines are co-released with NO from inhibitory nitrergic motor neurons or independently from ‘purinergic’ motor neurons in the intestine.

Within the circuitry of the ENS, neurally-released purines are involved in initiating enteric reflexes by contributing to the descending inhibitory limb of the peristaltic reflex; thus a portion of synaptic excitation between enteric interneurons and from descending interneurons to inhibitory motor neurons is mediated by purines

(Galligan, 2002). Fast purine-mediated EPSPs are elicited predominantly via activation of

P2X receptors localized on neuronal cell bodies (Galligan & North, 2004), the P2X2 and

P2X3 receptor subtypes appearing to be important particularly in the small intestine (Ren et al., 2003; Bian et al., 2003). Recently a role of P2Y1 metabotropic receptors in interneuronal excitation in the myenteric plexus has been reported (Thornton et al.,

2013). In addition to their involvement in descending inhibition, P2X receptors contribute to interneuronal transmission in ascending and descending excitatory cholinergic pathways in the distal colon (Spencer et al., 2000; Monro et al., 2002). Thus purines released from myenteric interneurons have roles in regulating both the contractions and relaxations of the circular smooth muscle.

Neurally-released purines can also activate P2X and P2Y receptors on enteric glial cells (Gulbransen et al., 2010; Boesmans et al., 2013) which are involved in modulating neuronal signaling through their expression of neurotransmitter receptors, 28 biosynthetic enzymes, reuptake transporters and degradation enzymes (Gulbransen et al.,

2010). Altered motility after disrupting glial cells with the gliotoxin fluorocitrate is suggestive of important roles of glia in regulating intestinal function (Nasser et al., 2006); however the precise roles of purines on glial cell function are currently not clear

(Bornstein, 2008).

1.6.2.2 Regulated release from non-neuronal sources

In response to physiologic stimuli such as distention or under pathological conditions such as during inflammation, regulated release of purines from cells in the gut wall initiates enteric reflexes. For example purines released from epithelial and enterochromaffin cells can activate P2 receptors (putative P2X2, P2X3 or P2Y receptors) on sensory nerve endings which transduce sensory stimuli to neurons in the submucosal and myenteric plexus to initiate gut secretory and motor reflexes, respectively (Bertrand

& Bornstein, 2002; Raybould et al., 2004). In addition, autocrine or paracrine activation of purine receptors on epithelial and enterochromaffin cells stimulates fluid secretion and serotonin (5-HT) release, which also transmits sensory information to enteric neurons

(Cooke et al., 2003). Regulated release of purines from enteric glial cells has also been suggested (Zhang et al., 2003). ATP transporters, exocytosis, hemichannels, receptor activation or ATP-conducting ion channels are possible mechanisms by which ATP (or other purines) may be released from non-neuronal cells (Maroto & Hamill, 2001).

1.6.2.3 Non-regulated release from damaged and inflamed tissues 29

During infection or inflammation large amounts of purines are released into the extracellular compartment by unregulated mechanisms as a result of cell damage, cytolysis or apoptosis (Cooke et al., 2003). This is thought to occur during the early stages of apoptosis in order to recruit monocytes and macrophages to the site of inflammation for efficient apoptotic cell clearance (Elliott et al., 2009). In the myenteric plexus, non-regulated purine release can have critical implications in neuronal survival

(Gulbransen et al., 2012). Elevated levels of extracellular ATP during tissue damage or inflammation chronically activates neuronal P2X7 receptor/pannexin Panx1 channel complex triggering intracellular signaling events that lead to myenteric neuron cell death

(Gulbransen et al., 2012). Thus unregulated purine release may contribute to colonic motility dysfunction in inflammatory GI disorders such as colitis (Roberts et al., 2012).

1.7 Road to identifying the enteric purine motor transmitter

As mentioned, evaluating presynaptic release of purines is challenging with available detection methodologies causing most investigators to make indirect assumptions about the release of ATP in enteric smooth muscles. For example, Burnstock and colleagues demonstrated that exogenous ATP elicited relaxation in guinea pig and human colonic smooth muscles which was similar the response produced by NANC nerve stimulation (Burnstock et al., 1970; Burnstock et al., 1972); thereby ATP was assigned as the inhibitory neurotransmitter. Other indirect evidence included the presence of ADO (assumed to be derived from ATP) in samples collected during nerve stimulation 30

(Burnstock et al., 1970), the existence of ecto-nucleotidases in GI muscles (Burnstock,

2008b; Burnstock, 2009), the fact that ATP is a potent agonist at purinergic P2 receptors

(von Kugelgen, 2006) including P2Y1 receptors (Gallego et al., 2006; Gallego et al.,

2008b) and evidence demonstrating the presence of ATP in synaptic vesicles (Dowdall et al., 1974; Aberer et al., 1978; Volknandt & Zimmermann, 1986).

Worth mentioning, many earlier studies monitored uptake and release of radiolabelled ADO during nerve stimulation as evidence for vesicular storage of ATP in smooth muscles (Su et al., 1971; White, 1988). Additionally quinacrine-binding has been used in early studies to localize ATP: for example quinacrine-labeling of myenteric nerves has been used to demonstrate storage of ATP in enteric neurons (Olson et al.,

1976). However quinacrine is very non-selective and binds other adenine nucleotides, guanylic acid and nucleic acids (Irvin & Irvin, 1954a; Irvin & Irvin, 1954b; Kurnick &

Radcliffe, 1962). Thus there are clearly significant specificity problems with the use of radioactive tracers and quinacrine for ATP detection.

In several GI smooth muscle preparations, ATP does not appear to play any role in producing apamin-sensitive IJPs (e.g. rat duodenum (Serio et al., 1990), guinea pig gastric fundus (Ohno et al., 1996), rat colon (Serio et al., 1992), rat cecum (Serio et al.,

1996) and guinea pig ileum (Ivancheva et al., 2000)). Moreover, upon rigorous examination of both presynaptic release mechanisms and postsynaptic effects, we reported that ATP does not fulfill criteria for a motor neurotransmitter in mouse and primate colon (Mutafova-Yambolieva et al., 2007; Hwang et al., 2011; Durnin et al.,

2013); see Chapters 3, 4 and 5). As a result, many investigators today refer to the 31 substance as a ‘purine’ or ‘purine-like’ (Serio et al., 2003; Gallego et al., 2006). Ongoing debate surrounds the true identity of the purine involved in regulating colonic motility.

1.7.1 Characterization of the enteric purine neurotransmitter

Resolving the identity of the purine mediating colonic inhibition may have important implications therapeutically. It could lead to development of novel selective and specific treatments for GI motility disorders associated with altered purinergic transmission that target steps in biosynthesis, release or degradation of the purine or manipulation of its receptor’s activity. Therefore intensive research has been carried out in our laboratory in an effort to fully characterize the enteric purine motor neurotransmitter(s). With the emergence of extracellular and putative neurotransmitter roles of NAD+ in the peripheral nervous system, a major goal of this dissertation was to determine if NAD+ could also be considered a neurotransmitter in the enteric nervous system and perhaps better fulfill neurotransmitter criteria than ATP. Of note, NAD+ is an agonist at purinergic P2Y1 receptors as demonstrated by receptor-mediated responses in

HEK293 cells expressing P2Y1 receptors (Mutafova-Yambolieva et al., 2007). Equipped with the tools to achieve an integrated view of the mechanisms of release, metabolism and extracellular actions of purines, our innovative work challenges the concept that has for 40 years suggested that purinergic inhibitory control of colonic motility is mediated exclusively by ATP (described in detail in Chapters 3, 4 and 5).

Although we are just beginning to scratch the surface, we have made several significant contributions to our understanding of purinergic motor neurotransmission in colon. In 2007 a seminal paper demonstrated several key differences in release and 32 extracellular action of the purines, ATP and NAD+, in the mouse colon (Mutafova-

Yambolieva et al., 2007). For example, release of NAD+, but not ATP, during nerve stimulation depended on the frequency of stimulation and was sensitive to inhibitors of neural activity, TTX and ω-conotoxin GVIA (Mutafova-Yambolieva et al., 2007). These neural blockers have been shown previously to block purinergic inhibitory neurotransmission in the gut (Banks et al., 1979; Bridgewater et al., 1995; Shuttleworth et al., 1997; Rae et al., 1998; Gil et al., 2010) establishing NAD+ as a candidate purine motor transmitter in colon. Postsynaptically, while exogenous ATP and NAD+ both caused hyperpolarizations in the gut, only the hyperpolarizations to NAD+ were blocked by non-selective P2 receptor antagonists and by the selective P2Y1 receptor antagonist,

MRS2179. The SK channel blocker, apamin, also blocked membrane potential changes to

NAD+. Thus the pharmacological profile of NAD+ mimicked that of the endogenous purine motor transmitter in colon better than ATP and for the first time, NAD+ was considered as an enteric inhibitory neurotransmitter in mouse colon.

Mice are intensively used as an experimental system due to similarity to humans with respect to genome organization, development and physiology. Moreover, mouse models can be used to verify the functional significance of a given gene. However, the motor patterns in the colon of different species can vary significantly due to differences in diet and anatomical arrangements of the colon between species (Christensen, 1985).

Thus it is fundamental that data obtained in small mammals such as mice can be translated to higher species in order to be functionally relevant to mechanisms of colonic motility in humans. Described in detail in Chapter 3, and in accordance with findings mice, nerve stimulation of colonic preparations from human and non-human primates 33 also evokes release of NAD+ in a frequency-dependent and neurotoxin-sensitive manner and postsynaptic hyperpolarizations to NAD+ mimic purinergic responses to enteric inhibitory neurotransmission (Hwang et al., 2011). However frequency- and neurotoxin- insensitive ATP release and failure of MRS2179 and the more selective P2Y1 antagonist,

MRS2500, to block membrane potential changes to ATP indicates that ATP is not the enteric inhibitory purine neurotransmitter in murine and primate colons. Evidently, an integrated examination of presynaptic release mechanisms and postsynaptic effects of purines is essential to understanding purinergic mechanisms regulating colonic motility.

Moreover our studies show that mice are an excellent model for examining purinergic neurotransmission in the gut and are therefore suitable for investigating changes that occur with GI motility disorders. This could validate the power of using transgenic animals in the study of enteric purinergic signaling.

1.7.2 The role of purine metabolites in the neuroeffector junction

There are a number of possibilities that might explain why the release and effects of ATP in primate and murine GI tracts are not consistent with the endogenous motor neurotransmitter. For example, in addition to some neuronal release, a larger proportion of ATP might originate from non-neuronal sources, such as glia, interstitial cells, or smooth muscle cells, as a consequence of activity induced in these cells by nerve stimulation. Alternatively, neuronal release of ATP could occur from extra-junctional sites such as from nerve cell bodies or from nerve terminals by mechanisms not associated with Ca2+-influx and vesicle exocytosis (see Section 1.9 ‘Non-synaptic transmission’). 34

Other possibilities that might explain some of the discrepancies between ATP and the endogenous transmitter in colon are activation of multiple postjunctional receptors by

ATP or rapid degradation of ATP once released. The enzymes involved in the degradation of both ATP and NAD+ are present in GI tissues (Peri et al., 2013) where, in addition to controlling the lifetime of purine nucleotides, they produce agonists for additional purine receptors expressed by cells within GI muscles (Abbracchio et al.,

2009). ADP and ADPR, direct metabolites of ATP and NAD+, respectively, also activate

P2Y1 receptors (von Kugelgen, 2006; Mutafova-Yambolieva et al., 2007; Gustafsson et al., 2011). These nucleotides are present in the biophase thus potentially contribute to postjunctional responses elicited by purinergic inhibitory neurotransmission in GI muscles. This possibility was investigated in murine and primate colons by examining postsynaptic effects of ADP and ADPR (Durnin et al., 2012b; Chapter 4). Interestingly, these studies revealed that ADPR is bioactive in GI muscles and, similar to NAD+, mimics the pharmacology of the endogenous neurotransmitter in colon better than both

ATP and ADP (Durnin et al., 2012b). Discrepancies between ATP and the endogenous purine transmitter therefore cannot be explained by rapid metabolism in the interstitium as ADP also failed to mimic responses to inhibitory neurotransmission. Moreover, purinergic motor transmission is far more complex than typically considered with multiple purines likely contributing to inhibitory control of colonic motility (see Fig 1.7).

Consequently, we are compelled to move beyond the overly simplistic view that purinergic signaling in the gut is mediated purely by ATP. Additional studies are now warranted to establish whether ADPR is generated in the neuroeffector junction after release of NAD+ from enteric neurons or can be released as a primary neurotransmitter. 35

1.7.3 Differential sites of release of ATP and NAD+ in the myenteric plexus

At low stimulation frequencies, EFS-evoked release of NAD+ in monkey colon preparations appears exclusive to the nerve terminal whereas at higher stimulation frequencies some ganglionic release may also occur (Hwang et al., 2011; Chapter 3).

While EFS is a common approach for stimulating neural activity, it fails to differentiate the precise sources of released molecules. Therefore in order to clarify the potential sources of ATP and NAD+ in the myenteric plexus we used an alternative approach to stimulate motor neurons – that is selectively activating ligand-gated ion channel receptors localized on cell bodies and dendrites of myenteric inhibitory motor neurons (Durnin et al., 2013; Chapter 5). In addition to the P2X receptor-mediated signaling to motor neurons, ACh and 5-HT released from descending interneurons activate nicotinic ACh receptors and serotonin 5-HT3 receptors, respectively, on inhibitory motor neurons to elicit fast EPSPs, propagation and inhibitory neurotransmitter release

(Zhou & Galligan, 1999; Dickson et al., 2010b). We recently demonstrated that activation of these receptors in murine and primate colons stimulates release of ATP and

NAD+ via different mechanisms and thus likely from different sources: the release of

NAD+ required nerve action potentials and Ca2+ influx via N-type voltage-dependent

Ca2+ channels verifying release from nerve terminals. However the majority of ATP appears to be released directly from the nerve cell bodies or glia as it was not altered by disrupting action potential propagation to the nerve terminal (Durnin et al., 2013; depicted in Fig 1.6 from Smith, 2013). Further studies are warranted to determine the precise stimuli and mechanisms mediating extracellular release of ATP in GI tissues. 36

Figure 1.6. Ganglionic stimulation of motor neurons causes release of β-NAD+ at nerve + varicosities. Abbreviations: β-NAD , β-nicotinamide adenine dinucleotide; 5-HT3R, serotonergic 5-HT3 receptors; nAChR, nicotinic acetylcholine receptors; SIP, smooth muscle cells, interstitial cells of Cajal, platelet-derived growth factor receptor. Image from Smith, 2013.

1.7.4 Differential sites for degradation of ATP and NAD+ in the colon

In addition to different release sites of ATP and NAD+ in gut, there are also differences in the preferred site for degradation of these nucleotides (Durnin et al.,

2012b; Chapter 4). More pronounced degradation of exogenous NAD+ (and ADPR) in monkey colon circular muscle preparations, containing only nerve terminals, compared to whole muscles that also contain ganglia suggests that metabolism occurs close to the site of NAD+ release (i.e. nerve terminals). However ATP metabolism is not exclusive to nerve terminals (Durnin et al., 2012b) which would rather suggest that ATP may be released as a paracrine substance in ganglia or from non-neuronal sources (see Section

1.9 ‘Non-synaptic transmission’).

Interestingly in the colon, comparable degradation of exogenous NAD+ was reported in wildtype mice and in mice lacking Cd38 (Durnin et al., 2012b; Chapter 4).

Other enzymes that degrade NAD+ might be upregulated as a compensatory mechanism 37 in CD38-/- mice, or alternatively other enzymes might normally be responsible for degradation of NAD+ in the colon. CD157 shares several characteristics with CD38 exhibiting NAD-glycohydrolase and cyclase activities (Ortolan et al., 2002) and recently

CD73, a 5’-nucleotidase involved in formation of ADO from AMP, has been shown to target NAD+ directly (Garavaglia et al., 2012). Future studies are necessary to clarify the potential roles of these proteins in NAD+ degradation in GI smooth muscles.

1.7.5 Effects of purines in the colon of P2Y1R-/- mice

Pharmacological manipulation of receptor activity is a common approach to examine postsynaptic purinergic responses. Although antagonists of purine receptors are not always selective, the P2Y1 receptor antagonist, MRS2500, is thought to be highly specific for this receptor (Kim et al., 2003; Cattaneo et al., 2004). Nevertheless pharmacological approaches, as used in the studies described here, require further examination for verification of receptor mediated responses. The importance of purinergic inhibitory neurotransmission in regulating colonic motility was recently demonstrated using a mouse model with genetic deactivation of P2ry1 (Hwang et al.,

2012). Digital video recording and monitoring of artificial pellet movement down the colon revealed significantly delayed and altered transit in animals lacking the P2Y1 receptor and fIJPs were significantly reduced (Hwang et al., 2012). Moreover, membrane potential changes to ATP and ADP remained largely intact in P2Y1-/- animals whereas

NAD+ and ADPR failed to produce hyperpolarizations. This study provides particularly compelling evidence as to the identity of the enteric purine motor transmitter, favoring

NAD+/ADPR as the purine(s) involved. Clearly ATP and ADP activate receptors other 38

than P2Y1 to cause hyperpolarization in enteric smooth muscle, inconsistent with the endogenous purine neurotransmitter.

1.8 Targets of neurogenic purines in the GI tract

Neurotransmitters released from enteric motor neurons elicit effects postsynaptically via direct activation of the smooth muscle itself, or indirectly via interstitial cells that are in close apposition to varicosities of enteric motor neurons

(Sanders et al., 2010). There has been considerable debate as to whether purinergic neuromuscular transmission involves a direct action of purines on smooth muscle cells or if another cell type transduces the response. There are at least two types of interstitial cells in the tunica muscularis of the GI tract – interstitial cells of Cajal (ICC) and fibroblast-like cells, also known as platelet-derived growth factor receptor α-positive

(PDGFRα+) cells due to robust labeling for PDGFRα (Komuro et al., 1999; Pieri et al.,

2008; Sanders et al., 2010; Kurahashi et al., 2012). The anatomical localization of these cells to neurotransmitter release sites is suggestive of important physiological functions.

Together, smooth muscle cells, ICC and PDGFRα+ cells constitute the enteric SIP syncytium.

ICC are pacemaker cells of the GI tract. They provide propagation pathways for slow waves that are responsible for timing phasic contractions and constitute an interface between terminals of enteric motor neurons and smooth muscle cells to mediate a portion of the motor input from the ENS (Sanders et al., 2010). Specifically, ICC are believed to mediate responses to nitrergic (Burns et al., 1996) and cholinergic (Ward et al., 2000) 39 enteric neurotransmission. Purinergic neurotransmission, however, does not appear to be mediated by ICC (Alberti et al., 2007) and in fact enhanced purinergic responses (Burns et al., 1996) and increased expression of purinergic P2Y receptors (Sergeant et al., 2002) have been described in the W/Wv mutant mouse lacking intramuscular ICC, perhaps as a compensatory mechanism to overcome reduced nitrergic transmission in W/Wv mutants

(Ward et al., 2000; Ward et al., 2004; Ward & Sanders, 2006; Alberti et al., 2007).

Likewise, purines are unlikely to directly target the smooth muscle in colon as activation of non-selective cation currents and membrane depolarization occurs in isolated smooth muscle cells in response to purines (Hwang et al., 2011; Chapter 3).

The role of PDGFR+ cells in enteric neurotransmission has been difficult to study functionally until the development of a new reporter mouse line that has the receptor tagged with bright green fluorescent protein (Hamilton et al., 2003). PDGFR+ cells, which are also closely apposed to terminals of enteric motor neurons, connect to smooth muscle cells via gap junctions (Horiguchi & Komuro, 2000; Fujita et al., 2003) and have been revealed as a new class of excitable cells in enteric smooth muscle

(Kurahashi et al., 2011). Abundant expression of P2Y1 receptors and apamin-sensitive

SK3 channels (Kurahashi et al., 2011; Peri et al., 2013) suggested that these cells might be important in transducing purinergic responses to smooth muscles. Indeed, application of purines, including NAD+, activates apamin-sensitive, large outward currents in

+ isolated PDGFRα cells that are blocked by P2Y1 receptor antagonists indicating that

NAD+ released from enteric neurons likely targets PDGFRα+ cells in the (Kurahashi et al., 2011). Additionally Cd38 is highly expressed by PDGFRα+ cells implicating these cells as targets for NAD+ degradation (Peri et al., 2013). 40

Morphological studies have identified PDGFRα+ cells in the human colon (Kurahashi et al., 2012) suggesting a potential role in purine neurotransmission in humans.

Unfortunately PDGFRα mutant mice are embryonic lethal (Sun et al., 2000) hampering verification of the role of PDGFRα+ cells in purine motor transmission. Future

+ experiments using cell specific knockouts of P2Y1 receptors in PDGFRα cells may help confirm whether these cells are primary mediators of enteric inhibitory purinergic neurotransmission.

1.9 Non-synaptic transmission

Unlike direct synaptic communication between neurons, there is a lack of clear synaptic specializations between nerves and smooth muscles; communication at the neuromuscular junction is therefore termed non-synaptic or junctional (Vizi, 1984;

Agnati et al., 1986; Vizi et al., 2004; Burnstock, 2008a; Goyal & Chaudhury, 2013). In some cases non-synaptic transmission, which is also called volume transmission (Agnati et al., 1986), can occur when neurotransmitters spill out from the cleft and diffuse to receptors localized outside the neuroeffector junction. Alternatively, non-synaptic volume transmission can involve transmitters which are released extra-junctionally outside of nerve terminals. This release occurs largely via non-exocytotic mechanisms such as through connexin hemichannels (Stout et al., 2002), P2X7 receptor pores (Duan & Neary,

2006), volume regulated anion channels (Darby et al., 2003) or, as is the case with gasotransmitters, via diffusion (Kiss & Vizi, 2001). With the apparent extra-junctional 41 release sites of ATP in the colon (Durnin et al., 2013), these features of neurotransmission deserve consideration.

As mentioned, in the colon purine-mediated fIJPs are absent in mice lacking P2Y1 receptors (Hwang et al., 2012). Therefore if the authentic purine transmitter is released

‘in volume’ it must display selectivity at P2Y1 receptors over other purine receptors localized in enteric smooth muscles. However persistence of ATP-mediated hyperpolarizations in P2Y1-/- mice colons suggests ATP activates receptors in addition to

+ P2Y1. NAD and ADPR, on the other hand, appear to selectively activate P2Y1 receptors as hyperpolarizations to these agonists are absent in P2Y1-/- mice (Hwang et al., 2012).

+ Thus if released ‘in volume’, responses to NAD /ADPR could be mediated by P2Y1 receptors expressed throughout GI muscles.

At some smooth muscle neuroeffector junctions, however, close juxtaposition of nerve terminals and effector cells allows communication over a short distance similar to that of a synapse. Thus transmission can occur at organized compartments where released transmitters interact with restricted pools of receptors clustered on the postjunctional membrane (Hirst et al., 1992). This appears to be the case for purinergic motor transmission where purines released from inhibitory motor neurons likely target closely apposed postjunctional PDGFRα+ cells (described in Section 1.8) that abundantly express purinergic P2Y1 receptors (Kurahashi et al., 2011). Therefore if ATP functions as a purine neurotransmitter, it cannot be released ‘in volume’ into the interstitial space surrounding smooth muscle cells as receptors other than P2Y1 would be activated to contribute to enteric inhibitory responses. This is not consistent with our evidence demonstrating extra-junctional release sites of ATP in the colon (Durnin et al., 2013). 42

In summary, NAD+ and/or ADPR appear to be the primary neurotransmitters mediating purinergic inhibitory regulation of colonic motility. Upon action potential firings, NAD+/ADPR can be released from vesicles in myenteric nerve terminals into the

+ neuroeffector junction. NAD /ADPR then bind to P2Y1 receptors localized on postjunctional PDGFRα+ cells activating intracellular Ca2+ release pathways that lead to apamin-sensitive SK3 channel activation and generation of large outward currents.

Hyperpolarization transients then conduct from PDGFRα+ cells to the smooth muscle via gap junctions causing smooth muscle relaxation (Fig 1.7). Our studies demonstrate the complexity of enteric purinergic signaling and suggest that the NAD+/ADPR system may be a novel target for treating motility disorders of the large intestine.

43

Figure 1.7. A simplified model of purine-mediated neurotransmission in the colon smooth muscle neuroeffector junction (NEJ). NAD+ is the primary neurotransmitter mediating purinergic inhibitory regulation of colon motility. ADPR, a metabolite of NAD+, could also be a factor in inhibitory neurotransmission, either as a neurotransmitter or as a metabolite of extracellular NAD+. NAD+ is stored in and released from neural vesicles in nerve varicosities into + the NEJ upon action potential firings (APs).NAD binds to postjunctional P2Y1 receptors on PDGFRα+ cells and mediates activation of phospholipase (PLC), formation of inositol 1,4,5- 2+ trisphosphate (IP3), release of Ca from intracellular stores (i.e., sarcoplasmic reticulum, SR), and activation of small-conductance K+ channels (SK3). Hyperpolarization transients (i.e. IJPs) conduct to SMCs via gap junctions (GJs) between PDGFRα+ cells and SMC. IJPs reduce excitability and cause muscle relaxation. NAD+ is rapidly degraded to ADPR by CD38 and/or an unknown metabolic pathway. ADPR is further degraded to AMP and adenosine (ADO) by well- known pathways (not shown). ADPR also binds to postjunctional P2Y1 receptors and produces responses identical to NAD+. ATP is also released in colonic muscles upon electrical field stimulation, but it appears to originate primarily from non-neuronal sources. ATP is quickly degraded to ADP, AMP, and ADO. ADO can activate P1 -coupled receptors on SMC membrane. ATP and ADP may participate in the postjunctional responses mediated by P2Y1 receptors (dotted arrows), but data suggest that ATP also binds to additional P2Y receptors (P2YRs) expressed by SMCs and possibly other cell types. Purine binding to P2YRs in SMCs activates SK channels and nonselective cation channels (NSCC). With cells at physiological gradients and potentials, the dominant responses of SMCs to ATP and NAD+ are activation of inward current. Thus, it is unlikely that IJPs due to transduction of the purinergic neurotransmitter signal could be mediated by SMCs. Image from Mutafova-Yambolieva, 2012.

44

1.10 Purinergic neurotransmission in the central nervous system

1.10.1 Overview of purinergic signaling in the central nervous system

There is surprisingly little known about purinergic neurotransmission in the CNS.

In the late 1970s, using the luciferin-luciferase chemiluminescence assay ATP was reported to be released from whole brain synaptosomes treated with the sodium channel activator, veratridine, or high extracellular K+ (White, 1977) stimulating interest in the potential functions of ATP in the brain. Later, exogenous ATP was shown to elicit robust inward currents in dorsal horn neurons (Jahr & Jessell, 1983) and isolated sensory neurons (Krishtal et al., 1983) but it wasn’t until the 1990s that endogenous P2X receptor-mediated excitatory postsynaptic currents were recorded in medial habenula neurons (Edwards et al., 1992). While not identified in every brain region, P2X-mediated currents are apparent in the spinal cord (Bardoni et al., 1997), hippocampus (Pankratov et al., 1998; Mori et al., 2001), locus coeruleus (Nieber et al., 1997) and somatosensory cortex (Pankratov et al., 2002; Pankratov et al., 2003).

In the brain, ATP is almost exclusively believed to be the purine neurotransmitter responsible for excitatory currents (Burnstock, 2013). However, just as in the periphery, this claim is made primarily with indirect evidence such as similar response of the endogenous transmitter and exogenous ATP and block of responses by P2 antagonists

(Bardoni et al., 1997; Nieber et al., 1997; Pankratov et al., 1998; Pankratov et al., 2003).

Studies which have examined release of ATP in brain regions including cortex (White et al., 1980; Salgado et al., 1997), cerebellum (Terrian et al., 1989), spinal cord (Sawynok et al., 1993), hippocampus (Wieraszko et al., 1989) and hypothalamus (Sperlagh et al., 45

1998) claim that ATP is likely released as a neurotransmitter due to the Ca2+-dependence of release. However reference must again be made to the possible vesicular localization of ATP for regulating steps neurotransmitter uptake and in exocytosis (Sperlagh & Vizi,

1996). In fact, some investigators have suggested that without any regulatory mechanism to limit uptake, ATP can accumulate in all synaptic vesicles and through all stages of vesicle formation and recycling in the CNS (see (Pankratov et al., 2006). Thus it is acknowledged that ATP may be present in vesicles for functions other than neurotransmission.

Accumulating evidence also demonstrates ATP release from non-neuronal cells in the CNS. For example ATP is released from glial cells by numerous mechanisms including connexin hemichannels (Stout et al., 2002), P2X7 receptor pores (Duan &

Neary, 2006), volume regulated anion channels (Darby et al., 2003) as well as by vesicular exocytosis (Coco et al., 2003; Montana et al., 2006). Additionally, vesicular release of ATP from has recently been demonstrated (Imura et al., 2013). Thus in the CNS ATP could be released from both neuronal and non-neuronal sources by both

Ca2+-dependent and Ca2+-independent mechanisms. As a result the question of whether

ATP in fact functions as a fast neurotransmitter in the brain remains unanswered. These observations as well as the strong evidence accumulated for possible neurotransmitter role of NAD+ in the peripheral nervous system led us to also examine the potential role of

NAD+ in the CNS.

1.10.2 NAD+ as a putative neurotransmitter in the central nervous system 46

While recent studies have demonstrated the release of ADO from neurons in the hippocampus (Lovatt et al., 2012) and cerebellum (Wall & Dale, 2007) which is involved in modulating neuronal excitability, studies examining the roles of other purines in the brain are generally lacking. Previously release of NAD+ from cultured hippocampal has been demonstrated (Verderio et al., 2001), and earlier studies suggested a potential neuromodulatory role of NAD+ in rat and guinea-pig hippocampus (Richards et al., 1983; Snell et al., 1985; Galarreta et al., 1993). However until recently, the potential role of NAD+ as a neurotransmitter in the CNS had not been proposed or determined.

Work in our laboratory has demonstrated that NAD+ may participate in neurotransmission in the CNS (Durnin et al., 2012a; Chapter 6) In isolated rat forebrain synaptosomes, high-K+ membrane depolarization evoked release of NAD+ that was proportional to the membrane depolarization and required intact vesicle exocytosis machinery, i.e. release was attenuated by cleavage of SNAP-25 with BoNT/A, by inhibition of N-type voltage dependent Ca2+ channels with ω-conotoxin GVIA and by inhibition of the proton gradient of synaptic vesicles with bafilomycin A1 (Durnin et al.,

2012a), which is consistent for a neurotransmitter. In addition, we demonstrated that localized application of exogenous NAD+ in cortical neurons caused rapid Ca2+ transients that were similar to those produced by exogenous ATP (see below) suggesting that endogenously released NAD+ could participate in neuron-to-neuron communication in the brain. Exogenous ADPR similarly produced Ca2+ transients in cortical neurons

(Durnin et al., 2012a). Thus once again our studies highlight that neuronal excitability in the brain may also be regulated by multiple purine substances. Interestingly, we found that subpopulations of cortical neurons appeared to respond preferentially to ATP or to 47

NAD+/ADPR which might suggest different mechanisms of action of these purines in the brain. Defining these distinct subpopulations of neurons may shed light on the functional significance of ATP and NAD+/ADPR in the brain.

In this study we also demonstrated expression of CD38 on synaptosomes expressing markers synaptophysin and chromogranin B, and degradation of exogenous NAD+ in contact with synaptosomes; thus a mechanism for terminating the extracellular actions of NAD+ is present in the brain. An additional uptake mechanism also appears to be present possibly as an alternative means for terminating or recycling extracellular NAD+. At present we do not know the nature of this transporter. Import of

NAD+ through connexin hemichannels has been suggested previously (Bruzzone et al.,

2001; Song et al., 2011; Okuda et al., 2013) but our studies rule against the involvement of these channels in NAD+ transport in brain synaptosomes (Durnin et al., 2012a). An alternative candidate is the chloride-dependent vesicular nucleotide transporter,

SLC17A9, which is highly expressed in the brain (Abbracchio et al., 2009) and has been shown to transport in vitro other purine nucleotides such as ATP, ADP and GTP (Sawada et al., 2008). Further studies are necessary to elucidate the potential role of SLC17A9 in

NAD+ transport in the CNS.

In summary our experiments provide the first evidence demonstrating a putative neurotransmitter role of NAD+/ADPR in the CNS. While we do not exclude the possibility that ATP may function as a neurotransmitter in some CNS neurons, we emphasize the importance in considering purines other than ATP that could also participate in regulating neuronal excitability in the brain. As in the periphery, the 48

NAD+/ADPR system could represent a novel therapeutic target for disorders associated with disrupted purinergic mechanisms in the brain.

1.11 Summary

NAD+ is a key intracellular regulator of energy metabolism. Increasing evidence also demonstrates important extracellular roles of NAD+ in cellular functions such as

ADP-ribosylation of proteins, regulation of plasma membrane channels and enzymes and activation of P2 receptors (Billington et al., 2006). Also indicative of important extracellular roles of NAD+ is the extracellular facing catalytic site of CD38. However the source and mechanisms of NAD+ efflux and the precise extracellular actions of NAD+ in different systems are not entirely clear.

Throughout the last decade ongoing work in Mutafova-Yambolieva’s laboratory has provided novel information about the extracellular signaling roles of NAD+ in the peripheral and central nervous systems. In 2004 evidence was presented demonstrating regulated release of NAD+ during stimulation of nerves in the peripheral nervous system; a putative neurotransmitter or neuromodulator role of NAD+ in smooth muscle was suggested. These pioneering studies are among the first to demonstrate regulated release of NAD+ into the extracellular compartment and the first to show this phenomenon in complex smooth muscle tissues. As well as providing novel information about NAD+ efflux from cells, these original findings have also had an impact in the field of intracellular NAD+ signaling as they revealed novel intracellular storage sites of NAD+ in vesicles that had not been previously identified. A summary of the known mechanisms of 49 release and actions of extracellular NAD+ is shown in Fig 1.8 (Billington et al., 2006) which is illustrated to include the vesicular release of NAD+ as put forward by our laboratory.

The work described in this dissertation continues to explore the extracellular role of NAD+ in the nervous system by examining mechanisms of release, metabolism and action of NAD+ in the enteric and central nervous systems. We provide the first direct evidence for storage of NAD+ in synaptic vesicles and release via exocytosis (Chapter 2).

We demonstrate that NAD+ is a better candidate than ATP as the enteric inhibitory neurotransmitters in the colon, challenging the decades old notion claiming ATP to be the transmitter (Chapters 3, 4 and 5). We also identify a novel extracellular signaling role of the NAD+ metabolite ADPR in the gut that has never before been demonstrated (Chapter

4). Finally, we provide the first evidence that NAD+/ADPR are candidate neurotransmitters in the CNS (Chapter 6): we show vesicular release, extracellular degradation and reuptake of NAD+ in rat brain synaptosomes and stimulation of postsynaptic neurons by exogenous NAD+ and ADPR.

Together our work uncovers novel information on the extracellular roles of

NAD+/ADPR in the nervous system, moves the field beyond the sovereign role of ATP and highlights the complexity in understanding purinergic regulation of enteric and central nervous system function. The NAD+/ADPR system may represent a novel therapeutic target for pathological conditions resulting from altered purinergic signaling.

50

Figure 1.8.The Known Signaling Functions of Extracellular NAD(P). NAD acts as a substrate for ectoenzymes on the extracellular surface of the plasma membrane. ADP-ribosylation by ARTs regulates the function of a wide range of cell surface enzymes, receptors and channels while CD38 produces a number of active metabolites from NAD(P). The Ca2+-releasing CD38 metabolites cADPR and NAADP are transported across the plasma membrane to reach their intracellular sites of action while NAD can be released or transported out of the cell. In addition, NADP and NAADP have been shown to act on P2X receptors, while ADPR and NAD act directly on yet unidentified plasma membrane receptors/channels. CD38 plays a central role in these processes by regulating the concentrations of all of the nucleotide players. Image from Billington et al., 2006.

51

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71

Chapter 2

Storage and secretion of β-NAD, ATP and dopamine in NGF-differentiated rat pheochromocytoma PC12 cells

Ilia A. Yamboliev, Lisa M. Smyth, Leonie Durnin, Yanping Dai and Violeta N. Mutafova-Yambolieva Published in European Journal of Neuroscience 2009 September; 30(5):756- 768.

72

2.1 Abstract

In nerve-smooth muscle preparations β-nicotinamide adenine dinucleotide (β-

NAD) has emerged as a novel extracellular substance with putative neurotransmitter and neuromodulator functions. Thus, β-NAD is released along with noradrenaline and adenosine 5'-triphosphate (ATP) upon firing of action potentials in blood vessels, urinary bladder and large intestine. At present it is unclear whether noradrenaline, ATP and β-

NAD are stored in and released from common populations of synaptic vesicles. This matter is unattainable to study in complex systems such as nerve-smooth muscle preparations. Adrenal chromaffin cells are used here as a single-cell model to examine mechanisms of concomitant neurosecretion. Using high-performance liquid chromatography techniques with electrochemical and fluorescence detections we simultaneously evaluated secretion of dopamine (DA), ATP, adenosine 5'-diphosphate, adenosine 5'-monophosphate, adenosine, and β-NAD and its immediate metabolites

ADP-ribose and cyclic ADP-ribose in superfused nerve growth factor-differentiated rat pheochromocytoma PC12 cells. β-NAD, DA, and ATP were released constitutively and upon stimulation with high-K+ solution or nicotine. Botulinum neurotoxin A tended to increase the spontaneous secretion of all substances and abolished the high K+-evoked release of β-NAD and DA but not of ATP. Subcellular fractionation by continuous glycerol and sucrose gradients along with immunoblot analysis of the vesicular marker proteins synaptophysin and secretogranin II revealed that β-NAD, ATP and DA are stored in both small synaptic-like vesicles and large dense-core-like vesicles. Yet, the three substances appear to have different preferential sites of release upon membrane 73 depolarization including sites associated with SNAP-25 and sites non-associated with

SNAP-25.

2.2 Introduction

In the peripheral sympathetic nervous system noradrenaline (NA) and adenosine

5'-triphosphate (ATP) are assumed to be primary cotransmitters (Burnstock, 1990;

Todorov et al., 1996). We have recently found that β-nicotinamide adenine dinucleotide

(β-NAD) is also released along with NA and ATP upon nerve stimulation in blood vessels (Smyth et al., 2004), urinary bladder (Smyth et al., 2004; Breen et al., 2006), and large intestine (Mutafova-Yambolieva et al., 2007). In these nerve-smooth muscle preparations the release of β-NAD correlates with neural activity and requires intact neural fast Na+ channels, N-type voltage-operated calcium channels, and the 25-kDa synaptosomal associated protein SNAP-25 (Smyth et al., 2004; Smyth et al., 2006b;

Breen et al., 2006; Mutafova-Yambolieva et al., 2007; Smyth et al., 2009). Exogenous β-

NAD reduces the release of NA in blood vessels (Smyth et al., 2004), inhibits spontaneous contractions in the bladder (Breen et al., 2006), induces vasodilatation or vasoconstriction in the mesenteric vasculature (Smyth et al., 2009) and causes membrane hyperpolarization and relaxation in the murine colon (Mutafova-Yambolieva et al.,

2007). Therefore, β-NAD is likely a novel neurotransmitter and a novel neuromodulator.

Moreover, in some systems β-NAD, but not ATP, may be the purine neurotransmitter

(Mutafova-Yambolieva et al., 2007) and hence, the release of ATP and β-NAD might occur at different sites or be mediated by different mechanisms. Understanding these 74 mechanisms will forward our knowledge of the roles of extracellular β-NAD in the context of adrenergic-purinergic cotransmission and may suggest new possibilities for synaptic regulation. At present only a few studies in nerve-smooth muscle preparations provide fairly indirect information about possible co-storage and co-release of β-NAD with either NA or ATP (i.e., Smyth et al., 2009; Mutafova-Yambolieva et al., 2007;

Bobalova & Mutafova-Yambolieva, 2006).

Nerve-smooth muscle preparations contain multiple cell types including smooth muscle cells, nerve endings, endothelial cells, and fibroblasts that make studies on vesicular storage and release of neurotransmitters in smooth muscles a challenging task.

Instead, we employed cultured rat pheochromocytoma PC12 cells, a single cell-type model commonly utilized for elucidating basic mechanisms of neurosecretion. These cells phenotypically resemble sympathetic neurons and contain both small vesicles and dense core granules (Greene & Tischler, 1976). ATP is stored together with catecholamines (mainly dopamine, DA) in secretory vesicles and the contents are released by Ca2+-dependent exocytosis (Wagner, 1985). To the best of our knowledge no information is available about storage and secretion of β-NAD in adrenal chromaffin cells. Here we show that in nerve growth factor-differentiated PC12 cells β-NAD, ATP, and DA are all present in small synaptic-like vesicles (SSVs) and large dense-core-like vesicles (LDCVs). All compounds, including β-NAD, are subject to spontaneous and regulated release. Yet, disruption of SNAP-25 by botulinum neurotoxin A (BoNT/A) abolished the high K+-evoked release of β-NAD and DA, but not of ATP, suggesting that the two purines β-NAD and ATP might have different preferential sites of release upon membrane depolarization. 75

2.3 Methods

2.3.1 Cell culture

PC12 (rat adrenal pheochromocytoma) cells were obtained from the American

Type Culture Collection (ATCC, Manassas, VA; #CRL-1721) and grown on 225 cm2 collagen-coated cell culture flasks in F12K medium (ATCC, #30-2004) supplemented with 2 mM L-glutamine, 5% fetal bovine serum (FBS), 10% horse serum, 100 units/ml penicillin, and 100 units/ml streptomycin until 75–80% confluence. To induce differentiation, cells were incubated in F12K medium supplemented with 0.1% FBS and

50 ng/ml murine nerve growth factor (NGF, 2.5S, G514, Promega Corporation, Madison,

WI): incubation continued for 7 days with change of culture medium every second day.

2.3.2 Overflow experiments

PC12 cells (3×106) were applied under suction onto Whatman GF/B filter paper

(3 mm diameter) which was placed in a 0.45 μm Cameo 3N syringe filter serving as a perfusion chamber (modification of a method described by (Todorov et al., 1996). The cells in the chamber were perfused with a superfusion solution at a rate of 0.8 ml/min.

The composition of the superfusion solution was as follows (mM): NaCl, 140; KCl, 4.7;

MgCl2, 1.2; CaCl2, 2.5; dextrose, 11; HEPES, 10, pH 7.4. After an equilibration period of

30 min, 800 μl of superfusate were collected as pre-stimulation samples. The cells were then stimulated for 20 min with 60 mM KCl. The high-potassium-containing solution was prepared by substituting NaCl for an equal amount of KCl to maintain isotonicity.

Samples of superfusate (approximately 800 μl) were collected at 1', 2', 5', 10' and 20' time 76 intervals in ice-cold Eppendorf tubes. Each sample was divided into two parts for measuring the content of catecholamines and purines as previously described (Mutafova-

Yambolieva et al., 2007; Smyth et al., 2004). In some experiments stimulation with either

25 mM KCl or 100 μM nicotine was also performed.

2.3.3 Treatment with BoNT/A

Cells grown to 80% confluence were exposed to 30 nM botulinum neurotoxin A

(BoNT/A, List Biological Laboratories, Campbell, CA) for 16 h before performing either overflow experiments or western immunoblot analysis of SNAP-25. BoNT/A was dissolved in BSA (1 mg/ml) as recommended by the manufacturer and further diluted in cell culture medium.

2.3.4 Fractionation of synaptic vesicles by glycerol and sucrose gradient centrifugation

For velocity sedimentation, we applied a modified method by Clift-O'Grady et al.

(1990) and Melikian & Buckley (1999) summarized in Fig. 2.1. Differentiated cells were scraped into homogenization buffer A containing (mM): NaCl, 150; HEPES 10, pH 7.4;

EGTA 1.0; MgCl2 0.1, pelleted (300xg, 5 min, room temperature) and resuspended in 2 ml buffer A containing protease inhibitors (1.0 mM PMSF and 1.0 μg/ml each leupeptin, aprotinin, and pepstatin). The cells were then homogenized (20 strokes up-and-down) in a teflon/glass homogenizer and postnuclear supernatant was obtained after centrifugation at

1000xg for 7 min at 4°C. The pellet was rewashed and centrifuged again, and the two postnuclear supernatants were pooled together. The sample was centrifuged at 10,000xg for 5 min at 4°C to obtain LDCVs in the P2 pellet, and SSVs in the S2 supernatant. S2 77 supernatant was then layered onto a continuous 5–25% glycerol gradient over a 1-ml

60% (1.75 M) sucrose pad, and SSVs were fractionated by centrifugation at 220,000xg for 1 hour at 4°C (SW41 Ti rotor, on a Beckman L8-70M Ultracentrifuge, Beckman

Coulter, Inc., Fullerton, CA). P2 pellet was resuspended in 1 ml buffer containing 10 mM

HEPES/KOH, pH 7.4, 0.3 M sucrose, and protease inhibitors, and layered onto a continuous 0.3–1.5 M sucrose gradient over a 1-ml 60% (1.75 M) sucrose pad. LDCVs were fractionated by centrifugation at 85,000xg for 3 hours at 4°C (SW41 Ti rotor, on a

Beckman L8-70M Ultracentrifuge, Beckman Coulter, Inc., Fullerton, CA). After centrifugation, tubes were punctured with 18-gauge syringe needles just above the sucrose pads, and 0.5-ml fractions were collected from the glycerol and sucrose gradients. Each sample was sonicated 3×15 sec, vortexed and then centrifuged at

15,000xg for 15 min at 4°C to remove insoluble material. Glycerol and sucrose concentrations were determined by measuring the refractive index of each fraction and converting to percentage glycerol or moles sucrose using a glycerol or sucrose standard/calibration curves. Four hundred μl aliquots were processed for HPLC analysis of purines (80 μl), catecholamines (60 μl) and HPLC fractions (260 μl) as described in

Preparation of samples for purine detection, HPLC assay of etheno-purines, HPLC fraction analysis, Preparation of samples for catecholamine detection, and HPLC assay of catecholamines, The remaining samples (~ 100 μl) were for analysis of vesicular markers by immunoblotting.

2.3.5 Preparation of samples for purine detection 78

A modified method of Levitt et al., 1984 was employed to detect 1,N6-etheno- derivatives of the endogenous purines present in the cell superfusates as described previously (i.e., Bobalova et al., 2002). Briefly, 100 μl of a citrate phosphate buffer (pH

4.0) was added to 200 μl of the superfusate sample in Eppendorf tubes. Ten μl of 2- chloroacetaldehyde was added to the samples in a fume hood; the samples were then heated for 40 min at 80°C in a dry bath incubator (Fisher Scientific, USA). Using this procedure endogenous ATP, ADP, AMP, and adenosine (ADO) are derivatized to 1,N6- etheno-ATP (eATP), 1,N6-etheno-ADP (eADP), 1,N6-etheno-AMP (eAMP), and 1,N6- etheno-ADO (eADO), respectively (Bobalova et al., 2002). The endogenous β-NAD,

ADP-ribose (ADPR) and cyclic ADPR (cADPR) are derivatized to 1,N6-etheno-ADP- ribose (eADPR) as described previously (Smyth et al., 2004). The reactions were stopped by placing samples on ice.

2.3.6 HPLC assay of etheno-purines

The HP1100 liquid chromatography module system (Agilent Technologies,

Wilmington, DE) was used throughout this study and has been previously described

(Bobalova et al., 2002). The mobile phase consisted of 0.1 mol/L KH2PO4 (pH 6.0) as eluent A; eluent B contained of 35 % methanol and 65 % eluent A. Gradient elution was employed according to the following linear program: time 0, 0 % eluent B; 18 min, 100

% eluent B. Flow rate was 1 ml/min and run time was 20 min. Column temperature was ambient while the autosampler temperature was 4°C. The fluorescent detector recorded signals at an excitation wavelength of 230 nm and emission wavelength of 420 nm, which are the optimum conditions for detecting etheno-derivatives of nucleotides and 79 nucleosides (Bobalova et al., 2002). The amounts of purines in each sample were calculated from calibration curves of purine standards run simultaneously with every set of unknown samples. Results were normalized for sample volume and cell count and the overflow of purines was expressed in pmol/106 cells.

2.3.7 Sample concentration and HPLC fraction analysis

As aforementioned β-NAD, ADPR and cADPR elute as a single peak of eADPR at 11.2 min. To identify the ratio of β-NAD:ADPR:cADPR forming eADPR we carried out an HPLC fraction analysis as described previously (Smyth et al., 2004). Briefly, in some overflow experiments 2.4-ml cell superfusate samples containing either 5 mM KCl

(pre-stimulation sample) or 60 mM KCl for 5 min (stimulation sample) were collected in

Eppendorf tubes and were immediately immersed in liquid N2. Control samples from solutions containing either 5 mM KCl or 60 mM KCl with no contact with cells were also collected. The samples were then concentrated by Speed Vacuum (Savant SVC100,

Thermo Electron Corp., Westmont, IL) to 1 ml volume. Nine hundred μl of each concentrated sample were injected into the HPLC system and 400 μl-fractions corresponding to the retention times of cADPR (7.0–7.4 min, “7.2-min fraction”), ADPR

(8.3–8.7 min, “8.5-min fraction”), and β-NAD (10.3–10.7 min, “10.5-min fraction”) were collected in Eppendorf tubes containing 180 μl citric buffer. The exact retention times for the three nucleotides were determined by injecting authentic β-NAD, cADPR and ADPR standards (20 nmol/injection) in the same sequence prior to the concentrated superfusate samples. The HPLC fractions were further subjected to etheno-derivatization with 20 μl

2-chloroacetaldehyde as described in Preparation of samples for purine detection. The 80 derivatized samples were injected into the HPLC and analyzed for eADPR content.

HPLC-fraction analysis was also performed with samples collected during glycerol and sucrose fractionation. Since these experiments were carried out with larger number of cells (>50×106 cells) there was no need of concentrating the samples prior collecting

HPLC fractions. For these experiments 240 μl of each glycerol or sucrose fraction was injected in the HPLC system and the samples were processed further as described above.

2.3.8 Preparation of samples for catecholamine detection

One hundred and fifteen μl of superfusate solution was acidified to pH 2.6 with 3

μl 1 N perchloric acid and the samples were filtered through 0.22 μm Cameo 3N syringe filters.

2.3.9 HPLC assay of catecholamines

The overflow of catecholamines was assayed as described previously (Mutafova-

Yambolieva et al., 2007). Briefly, 115-μl aliquots from the samples were acidified with 3

μl 1 M perchloric acid to pH 2.6 and injected (70 μl) into an isocratic HP1100 HPLC system equipped with an HP1049A electrochemical detector (Agilent Technologies,

Wilmington, DE, USA) and a MD-150 column (ESA Inc., Chelmsford, MA, USA). The mobile phase for separation consisted of the following (mmol/L): 50 Na2PO4; 0.2 EDTA;

3.0 l-heptanesulfonic acid, 10 LiCl, and methanol 3 % v/v in deionized water (pH 2.6).

The HPLC systems were controlled, and data collected, by a Dell computer equipped with HP ChemStation (A.10.02) software from Agilent Technologies (Wilmington, DE,

USA). The amounts of DA in each sample were calculated from calibration curves of 81 catecholamine standards run simultaneously with every set of unknown samples. Results were normalized for sample volume and cell count and the overflow of DA was expressed in pmol/106 cells.

2.3.10 Western immunoblot analysis of SNAP-25 in total PC12 cell extracts

PC12 cells from control and BoNT/A-treated groups were washed 3×5 ml cell culture medium, scraped, solubilized in 200 μl RIPA buffer, sonicated 3×30 s, incubated on ice, vortexed, and centrifuged at 15,000xg for 20 min at 4°C. Total protein concentration of the supernatant was determined by the Bradford assay (BioRad kit,

Hercules, CA) using bovine serum albumin (BSA) for standards. Cell homogenates were reduced with Laemmli sample buffer and equal amounts of total protein (10 μg) were resolved by SDS-PAGE (15% acrylamide) and transferred onto nitrocellulose membranes for 1.5 hours at 24V and 4°C (Genie blotter, Idea Scientific Company, Minneapolis,

MN). Membranes were blocked for 1 hour with LI-COR blocking buffer (LI-COR, Inc.,

Lincoln, NE) and probed for 18 hours at 4°C with a SNAP-25 primary mouse monoclonal antibody (Sternberger Monoclonals Inc., Lutherville, MD), diluted 1000-fold in LI-COR buffer. After removal of excess primary antibody, membranes were incubated for 45 min at room temperature with a secondary mouse antibody coupled to IR800 infrared marker (emission wavelength 800 nm, Rockland Immunochemicals, PA), diluted

100,000-fold in LI-COR buffer. Images were obtained with an infrared Odyssey scanner

(LI-COR, Inc., Lincoln, NE). For positive controls on immune blots we used 30 μg of total protein, obtained from rat brain tissue homogenized in RIPA buffer.

82

2.3.11 Western immunoblot analysis of vesicular protein markers and SNAP-25 in glycerol and sucrose gradient fractions

Cell fractions were prepared as described in Fractionation of synaptic vesicles by glycerol and sucrose gradient centrifugation. Proteins from equal fraction volumes were resolved by SDS-PAGE and transferred onto nitrocellulose membranes. Membranes were blocked and incubated with mouse anti-synaptophysin (Chemicon International,

Tamecula, CA, MAB368, dilution 1:1000) as an SSV marker (Jahn et al., 1985; Cutler &

Cramer, 1990), rabbit anti-secretogranin II (Santa Cruz Biotechnologies, Santa Cruz, CA, sc-50290, dilution 1:100) as a LDCV marker (Fischer-Colbrie et al., 1995; Cutler &

Cramer, 1990), and mouse anti-SNAP25 (Sternberger Monoclonals Inc., Lutherville,

MD, SMI-81, dilution 1:1000) as a marker of plasma membranes (Sollner et al., 1993;

McMahon & Sudhof, 1995; Sorensen, 2005) and were further processed as described in the Western immunoblot analysis of SNAP-25 in total PC12 cell extracts section.

2.3.12 Statistics

Data are presented as means ± SEM. One-way ANOVA with Bonferroni's multiple comparison post test was performed using GraphPadPrism v. 3 for Windows

(GraphPad Software, San Diego, CA) when three or more groups of data were compared.

Unpaired two-tailed Student's t-test was performed using the same software when two groups of data were compared. A probability value of less than 0.05 was considered significant.

83

2.4 Results

2.4.1 Spontaneous secretion of catecholamines and purines in NGF-treated PC12 cells

Differentiated PC12 cells perfused with normal K+ solution (5 mM KCl) spontaneously secreted DA (1.93±0.48 pmol/106 cells), ATP (0.106±0.03 pmol/106 cells), ADP (1.023±0.16 pmol/106 cells), AMP (0.752±0.134 pmol/106 cells), and ADO

(0.467±0.075 pmol/106 cells), n=13–21. Figs. 2.2 and 2.3 show the values of spontaneously secreted DA and purines (at KCl 5 mM) in the controls for BoNT/A- treated cells, discussed below. Spontaneous release of a mixture of β-NAD, ADPR and cADPR was also detected; due to etheno-derivatization at 80°C, the components of this cocktail elute as one peak of eADPR with a retention time of 11.2 min (Fig. 2.3A, KCl 5 mM). Thus, the superfusates collected before stimulation contained 0.410±0.061 pmol/106 cells, n=21, of the mixture; these amounts were calculated based on the assumption that β-NAD is the major component of the peak (see results from fraction analysis shown in Distribution of β-NAD, ADPR and cADPR in cell superfusates determined by HPLC fraction analysis). Total purines (ATP+ADP+AMP+ADO+β-

NAD+ADPR+cADPR) secreted in the superfusates were 3.00±0.42 pmol/106 cells, n=21.

Noradrenaline was not detected in the samples collected during perfusion of the PC12 cells with 5 mM KCl solution, likely due to concentrations of noradrenaline below detection limits.

2.4.2 Secretion of catecholamines and purines evoked by 60 mM KCl and effects of

BoNT/A 84

Perfusion of PC12 cells with 25 mM KCl for 20 minutes caused no additional release of DA and purines (data not shown), suggesting that higher concentrations of KCl are necessary to evoke measurable effects in these cells. Therefore, in the majority of the study 60 mM KCl was used to stimulate the PC12 cells. The high-K+ (60 mM KCl) solution evoked additional release of DA and purines with greatest release occurring within 2 min of contact with the cells (Fig. 2.2A,C-control and Fig. 2.3A, C–H-control).

Secretion of DA and purines returned to basal (pre-stimulation) levels by about the 5th minute of superfusion of 60 mM KCl. Thus, DA was 3.045±0.971 pmol/106 cells before stimulation, 10.391±2.615 pmol/106 cells at 2' (P<0.01, t=4.184, F=4.857), and

4.335±0.661 pmol/106 cells at 5' (P>0.05), one-way ANOVA, followed by Bonferroni's multiple comparison posttest, n=5. ATP was 0.128±0.026 pmol/106 cells before stimulation, 0.324±0.064 pmol/106 cells at 2' (P<0.05, t=3.363, F=4.021), and

0.192±0.041 pmol/106 cells at 5' of stimulation with high K+ solution (P>0.05, t=1.092), one-way ANOVA, followed by Bonferroni's test, n=5. ADP and AMP greatly exceeded the amounts of ATP in the superfusate samples collected before and during stimulation, which is in agreement with previously published results in bovine chromaffin cells (i.e.

Todorov et al., 1996; Kasai et al., 1999). The chromatographic peaks formed by β-

NAD+ADPR+cADPR appear small due to the very low fluorescence coefficient of β-

NAD generating eADPR (see Fig. 2.3 inset). However, when compared to standards the amounts of β-NAD+ADPR+cADPR (which comprises mainly β-NAD, see below) is much greater than other purines, including ATP (Fig. 2.3 C,F). Thus, the overflow of β-

NAD was 0.379±0.071 pmol/106 cells before stimulation, 0.975±0.147 pmol/106 cells at

1' (P<0.05, t=3.351), 0.994±0.210 pmol/106 cells at 2' of stimulation (P<0.05, t=3.459), 85 and 0.627±0.102 pmol/106 cells at 5' of stimulation with KCl 60 mM (P>0.05, t=1.457), one-way ANOVA, followed by Bonferroni's test. At 10' and 20' of KCl stimulation the overflow of ATP and β-NAD was not significantly different from pre-stimulation values

(P>0.05), Fig. 2.3.

We next examined whether disruption of SNAP-25 with BoNT/A would affect the spontaneous and evoked by 60 mM KCl release of DA and purines. Immunoblot analysis of PC12 cells revealed a single immunoreactive band with molecular size of ~25 kDa, representing intact SNAP-25 (Fig. 2.2D, lane 1). Incubation of PC12 cells with

BoNT/A produced a secondary immunoreactive band with apparent molecular size of

~24 kDa (lane 2), which was not found in control cell protein extracts (lane 1). The 24- kDa band likely represents degradation product(s) of SNAP-25 owing to cleavage by

BoNT/A (Blasi et al., 1993b). BoNT/A tended to increase the spontaneous secretion of

DA and all purines (see pre-stimulation samples at KCl 5 mM in Figs. 2.2 and 2.3, controls vs. BoNT/A-treated, P<0.05 for ADP, AMP, and total purines, P>0.05 for DA,

ATP, β-NAD and ADO, unpaired t-test, two-tailed. The high K+-evoked release of DA

(Fig. 2.2C), ADP, AMP, ADO and the mixture of β-NAD, ADPR and cADPR (Fig.

2.3D–H) was significantly attenuated and shortened within 1 minute of perfusion with 60 mM KCl in BoNT/A-treated cells. The amount of released ATP was not significantly changed in BoNT/A-treated cells (Fig. 2.3B, C): thus, in BoNT/A-treated cells ATP overflow was 0.154±0.006 pmol/106 cells before stimulation and 0.318±0.049 pmol/106 cells at 1' of stimulation (P<0.05, t=3.387, one-way ANOVA, followed by Bonferroni's posttest). However, the overflow of β-NAD in the same samples was 0.528±0.123 pmol/106 cells before stimulation, 0.616±0.015 pmol/106 cells at 1'of stimulation 86

(P>0.05, t=0.674), and 0.565±0.03 pmol/106 cells at 2' of stimulation with 60 mM KCl

(P>0.05, t=0.284). Continuous stimulation of the PC12 cells for 10 minutes and 20 minutes led to reduced levels of DA. Thus, DA overflow was 3.673±0.028 pmol/106 cells at 5', 3.510±0.175 pmol/106 cells at 10', and 3.657±0.431 pmol/106 cells at 20' of stimulation, which was significantly lower than the overflow of DA at 1' of stimulation reaching 6.087±0.658 pmol/106 cells (P<0.05, one-way ANOVA, followed by

Bonferroni's posttest). Likewise, in BoNT/A-treated cells the overflow of β-NAD, AMP, and total purines at 10' and 20' was significantly lower than the spontaneous overflow in pre-stimulation samples (Fig. 2.3 E,F,H).

2.4.3 Secretion evoked by nicotine 100 μM

Similar to the effects of high-K+ solution, 100 μM nicotine transiently increased the secretion of both ATP and β-NAD (Fig. 2.4A and D). In some cases secretion of purines at 5–20 min was significantly lower than secretion at 1 min of stimulation, which was not significantly different from pre-stimulation values. DA secretion was increased from 0.396 ± 0.058 pmol/106 cells before nicotine to 0.583 ± 0.060 pmol/106 cells at

1 min (P > 0.05, t = 1.239), 0.858 ± 0.16 pmol/106 cells at 2 min (P < 0.05, t = 3.055,

F = 4.116) and 0.82 ± 0.114 pmol/106 cells at 10 min (P > 0.05, t = 2.823) of stimulation with nicotine; one-way anova, followed by Bonferroni’s test, n = 8. Secretion of ATP was increased from 0.097 ± 0.023 pmol/106 cells at 0 min to 0.649 ± 0.246 pmol/106 cells at 1 min of stimulation (P < 0.05, t = 3.347, F = 3.188, one-way anova, followed by

Bonferroni’s test). Secretion of ATP at 2, 5, 10 and 20 min was not significantly different from secretion of ATP before stimulation (P > 0.05). Likewise, secretion of β-NAD was 87 increased from 0.552 ± 0.074 pmol/106 cells at 0 min to 1.629 ± 0.471 pmol/106 cells at

1 min (P < 0.05, t = 3.187, F = 3.411), whereas no significant increase was observed at

2–20 min of stimulation with nicotine (P > 0.05); one-way anova, followed by

Bonferroni’s test. No significant increase in ADP, AMP, ADO or total purines was observed at 1–20 min of stimulation with nicotine (P > 0.05; Fig. 2.4). In general, the results with KCl 60 mM simulation and nicotine stimulation were qualitatively similar, although under these experimental conditions the effect of nicotine seemed more transient than the effect of KCl.

2.4.4 Distribution of β-NAD, ADPR, cADPR in cell superfusates determined by HPLC fraction analysis

β-NAD, ADPR and cADPR co-elute as 1,N6-etheno-ADPR at ∼11.2 min due to conversion of β-NAD and cADPR to ADPR during the etheno-derivatization at 80°C

(Smyth et al., 2004; Breen et al., 2006). HPLC fraction analysis of the 11.2-min peak showed that β-NAD is the predominant compound, comprising 83.56 ± 2.47% of purines in the samples collected before stimulation (KCl 5 mM) (Fig. 2.5C) and 95.72 ± 2.9% during stimulation (KCl 60 mM); n = 4, P = 0.02, t = 3.143, unpaired t-test, two-tailed.

ADPR comprised 16.34 ± 2.47 and 4.277 ± 2.93% of the mixture before and during stimulation, respectively (n = 4, P = 0.02, t6 = 3.141, unpaired t-test, two-tailed). Thus, during stimulation with 60 mM KCl the release of β-NAD + ADPR + cADPR was increased (Fig. 2.3A and F) mainly due to increased secretion of β-NAD and not of

ADPR or cADPR (Fig. 2.5B and C, KCl 60 mM). It is reasonable, therefore, to refer to the compound released during stimulation with high-K+ solution as β-NAD. 88

2.4.5 Fractionation of synaptic vesicles by glycerol gradient: neural markers, content of

DA and purines, HPLC fraction analysis of β-NAD, ADPR, and cADPR

Aliquots of samples collected from glycerol gradient centrifugation were processed for Western immunoblot analysis of synaptophysin and secretogranin II.

Aliquots of the samples were also processed for analysis of purine and catecholamine contents. Synaptophysin, an SV marker, labeled vesicle population in the fractions containing ∼12–25% glycerol (Fig. 2.6A, top row, F11–F20), whereas the immunoreactive bands of secretogranin II, an LDCV marker, were negligible in all glycerol fractions (Fig. 2.6A, bottom row). Synaptophysin-like immunoreactivity was also present in F1, and this may be due to the presence of synaptophysin in membranes of very small low-density vesicles such as small synaptic-like microvesicles (SLMV), which fractionate at the top of the gradient. Parallel HPLC measurements of content of neurotransmitter substances in the glycerol fractions showed that all fractions contained

DA and purines, but in different proportions. Fractions with low glycerol (< 6%) concentrations (e.g., F1–F6) contained the largest amounts of ATP and β-NAD +

ADPR + cADPR. Then the concentrations of ATP and β-NAD + cADPR + ADPR gradually declined as the glycerol concentrations rose from ∼6 to 20% (F6–F14) and increased again at F17–F19 (∼22–25% glycerol). A similar pattern of distribution was observed with ATP, ADP, AMP and ADO (data not shown). An HPLC fraction analysis of the peak representing β-NAD + ADPR + cADPR in selected samples from the glycerol gradient samples showed that β-NAD is the prevailing compound in all fractions. Thus,

F3 comprised of 99.49% β-NAD, 0.36% ADPR and 0.15% cADPR. Likewise, F9 was 89 composed of 99.40% β-NAD, 0.32% ADPR and 0.29% cADPR. F13–F20 contained 92%

β-NAD, 8% ADPR and no detectable amounts of cADPR. DA was not detected in the low-glycerol fractions (F1–F5, Fig. 2.6D). However, DA was detected in fractions with higher glycerol levels and peaked in F15–F19.

2.4.6 Fractionation of synaptic vesicles by sucrose gradient: neural markers, content of

DA and purines, HPLC fraction analysis of β-NAD, ADPR, and cADPR

Aliquots of samples collected from sucrose gradient centrifugation (presumably containing LDCVs) were also processed for Western immunoblot analysis of synaptophysin and secretogranin II expression and for HPLC analysis of contents of purines and catecholamines. Synaptophysin labeled the fractions containing 0.4–1.0 M sucrose (F12–F18), whereas secretogranin II labeled mainly the fractions with 0.9–1.4 M sucrose (F16–F21; Fig. 2.7A). Similar to glycerol gradient fractions, F1 showed high expression of synaptophysin, possibly due to greater amounts of microvesicles in this fraction. Low-sucrose fractions (F1–F8) showed similar amounts of ATP (Fig. 2.7B).

Likewise, the content of β-NAD + cADPR + ADPR was similar in F1–F8 fractions

(Fig. 2.7C). Fractions F9–F15 showed declining concentrations of ATP and β-

NAD + cADPR + ADPR, whereas the content of ATP and β-NAD + cADPR + ADPR increased again in the high-sucrose fractions and peaked in F17–F20, the fractions with highest expression of secretogranin II. HPLC fraction analysis showed that the distribution of β-NAD, ADPR and cADPR in F3 was 100% β-NAD and no ADPR or cADPR. F9 contained 94.7% β-NAD, 0% ADPR and 5.3% cADPR, whereas F11–F16 contained 98.2% β-NAD and 1.8% ADPR. F19 and F20 contained 94.7% β-NAD, 4.4% 90

ADPR and 0.9% cADPR. Therefore, β-NAD was the major compound in all examined fractions. DA was almost absent in the low-sucrose fractions (F1–F9, Fig. 2.7D).

However, DA was detected in fractions with higher sucrose levels and peaked in F16–

F20.

2.4.7 Western immunoblot analysis of SNAP-25 in glycerol and sucrose gradient centrifugation fractions

Figure 2.8 shows the presence of SNAP-25 in all fractions with increasing concentrations of glycerol (6–25%) and sucrose (0.4–1.4 M). It appears that the SNAP-25 bands are stronger in the sucrose gradient fractions (compare Fig. 2.8 A and B) taking into account that equal fraction volumes were loaded in the immunoblots. 91

Fig 2.1

Figure 2.1. Protocol for isolation of SSVs and LDCVs from cultured NGF-differentiated rat PC12 cells Different velocities of cold ultracentrifugation and glycerol and sucrose gradient purification were used to obtain fractions enriched in SSVs or LDCVs. RT, room temperature; WB, Western blot analysis; HPLC, high performance liquid chromatography; LDCV, large debse-cord-like vesicles; SSV, small synaptic-like vesicles.

92

Fig 2.2

93

Figure 2.2. Spontaneous and high-K+-evoked secretion of DA in NGF-treated PC12 cells (A) PC12 cells secrete DA in a solution containing 5 mM KCl. Superfusion of the cells with 60 mM KCl evokes additional secretion of DA, which is maximum at 2 minutes of superfusion. (B) Pretreatment of cells with BoNT/A (30 nM for 16 h) reduces the evoked release of DA. Note also that DA amounts in superfusates are significantly reduced at 5–20' below the initial levels in non- stimulated cells. (C) Quantified data from 4–5 experiments expressed as the mean ± SEM. *P<0.05 vs. controls at 0' (KCl 5 mM), *P<0.05 vs. the highest value in BoNT/A group at 1'. One- way ANOVA followed by Bonferroni's multiple comparison tests. (D) Immunoblot analysis of SNAP-25 shows a single band at 25 kDa in homogenates from control PC12 cells. An additional 24-kDa band appears in BoNT/A-treated cells indicating cleavage of SNAP-25 induced by BoNT/A.

94

Fig 2.3

95

Figure 2.3. Spontaneous and high-K+-evoked release of adenine purines in NGF-treated PC12 cells (A) Original chromatograms from samples collected during perfusion of the PC12 cells with either 5 mM KCl (pre-stimulation sample) or in the presence of 60 mM KCl for different time periods. KCl 60 mM evoked overflow of ATP, ADP, AMP, adenosine (ADO) and a mixture of β- NAD, cADPR, and ADPR. Inset: chromatographic peaks generated by etheno-derivatized purine standards (1 pmol each). Note that β-NAD has significantly lower fluorescence coefficient in generating eADPR than ATP, ADP, ADPR, AMP and ADO in generating corresponding e- purines. (B) Excerpts of original chromatograms of superfusates from BoNT/A-pretreated cells with an emphasis on ATP and the mixture of β-NAD, ADPR, and cADPR. (C) Quantified data from 4–5 experiments expressed as the mean ± SEM. Data are averaged for each individual purine as well as for the sum of all purines (total purines). *P<0.05 vs. controls at 0' (KCl 5 mM). One-way ANOVA followed by Bonferroni's multiple comparison tests. #P<0.05 vs. 0' in controls. Unpaired Student's t-test. BoNT/A increased the spontaneous release of ADP, AMP, and total purines. The evoked release of most purines, except for ATP, was significantly reduced by BoNT/A. In BoNT/A-treated cells secretion of AMP, β-NAD+ADPR+cADPR, and total purines at 5–20' was below the initial purine levels.

96

Fig 2.4

Figure 2.4. Spontaneous and nicotine (100 μM)-evoked release of adenine purines in NGF- treated PC12 cells Quantified data from 6–10 experiments expressed as the mean ± SEM. Data are averaged for each individual purine as well as for the sum of all purines (total purines). *P<0.05 vs. controls at 0' (KCl 5 mM). *P<0.05 vs. 1'. One-way ANOVA followed by Bonferroni's multiple comparison test. Note that nicotine induced a transient secretion of purines, which was statistically significant for ATP and β-NAD+ADPR+cADPR. Although secretion of ADO and total purines did not show statistically significant increase at 1', the values of these purines at 5–20' of stimulation showed significantly lower levels than at 1'.

97

Fig 2.5

98

Figure 2.5. HPLC fraction analysis of the mixture of β-NAD, ADPR and cADPR (A) Original chromatograms of the 7.2-min (cADPR-containing) fraction, the 8.5-min (ADPR- containing) fraction and the 10.5-min (β-NAD containing) fraction of cell superfusate samples collected in the first 5 min of superfusion with 60 mM KCl and etheno-derivatized with 2- chloroacetaldehyde at 80°C, pH 4.0 for 40 min. (B) Averaged data. The pre-stimulation sample (5 mM KCl) contained mainly β-NAD and small amounts of ADPR, but no detectable amounts of cADPR. Stimulation with 60 mM KCl caused increase in the β-NAD content in the cell superfusate. (C) Distribution of β-NAD, ADPR and cADPR in pre-stimulation (5 mM KCl) samples and in samples collected during stimulation with 60 mM KCl. In both samples β-NAD is the primary purine nucleotide in the 11.2-min fraction. The ratio β-NAD:ADPR increases in the samples collected during stimulation with a high-K+ solution. *P<0.05 vs. KCl 5 mM, unpaired t- test, two-tailed.

99

Fig 2.6

100

Figure 2.6. Fraction separation of small synaptic vesicles (SSV) by glycerol gradient centrifugation (A) Fractionation of synaptic vesicles was evaluated by the immunoreactivity of the vesicular markers synaptophysin (for SSV and SLMV) and secretogranin II (for LDCV) using equal fraction volumes. Synaptophysin-immunoreactive SSVs (top row) were present mostly in fractions with 10–23% glycerol, F13–F20. Secretogranin II immunoreactivity was faint (bottom row) or absent, indicating lack of LDCV in the glycerol fractions. (B, C, D) Contents of ATP, β- NAD+ADPR+cADPR, and DA in fractions separated by glycerol gradient centrifugation. Averaged data (means ± SEM), n=4. (E) Concentrations of glycerol in fractions isolated by glycerol gradient centrifugation. Averaged data (means ± SEM), n=4.

101

Fig 2.7

102

Figure 2.7. Fraction separation of large dense-core vesicles (LDCVs) by sucrose gradient centrifugation (A) Fractionation of LDCV was evaluated by the immunoreactivity of the vesicular markers synaptophysin (top row) and secretogranin II (bottom row) using equal fraction volumes. Synaptophysin focused in fractions (F13–18) with 0.7–1.3 M sucrose (E). Secretogranin II- labeled LDCVs in fractions with sucrose concentrations above 0.9M (F15–21), A and E. (B,C,D) Contents of ATP, β-NAD+ADPR+cADPR, and DA in fractions separated by sucrose gradient centrifugation. Averaged data (means ± SEM), n=3. (E) Concentrations of sucrose in fractions isolated by sucrose gradient centrifugation. Averaged data (means ± SEM), n=3.

103

Fig 2.8

Figure 2.8. Western immunoblot analysis of SNAP-25 in fractions isolated by glycerol and sucrose gradient centrifugation Expression of SNAP-25 was analyzed by immunoblotting using equal fraction volumes. (A) SNAP-25 was expressed in fractions containing >5% glycerol and peaked in fractions containing 10–25% glycerol. (B) SNAP-25 was also expressed in fractions containing >0.32 M sucrose and peaked in fractions containing 0.7–1.12 M sucrose.

104

2.5 Discussion

Studies in rat tail artery suggest that ATP and NA are probably stored in the same vesicles within the sympathetic nerve varicosity and are released in parallel by vesicular exocytosis (Stjarne, 1989; Stjarne et al., 1994; Brock & Cunnane, 1999). Likewise, ATP and catecholamines appear released in parallel in adrenal chromaffin cells (Wagner,

1985; Njus et al., 1986; Todorov et al., 1996) and hence are probably released from the same vesicles. In contrast, functional and overflow studies in the rodent vas deferens have suggested that ATP and NA are released differentially (e.g., Ellis & Burnstock,

1990; Trachte, 1985; Ellis & Burnstock, 1989; Todorov et al., 1996) and thus may originate from different synaptic vesicles. These and other studies illustrate decades-long controversy about the sources of storage and release of cotransmitter substances.

Understanding mechanisms of co-storage and co-release of neurotransmitters is important for understanding mechanisms of synaptic integration and plasticity. It may raise the potential for selective external control of synaptic events and may thus have potential clinical significance. This becomes particularly important when a novel neurotransmitter substance is discovered.

Work in our laboratory has contributed to the recent notion that, in addition to its well-known role in intracellular processes, β-NAD is also an extracellular molecule involved in cell-to-cell communications (Ziegler & Niere, 2004; Billington et al., 2006) and may function as a novel neuromodulator (Smyth et al., 2004; Breen et al., 2006) and a novel neurotransmitter (Mutafova-Yambolieva et al., 2007). β-NAD is released concomitantly with NA and ATP in blood vessels, urinary bladder, vas deferens, large 105 intestine (Smyth et al., 2004; Smyth et al., 2006a; Smyth et al., 2006b; Smyth et al.,

2009; Bobalova & Mutafova-Yambolieva, 2006; Mutafova-Yambolieva et al., 2007;

Breen et al., 2006) and brain (V. Mutafova-Yambolieva, I. A. Yamboliev, L. Durnin and

P. Dai). In a number of smooth muscles the release of NA and β-NAD is regulated in parallel (i.e., Smyth et al., 2009; Bobalova & Mutafova-Yambolieva, 2006), suggesting that the two substances may originate from the same synaptic vesicles. The release of

ATP and β-NAD, however, often appear differentially regulated (i.e., Smyth et al., 2009;

Mutafova-Yambolieva et al., 2007). Therefore, ATP and β-NAD may originate from different release sites including different nerves or different secretory vesicles. The present study provides a thorough analysis of simultaneous release of catecholamines and purines in NGF-differentiated PC12 cells. The major novel finding is that differentiated

PC12 cells exhibit constitutive and evoked release not only of DA and ATP but also of β-

NAD. This represents a significant advance in knowledge over previously published studies limited to secretion of catecholamines and ATP in chromaffin cells. β-NAD, ATP and DA appear to be co-stored in multiple vesicles with no well-defined distinction in the primary distribution of these substances. Nonetheless, β-NAD, ATP and DA appear to have different preferential sites of secretion.

We employed a small-volume superfusion assay for collecting neurotransmitters secreted at rest and upon stimulation with high-K+ solution or nicotine from NGF- differentiated rat PC12 cells. This technique allowed examination of the time course of neurotransmitter release under constant flow. NGF-treated PC12 cells showed constitutive release of DA, ATP, ADP, AMP, ADO, β-NAD and ADPR. Stimulation of cells with high-K+ solution (and hence causing Ca2+ influx due to membrane 106 depolarization) elicited release of DA and the purines listed above. Importantly, the maximum evoked release of all substances was achieved within the first 2 min of stimulation and returned to basal levels by the fifth minute of stimulation even though the stimulation lasted for 20 min. Thus, studies assaying mass release of neurotransmitters in

PC12 cells during stimulation with high-K+ stimulation for > 5 min may underestimate the evoked release of neurotransmitters at the expense of measuring basal release and more degradation products. As mentioned above, differentiated PC12 cells exhibited constitutive and evoked release of β-NAD. Small amounts of ADPR were also found in tissue superfusates. ADPR is the major direct metabolite of β-NAD, as 98% of β-NAD is degraded to ADPR by NAD glycohydrolase and ∼2% of β-NAD is degraded to cADPR by ADP-ribosyl cyclase (Lee, 2001). In samples collected in normal-K+ solution the β-

NAD:ADPR ratio was 6 : 1, whereas the high-K+ (60 mM)-evoked release of β-

NAD:ADPR was 70 : 1. Activation of nicotinic acetylcholine receptors with nicotine also caused a transient increase in secretion of DA, ATP and β-NAD, as did the high K+- solution.

SLMVs are present in PC12 cells (Liu & Edwards, 1997; Varoqui & Erickson,

1998) and the biogenesis and composition of SLMV is similar to the SSVs in presynaptic nerves in the central and peripheral nervous systems (Navone et al., 1989; Llona, 1995).

Synaptophysin is a component of the SLMV and SSV membrane (Navone et al., 1986;

Cutler & Cramer, 1990; Jahn & Sudhof, 1994; Llona, 1995) and to a smaller extent is also present in the LDCV membranes (Marxen et al., 1997). To isolate SSVs and

LDCVs, after removing nuclei, mitochondria, lysosomes, proxyosomes and other large particles, we loaded the samples onto continuous glycerol and sucrose gradients, 107 respectively. In agreement with previous studies (Melikian & Buckley, 1999; Liu &

Edwards, 1997; Varoqui & Erickson, 1998) synaptophysin labeled the SSV population in the fractions containing 10–23% glycerol. As shown earlier (Marxen et al., 1997), synaptophysin was present in fractions containing 0.7–1.3 M sucrose, presumably rich in

LDCVs. Synaptophysin was also present in membranes of SLMVs, which fractionated at the top (F1) of the two gradients. To label LDCVs we used secretogranin II (Cutler &

Cramer, 1990; Fischer-Colbrie et al., 1995). As anticipated, secretogranin II fractionated through the sucrose gradient with maximum occurrence in fractions with sucrose concentrations > 0.9 M. These observations are also in agreement with previous findings

(e.g., Yao & Hersh, 2007; Liu & Edwards, 1997; Melikian & Buckley, 1999).

Secretogranin II did not label the glycerol fractions well, suggesting that no major pool of

LDCVs was present in these samples. We concluded, therefore, that the glycerol and sucrose fractionation centrifugations led to isolation of fractions rich in SSVs–SLMVs and in LDCVs, respectively.

We next determined the content of purines and catecholamines in SSVs–SLMVs and in LDCVs isolated as described above. Previous studies have shown that ATP and catecholamines are stored in and concomitantly released from large secretory vesicles in chromaffin cells and PC12 cells (Wagner, 1985; Kasai et al., 2001), although these studies did not examine the role of small synaptic vesicles in the release of ATP and DA.

In the present study we also found that ATP and DA are co-stored in LDCVs: the high- sucrose fractions that labeled for secretogranin II showed high amounts of ATP, β-NAD and DA. β-NAD comprised > 95% of the mixture β-NAD + ADPR + cADPR in the high- sucrose fractions. Low-sucrose fractions, presumably representing the cytosol, contained 108 large amounts of ATP and β-NAD, but almost no DA. Likewise, fractions with low glycerol concentrations contained the largest amounts of ATP and β-NAD, but no DA.

Based on immunoblot analysis of the distribution of synaptophysin and secretogranin II we assume that these glycerol-free fractions largely represented the cytosol. Thus, a fraction of the neurotransmitter substances detected in F1–F6 may be the neurotransmitters synthesized in the cytoplasm ‘waiting’ to be packaged into synaptic vesicles. Another fraction may comprise neurotransmitters already transported into

SLMVs. The content of β-NAD was greater than that of ATP (and DA) in the glycerol- free fractions. Moderate increase in glycerol concentrations led to isolation of fractions containing low concentrations of ATP and β-NAD. However, the concentrations of ATP and β-NAD increased again in the fractions with highest glycerol concentrations, presumably containing SSVs. In all fractions β-NAD represented the major compound of the β-NAD + ADPR + cADPR mixture, comprising 92–99% of the mixture. The high- glycerol fractions also contained DA. Therefore, ATP, DA and β-NAD are probably stored not only in LDCVs but also in SSVs.

Neuronal release of noradrenaline and ATP is generally believed to occur via exocytosis. The release of β-NAD evoked by electrical field stimulation is severely disrupted by SNAP-25 cleavage in mesenteric blood vessels and large intestine and hence, in these systems, β-NAD appears released primarily by vesicular exocytosis

(Smyth et al., 2006b; Mutafova-Yambolieva et al., 2007). We next tested whether, in differentiated PC12 cells, β-NAD, ATP and DA are secreted by SNAP-25-mediated mechanisms. SNAP-25 is present in synaptic membranes and transport vesicles and is involved in the docking and fusion of synaptic vesicles (Sollner et al., 1993). 109

Biochemical and cellular fractionation studies have suggested that SNAP-25 is a part of the SNARE complex, which represents a critical step in synaptic exocytosis (Sollner et al., 1993; McMahon & Sudhof, 1995). In support of this, in the present study SNAP-25 showed parallel distribution in the fractions from glycerol and sucrose centrifugation gradients. Interestingly, samples collected before stimulation of PC12 cells treated with

BoNT/A contained higher levels of DA and purines than samples collected before stimulation of non-treated cells. Therefore, spontaneous release of neurotransmitters might be a regulated process that requires intact SNAP-25. Continuous stimulation with high-K+ solution caused a progressive decrease in the levels of secreted catecholamines and purines in the BoNT/A-treated cells, suggesting that disruption of SNAP-25 may lead to depletion of pool(s) of secretory vesicles that are available for release upon membrane depolarization and Ca2+ influx. This is in accordance with a previous study showing that in hippocampal slice cultures 5–10 min high-K+ stimulation causes progressive depletion of synaptic vesicles and redistribution of SNAP-25 from the plasma membrane to membrane-bound tubulovesicular structures in the (Tao-Cheng et al., 2000). It should be pointed out that the high-K+-evoked release of ATP remained unchanged by

BoNT/A, whereas the evoked release of β-NAD and DA was abolished. Therefore, despite similar distribution of ATP, β-NAD and DA in SSVs and LDCVs, there may be differences in the preferential sites of evoked secretion of neurotransmitter substances in the NGF-differentiated PC12 cells: the predominant release of β-NAD and DA appears to be from SNAP-25-associated sources, whereas the release of ATP seems to occur predominantly from sources not associated with SNAP-25 but still dependent on membrane depolarization. As some studies show, ATP can also be released through 110 native or overexpressed hemichannels (Ebihara, 2003; Belliveau et al., 2006; Huang et al., 2007) or secretion of ATP can occur independently of vesicular exocytosis (Hussl et al., 2007). However, these mechanisms cannot explain the results of the present study as these mechanisms usually occur independently of membrane depolarization. Further studies are warranted to determine the exact sources of differential secretion of β-NAD,

DA and ATP in these cells.

In summary, we demonstrate here that β-NAD, a putative neurotransmitter and a neuromodulator, is subject to constitutive and regulated release in NGF-differentiated

PC12 cells. β-NAD, ATP and DA are present in both SSV and LDCV populations of secretory vesicles with no definite distinction in the primary distribution of the three substances. SNAP-25-mediated mechanisms may be involved in the constitutive release of catecholamines and purines. Membrane depolarization-evoked release of DA, β-NAD and ATP is differentially affected by disruption of SNAP-25, suggesting that the three neurotransmitter substances may have different preferential sites of release, including sites associated with SNAP-25 (i.e., DA and β-NAD) and sites not associated with

SNAP-25 (i.e., ATP).

111

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Chapter 3

β-nicotinamide adenine dinucleotide (β-NAD) is an enteric inhibitory neurotransmitter in human and non-human primate colons

Sung Jin Hwang*, Leonie Durnin*, Laura Dwyer, Poong-Lyul Rhee, Sean M. Ward, Sang Don Koh, Kenton M. Sanders and Violeta N. Mutafova- Yambolieva Published in Gastroenterology 2011 Feb;140(2):608-617.e6.

*These authors contributed equally to this work

117

3.1 Abstract

Background & Aims: An important component of enteric inhibitory neurotransmission is mediated by a purine neurotransmitter, such as adenosine 5’- triphosphate (ATP), binding to P2Y1 receptors and activating small conductance K+ channels. In murine colon β-nicotinamide adenine dinucleotide (β-NAD) is released with

ATP and mimics the pharmacology of inhibitory neurotransmission better than ATP.

Here β-NAD and ATP were compared as possible inhibitory neurotransmitters in human and monkey colons. Methods: A small-volume superfusion assay and HPLC with fluorescence detection were used to evaluate spontaneous and nerve-evoked overflow of

β-NAD, ATP and metabolites. Postjunctional responses to nerve stimulation, β-NAD and

ATP were compared using intracellular membrane potential and force measurements.

Effects of β-NAD on smooth muscle cells (SMCs) were recorded by patch clamp. P2Y receptor transcripts and proteins were assayed by RT-PCR. Results: In contrast to ATP, overflow of β-NAD evoked by electrical field stimulation correlated with stimulation frequency and was diminished by neurotoxins, tetrodotoxin and ω-conotoxin GVIA.

Inhibitory junction potentials and responses to exogenous β-NAD, but not ATP, were blocked by P2Y receptor antagonists suramin, PPADS, MRS2179 and MRS2500. β-NAD activated non-selective cation currents in SMCs, but failed to activate outward currents.

Conclusions: β-NAD meets the criteria for a neurotransmitter better than ATP in human and monkey colons and therefore may contribute to neural regulation of colonic motility.

SMCs are unlikely targets for inhibitory purine neurotransmitters because dominant responses of SMCs were activation of net inward, rather than outward, current. 118

3.2 Introduction

Enteric neural control is essential for mixing and propulsion of luminal contents in the gastrointestinal (GI) tract (Kunze & Furness, 1999). Motility disorders, including diabetic enteropathy, slow-transit constipation, inflammatory neuropathy, and ageing- associated motility disorders are linked, directly or indirectly, to defects in the neural control of the GI tract (Bassotti & Villanacci, 2006; Camilleri et al., 2008; Lomax et al.,

2010). However, underlying mechanisms for neurotransmitter actions and post-junctional targets for neurotransmitters are not entirely understood. Thus, accurate design and targeting of treatments for motility disorders has proven difficult.

An important component of enteric inhibitory neurotransmission is mediated by a purine neurotransmitter binding to P2Y1 purinoreceptors (Gallego et al., 2008a; Grasa et al., 2009; Wood, 2006). Functional studies have typically suggested that adenosine 5’- triphosphate (ATP) is the inhibitory purinergic neurotransmitter in the gut (Burnstock,

2008b). However, previous studies have largely assayed integrated responses in complex muscular tissues that are possibly mediated by mixed purine receptors. Few studies have measured release of neurotransmitters directly. ATP does not mimic the effects of the endogenous neurotransmitter (Mutafova-Yambolieva et al., 2007) in some cases, and many investigators refer to the neurotransmitter as a purine or “purine-like” substance

(Auli et al., 2008; De Man et al., 2003; Gallego et al., 2006; Gallego et al., 2008b; Serio et al., 2003).

β-nicotinamide adenine dinucleotide (β-NAD) is another purine substance released by nerve stimulation in vascular and visceral smooth muscles and neuro- 119 secretory cells (Mutafova-Yambolieva et al., 2007; Smyth et al., 2004; Breen et al.,

2006; Smyth et al., 2006b; Yamboliev et al., 2009). We found that nerve stimulation of murine colon evoked release of β-NAD and ATP, and the pharmacological profile of β-

NAD mimicked the endogenous neurotransmitter better than ATP (Mutafova-

Yambolieva et al., 2007). Here we investigated whether β-NAD serves as an inhibitory neurotransmitter in larger mammals, such as humans and non-human primates.

3.3 Methods

3.3.1 Tissue Preparation

Proximal colons of 94 Cynomolgus monkeys (Macaca fascicularis) of both sexes were obtained from Charles River Laboratories Preclinical Services (Reno, NV).

Monkeys sedated with Ketamine (10 mg kg−1) and 0.7 ml Beuthanasia-D (Schering-

Plough AH, Kenilworth, NJ, USA) were exsanguinated (Charles River Laboratories

Institutional Animal Care and Use Committee).

Human colon samples were obtained as surgical waste from 48 male and female patients (ages 32-77) undergoing colon resections for neoplasm or diverticulitis at

Renown Medical Center and St. Mary’s Hospital (Reno, NV), or Samsung Hospital

(Seoul, South Korea). Tissues from a site distant to the disease-affected area were placed in oxygenated Krebs solution (10°C) containing (mmol/L): 118.5 NaCl; 4.2 KCl; 1.2

MgCl2; 23.8 NaHCO3; 1.2 KH2PO4; 11.0 dextrose; 1.8 CaCl2 (pH 7.4). Use of human colon was approved by Human Subjects Research Committees at Renown and St.

Mary’s, and Samsung hospitals, and the Biomedical Institutional Review Board at 120

University of Nevada, Reno. Tunica muscularis was processed as below after removal of mucosa and submucosa.

3.3.2 Purine overflow

Muscle strips (2 × 6 mm) were prepared from monkey and human colonic tunica muscularis. Pure circular muscles (CM) (2 × 6 mm) of monkey colon were prepared by peeling off longitudinal muscle (LM) with attached myenteric ganglia (MG). NADPH- diaphorase histochemistry (below) verified absence of MG in CM. Due to adhesions between smooth muscle layers in human colon, CM preparations were not viable.

Muscles (50-70 mg) were placed in 200-μl superfusion chambers equipped with platinum electrodes (Mutafova-Yambolieva et al., 2007; Bobalova & Mutafova-

Yambolieva, 2001). Nerves were stimulated by electrical field stimulation (EFS; 480 pulses; 0.3 ms (whole muscle, WM) or 0.5 ms (CM); 4 Hz or 16 Hz; supramaximal voltage). Superfusates were collected before and during EFS and processed for derivatization to 1,N6-etheno-purines (Levitt et al., 1984; Bobalova et al., 2002). ATP,

ADP, AMP and adenosine (ADO) were converted to their 1,N6-etheno-derivatives eATP, eADP, eAMP, and eADO, respectively (Bobalova et al., 2002). β-NAD, ADPR and cADPR all derivatized to eADPR (Smyth et al., 2004).

3.3.3 HPLC Assay of Etheno-purines

Reverse-phase gradient HP1100 liquid chromatography equipped with fluorescence detector (Agilent Technologies, Wilmington, DE) was used as previously described (Bobalova et al., 2002; Smyth et al., 2004). Amounts of 121 nucleotides/nucleosides were calculated against standards. Results, normalized for sample volume and tissue weight, were expressed as femtomoles per milligram of tissue wet weight (fmol/mg).

3.3.4 HPLC Fraction Analysis

The compounds forming eADPR were determined by HPLC fraction analysis

(Mutafova-Yambolieva et al., 2007; Smyth et al., 2004). Superfusates from 28 chambers were combined, concentrated and analyzed by HPLC. Samples were collected according to retention time (cADPR and ATP 7.0–7.4 min, ADPR 8.3–8.7 min, and β-NAD 10.3–

10.7 min) and were etheno-derivatization and reanalyzed by HPLC for eADPR.

3.3.5 Electrophysiology and Contractions

Muscles (10 × 10 mm) were perfused with oxygenated Krebs solution (mmol/L):

NaCl 118.5; KCl 4.5; MgCl2 1.2; NaHCO3 23.8; KH2PO4 1.2; dextrose 11.0; CaCl2 2.4, pH 7.4, at 37±0.5°C, and CM cells were impaled with glass microelectrodes in the presence of nifedipine (1 μmol/L) to reduce movement (Mutafova-Yambolieva et al.,

2007; Burns et al., 1996). Parallel platinum electrodes delivered EFS (0.5ms) from a

Grass S48 stimulator (Quincy, MA, USA).

Receptor antagonists were perfused into the recording chamber. ATP and β-NAD, were applied (10 ms pulses at 10 psi) via picospritzer micropipettes (Picospritzer;

General Valve, East Hanover, NJ) positioned close to electrical recording sites.

Single colonic smooth muscle cells (SMCs) were prepared from human and monkey colon as described (Kong et al., 2000). Membrane currents were recorded by 122 patch clamp (on-cell recording or permeabilized-patch conditions with amphotericin B)

(Kong et al., 2000). Currents were amplified (Axopatch 200B, Instruments) and digitized (12-bit A/D converter; Digidata 1322A, Axon Instruments) on-line via pCLAMP software (V9.2.0.09, Axon Instruments). Data were sampled at 2 kHz (whole cell) or 5 kHz (single channel recordings).

Bath solution (CaPSS) for whole-cell recording contained (mmol/L): 5 KCl, 135

NaCl, 2 CaCl2, 10 glucose, 1 MgCl2, and 10 HEPES, adjusted to pH 7.4 with Tris. Pipette solution contained (mmol/L): 110 Cs-aspartate, 30 TEACl, 0.1 EGTA, 10 HEPES, pH7.2. For single channel recordings, bath solution contained high K+ (HK, mmol/L):

150 KCl, EGTA 1, and 10 HEPES, pH 7.4. The pipette solution in single channel recordings was CaPSS.

Muscle strips (1 × 10 mm) were prepared for contractile studies and attached to

Fort 10 force transducers (World Precision Instruments, Sarasota, Fl) for force measurements (Mutafova-Yambolieva et al., 2007; Burns et al., 1996).

3.3.6 Expression of P2Y receptors

P2Y1, P2Y2, P2Y4, P2Y6 and P2Y11 receptor transcripts were measured in monkey and human colons. Detailed methods are given in Supplementary Materials

(Section 3.6) and Table S3.1.

3.3.7 NADPH Diaphorase Histochemistry

CMs were checked for MG using NADPH diaphorase histochemistry as described

(Ward et al., 1992; Young et al., 1992). After staining, muscles were mounted in 123

Aquamount (Lerner Laboratories, Pittsburgh, PA) and examined with a Zeiss Axiovert

200M and a Hamamatsu C5810 camera.

3.3.8 Data Analysis

Data presented are means ± SEM. In neurotransmitter overflow experiments means are compared by two-tailed, unpaired t tests (GraphPadPrism, GraphPad Software,

San Diego, CA). In intracellular electrical and mechanical experiments means are compared by two-tailed paired Student’s t tests and Mann Whitney rank sum tests. A probability of <.05 was considered significant. For analysis of the picospritizing data, membrane hyperpolarization area following picospritzing was plotted as a function of mV.ms-1 until the membrane repolarized to control level. For force measurements, relaxation responses (10 min) were calculated as percents of the maximal inhibition following application of β-NAD.

3.3.9 Drugs

ATP, ADP, AMP, adenosine, β-NAD, nifedipine, PPADS, suramin, apamin, L-

NNA, atropine, ω-conotoxin GVIA, and amphotericine B were purchased from Sigma-

Aldrich (St. Louis, MO). ADPR and cADPR came from Biolog (Germany). MRS2179 and MRS2500 came from Tocris Bioscience (Ellisville, MO). Nifedipine, dissolved in ethanol at 10mmol/L, was added to the perfusion to make 1μmol/L. Other drugs were dissolved in de-ionized H2O and diluted in perfusion solutions.

124

3.4 Results

3.4.1 Neural release of purines

Stimulation of intrinsic nerves caused accumulation of ATP and β-NAD and metabolites, ADP, AMP, ADO, ADPR and cADPR in tissue superfusates (Figs. 3.1 and

3.2, Tables S3.2 and S3.3 in Supplementary Materials). ADP is a product of ATP. AMP and ADO are products of ATP and β-NAD, and cADPR and ADP-ribose are products of

β-NAD (Zimmermann, 1996; Graeff et al., 1998b; Lee, 2001). Therefore, ATP and β-

NAD detected in superfusates are remnants of purines released less metabolic products.

Fig. 3.1 shows overflow of purines from human colonic muscles. EFS evoked β-NAD release at 4 Hz (Fig. 3.1A, C) and release increased at 16 Hz (Fig. 3.1C, P=.0227). In contrast, ATP did not increase significantly with stimulation frequency (Fig. 3.1B,

P=.3325). β-NAD was released from nerves, as it was reduced by 0.5 μmol/L TTX

(P=.0028), a blocker of neural Na+ channels, and 50 nmol/L ω-conotoxin GVIA

(P=.0469), an inhibitor of neuronal voltage-dependent, N-type Ca2+ channels (Fig. 3.1C).

ATP release was not significantly reduced by TTX (P=.2360) or ω-conotoxin GVIA

(P=.2994), Fig. 3.1B, suggesting that most ATP released during EFS may occur by TTX- insensitive mechanisms. EFS-evoked overflow of ADP, AMP, and ADO was not significantly reduced by TTX and ω-conotoxin GVIA (P>.05), but total purine release followed β-NAD overflow (Table S3.2, Supplementary Material).

In human colon β-NAD, ADPR and cADPR contributed 82%, 17.4% and 0.6% of the eADPR peak, respectively (Fig. 3.1D). Therefore, β-NAD was the dominant purine 125 nucleotide in β-NAD+ADPR+cADPR peaks. β-NAD release exceeded ATP by about 15 fold (P=.0072) (Fig.3.1E).

β-NAD released in intact muscles might originate from enteric ganglia rather than motor nerve terminals. Thus, we measured purine release from pure CMs with LM and

MG removed. Fig. 3.2 shows overflow of β-NAD and ATP and metabolites from monkey colonic WM (containing MG) and pure CM, with nerve processes but no cell bodies (Fig.

3.2A). ATP release was not increased between 4 Hz and 16 Hz (Fig. 3.2B, WM, P=.7831;

CM, P=.6706), and was not significantly reduced by TTX (WM, P=.2487; CM, P=.4649) or ω-conotoxin GVIA (WM, P=.1688; CM, P=.2371). In contrast β-NAD release increased with stimulus frequency (Fig. 3.2C, WM, P=.0398; CM, P=.0300). Release of

β-NAD from CM was 2- to 3-fold lower than from WM at 16 Hz (P=.0289), but no difference was observed at 4 Hz (P=.6117). Thus, at lower stimulation frequencies, β-

NAD may be released exclusively from nerve endings, but at 16 Hz, ß-NAD may also be released in ganglia. β-NAD release (16 Hz) was inhibited by TTX (WM P=.0374, CM

P=.0024) and ω-conotoxin GVIA (WM, P=.0092, CM, P=.0340), indicating that β-NAD was released from enteric motor nerves.

In monkey WM, β-NAD, ADPR and cADPR contributed ~64%, 33% and 3% of the eADPR peak, respectively (Fig. 3.2D). In CM, β-NAD, ADPR and cADPR contributed 86%, 12.5% and 1.5% of the eADPR peak, respectively. Therefore, β-NAD is the dominant nucleotide in β-NAD+ADPR+cADPR peaks. β-NAD exceeded the mass of

ATP by 25-fold in WM (P=.0094) and 15-fold in CM (P=.0043, Fig. 3.2E).

3.4.2 Purinergic component of inhibitory junction potentials (IJPs) 126

Functional responses of colonic muscles were tested by measuring membrane potentials and contractions (Supplementary materials; Figs. S3.1, S3.2). CM cells of monkey proximal colon and human colon had resting membrane potentials averaging -

51.2±1.1 mV (n=39) and -43.0±0.6 (n=41), respectively. The effects of antagonists, including suramin (100 μmol/L), PPADS (30 μmol/L), MRS2179 (10 μmol/L),

MRS2500 (1 μmol/L) and apamin (0.3 μmol/L) were tested on the purinergic component of IJPs (Fig. 3.3). Apamin reduced IJPs in monkey colon (by 70±3.9%; n=19, P<.00001), suramin (by 51±1.7%; P<.00005, n=10), PPADS (by 61±4.8%; P<.00001, n=15), and

MRS2179, selective P2Y1 antagonist (Camaioni et al., 1998; Hu et al., 2003) (by

85±6.4%; P<.00001, n=14), Fig. 3.3A. Apamin tended to reduce IJPs of human colon (by

16±4.7%; n=19; but did not reach significance; P>.05), Fig. 3.3B. Human IJPs were reduced by suramin (by 60±6.0%; P<.0001, n=5), PPADS (by 81±1.6%; P<.0005, n=5),

MRS2179 (by 83±2.1%; P<.00001, n=15), and MRS2500 (by 95±1.3%;

P<0.0000000001, n=10), Fig. 3.3B. Therefore, non-nitrergic inhibitory responses in human colon were mediated via P2Y1 receptors, as previously reported (Gallego et al.,

2006) and these responses have similar pharmacology in monkey colon.

Antagonists of purinergic inhibitory responses also inhibited responses to exogenous transmitters. With atropine (1 μmol/L) and L-NNA (100 μmol/L) present, picospritzing of ATP onto monkey colonic muscles caused hyperpolarization averaging

12±0.9 mV (n=21). Apamin (0.3 μmol/L) inhibited this response by 97±2.4% (n=5,

P<.05, Fig. 3.4A, left). Responses to exogenous ATP were not inhibited significantly by suramin (100 μmol/L, reduced by 5±5.2% of control, P>.05, n=5), PPADS (30 μmol/L, by 12±9.6%, P>.05, n=6), or MRS2179 (10 μmol/L, by 23±12%, n=5, P>.05) (Fig. 3.4B- 127

D, left). Picospritzing ATP onto human muscles under the same conditions caused hyperpolarization averaging 10.9±0.7 mV (n=11). Apamin did not reduce responses to

ATP significantly, and ATP responses were not reduced by MRS2179 (10 μmol/L, by

9±2.1%, n=5, P>.05, Fig. 3.4E, left) or MRS2500 (1 μmol/L, by 12.3±7.3%, n=3, P>.05).

Thus, IJPs and hyperpolarizations to ATP displayed different pharmacological profiles, suggesting responses to the endogenous neurotransmitter and exogenous ATP are mediated by different receptors.

Picospritzing β-NAD caused hyperpolarization of monkey colon averaging

7.9±0.8 mV (n=23). Responses to β-NAD were reduced by apamin (0.3 μmol/L) by

98±2.4% (n=6, P<.05), suramin (100 μmol/L) by 95±3.8% (n=5, P<.05), PPADS (30

μmol/L) by 88±3.2% (n=6, P<.05) and MRS2179 (10 μmol/L) by 98±1.3% (n=6,

P<.005), Fig. 3.4A-D, right. β-NAD hyperpolarized human colon by 6.8±0.5 mV (n=13).

β-NAD solutions are acidic, but picospritzing of acidified Krebs solution (pH 3.6) had no effect on resting membrane potential. Apamin did not affect the hyperpolarization responses to β-NAD (n=2). However, β-NAD responses were reduced by MRS2179 (10

μmol/L) by 68±6.5% (n=5, P<.05, Fig. 3.4E, right) and by MRS2500 (1 μmol/L) by

97±1.7% (n=3, P<.05, Fig. 3.4F, right). Thus, β-NAD mimicked the pharmacology of the purine inhibitory neurotransmitter.

Monkey and human colonic muscles expressed P2Y receptor transcripts, including P2Y1 receptors (Fig. 3.5).

3.4.3 Effects of β-NAD and ATP on SMC conductance 128

β-NAD induced hyperpolarization in human and monkey colonic muscles, which might be accomplished by activation of K+ channels or inhibition of a tonic inward current in colonic SMCs. The effects of β-NAD on isolated SMC were tested with cell- attached patch clamp recording. Monkey colonic SMC were held at -80 mV and depolarized by ramping potential to +80 mV. Control single channel openings at -80 mV were negligible, but β-NAD (1 mmol/L) increased channel openings (-53±17 pA, n=5,

P<.05, Fig. 3.6A). These effects returned to control level upon washout and were repeatable. Currents activated by β-NAD reversed at 0 mV (Fig. 3.6B), suggesting that

- + non-selective cation channels, but not Cl channels (ECl ≈ -30 mV) or K channels (EK ≈ -

80 mV) were responsible. ATP (1 mmol/L) on cell-attached patches also activated non- selective cation channels at -80 mV (n=2, Fig. 3.6C).

β-NAD was also tested on human colonic SMC using permeabilized patch, whole-cell recording. Contamination from K+ and Cl- currents were eliminated with Cs-

TEA pipette solution with ECl set to -40 mV. Under these conditions, β-NAD (5mmol/L) activated inward current, averaging -424±0.4 pA (-80mV; n=7, P<.01 control net inward current was -37±4 pA) (Fig. 3.6D). The currents activated reversed at about 0 mV (Fig.

3.6E). Therefore, β-NAD-activated currents were not due to K+ and Cl- conductances, but appeared to be due to activation of non-selective cation channels. Activation of non- selective cation channels would generate net inward current and depolarize SMC. ATP also activated non-selective cation current at -80 mV (-99±47pA, n=4, Fig. 3.6F). 129

Fig 3.1

130

Figure 3.1. Overflow of ATP and β-NAD in human colonic muscle. (A) Chromatograms of tissue superfusates collected during EFS (4 and 16 Hz) and with neural blockers at 16 Hz. EFS-evoked overflow of β-NAD, but not ATP, increased with stimulation frequency and decreased with TTX (0.5 μmol/L) or ω-conotoxin GVIA (50 nmol/L); LU, luminescence units. Note that each chromatogram shows data from a different experimental tissue. (B and C) Averaged data are means±SEM; (o) denote significant differences from 4 Hz controls (P<.05); (*) denote significant differences from 16 Hz controls (*P<.05, **P<.01). Experiment number shown in parentheses. (D and E) HPLC fraction analysis demonstrated that β-NAD is the primary purine nucleotide in β-NAD+ADPR+cADPR peak. (E) Fraction analysis showed that the amount of β-NAD exceeded ATP; (*) denote significant differences from ATP (**P<.01).

131

Fig. 3.2

132

Figure 3.2. ATP and β-NAD are released in monkey whole and circular muscle. (A) NADPH-diaphorase staining of LM and CM of monkey colon. Left panel shows NADPH- diaphorase+ neurons (arrows) in myenteric plexus LM preparation. Inset is magnification showing neurons within MG (*). Right panel shows that CM is free of ganglia. Nerve fibers parallel to CM remain (arrow heads). Inset is magnification showing nerve fibers. Scale bars are 500 μm in both panels and 50μm in insets. (B and C) ATP and β-NAD released from monkey WM and CM (4 and 16 Hz) and with TTX (0.5 μmol/L) and ω-conotoxin-GVIA (50 nmol/L). Averaged data (fmol/mg tissue) are means±SEM; (o) denote significant differences from 4 Hz controls (P<.05); (*) denote significant differences from 16 Hz controls (*P<.05, **P<0.01). (D and E) HPLC fraction analysis demonstrates that β-NAD is the primary nucleotide in β-NAD+ADPR+cADPR peak in WM and CM. β-NAD was greater in WM than CM. (o) denote significant difference from WM (P<.05). β-NAD exceeded ATP in WM and CM (**P<0.01).

133

Fig 3.3

Figure 3.3. Purinergic component of IJPs. IJPs generated by EFS (0.5 ms; 1 pulse; solid circles) in monkey (A) and human colonic muscles (B) in the presence of atropine (1 μmol/L), L-NNA (100 μmol/L) and nifedipine (1 μmol/L). Purinergic component of IJP in monkey colon was reduced by apamin, suramin, PPADS, and MRS2179. In human colon apamin only partially reduced IJPs, but suramin, PPADS, MRS2179 and MRS2500 significantly reduced IJPs.

134 Fig 3.4

135

Figure 3.4. Membrane responses to exogenous purines. Picospritzed ATP (10 mmol/L, left panels) and β-NAD (50 mmol/L, right panels) hyperpolarized monkey (A-D) and human muscles (E, F). Responses to β-NAD, but not to ATP, were inhibited by purine receptor antagonists (as labeled). In monkey muscles, apamin inhibited both ATP and β-NAD induced hyperpolarization (A, left and right panels). In some muscles β-NAD caused transient depolarization after MRS2179 (D, right). In human muscles MRS2179 and MRS2500 did not affect ATP response (E,F left), but significantly reduced β-NAD response (E, F right). Scale bars in D and F apply to traces in A-D and E-F, respectively.

136

Fig 3.5

Figure 3.5. Expression of P2Y receptors in monkey and human tissues. (A) Expression of P2Y1, P2Y2, P2Y4, P2Y6 and P2Y11 receptor transcripts in the tunica muscularis of the monkey fundus, antrum, jejunum and proximal colon. (B) Expression of P2Y1, P2Y2, P2Y4, P2Y6 and P2Y11 receptor transcripts in human brain, fundus, gastric body (G. Body), antrum, ascending colon (A. Colon), distal colon (D. Colon) and sigmoid colon (S. Colon). C ytoglobin was used as a house keeping gene and M represents base pair marker. 137

Fig 3.6

Figure 3.6. β-NAD and ATP activate inward currents in SMCs of monkey (A-C) and human (D-F). (A) In cell-attached patches (pipette CaPSS; bath HK), channel openings were negligible. β-NAD (1mmol/L) increased openings at -80 mV. (B) Stepping from -80 to +80 mV showed β-NAD- activated currents reversed at 0 mV in monkey SMC. a (control) and b (β-NAD) show expanded traces from panel A during ramp depolarization (-80 mV to +80 mV). (C) ATP (1 mmol/L) increased channel activity at -80 mV in monkey SMC. (D) Perforated whole-cell conditions (pipette Cs-TEA, bath CaPSS), β-NAD (5 mmol/L) activated inward current at -80 mV. β-NAD activated-currents reversed upon washout. (E) a, b, and c show expanded traces from panel D during ramp depolarization. β-NAD activated-currents reversed at 0 mV, demonstrating non- selective cation conductance was activated by β-NAD. Dotted lines in B and E denote 0 mV and 0 pA. (F) ATP (1 mmol/L) activated inward currents at -80 mV. 138

3.5 Discussion

This study demonstrates several new features of enteric inhibitory motor neurotransmission in human and non-human primate colonic muscles. Stimulation of intrinsic neurons caused release of ATP and β-NAD, and β-NAD was the dominant purine released. β-NAD was previously suggested as an enteric inhibitory neurotransmitter in murine colonic muscles (Mutafova-Yambolieva et al., 2007) but it has never been considered as a candidate for neurotransmission in human GI smooth muscles. The characteristics of β-NAD release suggest it is more likely to be a neurotransmitter than ATP. For example, release of β-NAD, but not ATP, was frequency- dependent and blocked by neurotoxins. Localized β-NAD application mimicked purinergic responses to enteric inhibitory neurotransmission; antagonists that successfully blocked neural responses also blocked responses to exogenous β-NAD. This was not true for ATP, because MRS2179 and MRS2500 effectively blocked purinergic IJPs but failed to block responses to ATP. These data suggest that β-NAD could be the primary purinergic enteric inhibitory neurotransmitter in human and non-human primate colonic muscles. Purinergic neurotransmission may be the dominant form of enteric inhibitory control in human colon as brief stimuli failed to elicit nitrergic responses (Supplementary

Data), as commonly elicited in laboratory animals, and block of NO synthesis failed to reveal tonic nitrergic inhibition, as commonly observed in animal models (Lyster et al.,

1995).

ATP and β-NAD were released from human and monkey colonic muscles spontaneously and during nerve stimulation. At low frequency stimulation, β-NAD 139 appeared to be released from motor nerve terminals, but ganglionic release may also occur at higher frequencies. ATP release was equivalent in muscles with and without ganglia, and ATP release was not significantly reduced by neurotoxins. EFS-evoked overflow of the ATP metabolite, ADP, however, was significantly reduced by ω- conotoxin GVIA in WM, suggesting that a portion of ATP might have originated in ganglia but was degraded to ADP. β-NAD release exceeded ATP by 15-25 fold, suggesting that higher concentrations of β-NAD might be achieved near post-junctional receptors. Taken together, these data suggest that β-NAD may serve as an inhibitory neurotransmitter.

Tissue superfusates contained not only β-NAD and ATP but also their metabolites

ADP, AMP, ADPR, cADPR, and adenosine. Thus, mechanisms for terminating actions of purines are available in colon muscles. As in murine colon (Mutafova-Yambolieva et al., 2007) β-NAD is the primary purine in β-NAD+ADPR+cADPR peaks in human and monkey colons. Unlike murine colon, where ADPR comprised only ~6% of the β-

NAD+ADPR+cADPR mixture, ADPR contributes 12%-33% in the human and monkey colons. Future studies should determine whether ADPR originates only from degradation of β-NAD, or whether ADPR is also released from nerves. If the latter is true, ADPR may also elicit responses in post-junctional cells and be a co-transmitter.

Comparisons of postjunctional responses to neurotransmitters released during

EFS with responses to exogenous, putative transmitters, ATP and β-NAD, shed new light on the nature of enteric inhibitory neurotransmission in the human colon. In agreement with previous studies (Gallego et al., 2006; Grasa et al., 2009; Keef et al., 1993;

Mutafova-Yambolieva et al., 2007; Serio et al., 2003) electrical and mechanical 140 responses of human and monkey colons to EFS demonstrated P2Y receptor- and NO- mediated components (Supplementary Data). In monkey and human muscles the primary component of IJPs, a non-nitrergic, fast hyperpolarization, was significantly reduced by the nonselective P2Y receptor antagonists PPADS and suramin and by the P2Y1 inhibitors MRS2179 and MRS2500 (Cattaneo et al., 2004; Camaioni et al., 1998) IJPs in the human colon were less sensitive to 0.3 μmol/L apamin than monkey responses, and this finding is consistent with previous reports demonstrating relative insensitivity of human colonic (Gallego et al., 2006; Keef et al., 1993) and intestinal (Xue et al., 1999) muscles to apamin. Therefore, our data are in agreement with previous studies (Gallego et al., 2008a; Gallego et al., 2006) showing that IJPs in human colon are mediated by

P2Y1 receptors through a pathway that involves, in part, activation of apamin-sensitive channels. The major K+ conductance activated during purinergic IJPs in the human is unknown. Of major importance in this study is the finding that β-NAD also elicited hyperpolarization via P2Y1 receptors, but ATP responses were not blocked by MRS2179 and MRS2500. These data, coupled with findings on neurotransmitter release, suggest that β-NAD is a better candidate for the purine inhibitory neurotransmitter than ATP.

There are, however, additional explanations (as below) why the P2Y1 receptor antagonists were effective in blocking neurogenic responses but incapable of blocking responses to exogenous ATP. Thus, we cannot completely rule out a role for ATP as a neurotransmitter.

IJPs may be mediated by different post-junctional receptors, possibly expressed by different cell types, than hyperpolarization responses to exogenous substances.

Utilization of specific receptors may result from compartmentalization of P2Y1 receptors 141 in restricted receptive fields close to sites of neurotransmitter release and broader expression of purinergic receptors in cells not closely associated with active zones of transmitter release. In fact tunica muscularis expresses multiple P2Y receptors, including

P2Y1, P2Y2, P2Y4, P2Y6, and P2Y11, but specific junctional and/or extrajunctional distribution could not be delineated because the antibodies we tested performed poorly in immunohistochemistry. β-NAD might be a more exclusive agonist for P2Y1 receptors than ATP, and therefore responses to exogenous β-NAD were readily blocked by

MRS2179 and MRS2500. In contrast, responses to exogenous ATP appear to be mediated by receptors other than P2Y1 receptors, and therefore insensitive to the antagonists. ATP, if released from nerves, might generate responses via a specialized, junctional population of P2Y1 receptors, and our data do not rule out this possibility.

β-NAD- and ATP-induced hyperpolarizations of human and monkey colonic muscles were modest in comparison to IJPs. Multiple factors may be involved in this difference, including degradation of purines, insufficient penetration of β-NAD and ATP to appropriate receptors, or simultaneous activation of opposing conductances.

Picospritzing drugs close to recording sites, instead of superfusing drugs, was meant to simulate rapid direct application and avoid some of the factors that might dampen responses. β-NAD and ATP in picospritz pipettes were likely comparable to concentrations of neurotransmitters achieved near nerve terminals following exocytosis, since synaptic vesicles have been estimated to contain 5-1000 mmol/L neurotransmitters

(Van der Kloot, 2003; Borycz et al., 2005). Close examination of electrical responses to

β-NAD in monkey colonic muscles (Fig. 3.4) after P2Y antagonists revealed transient depolarizations in response to β-NAD. Equivalent observations were made in contractile 142 responses of human muscles to β-NAD where transient contractions often occurred before relaxation (Supplementary Figure S3.2). We considered the possibility that β-

NAD might activate opposing conductances in different cells, and tested the effects of this compound on isolated SMC. The endogenous purine neurotransmitter and exogenous

β-NAD caused hyperpolarization. In monkey cells, hyperpolarization resulted, in part, from activation of apamin-sensitive (SK) channels. Thus, if the effects of β-NAD were mediated by SMC, one would expect activation of SK channels in these cells. In fact, the dominant effect of both ATP and β-NAD was activation of non-selective cation channels in SMC. These observations support the idea that purinergic neurotransmission is targeted toward specific cells in GI muscles, and SMC are unlikely to mediate purinergic

IJPs.

There are two additional cells in close proximity to nerve terminals and coupled to SMCs via gap junctions in GI muscles: i) interstitial cells of Cajal (ICC) (Sanders,

1996), and ii) fibroblast-like cells (FLC) (Horiguchi & Komuro, 2000). It has not been possible to identify either ICC or FLC in dispersions of human or monkey GI muscles.

Previous studies on mice, however, suggest that ICC are unnecessary for purinergic responses (Sergeant et al., 2002), but FLC express SK3 channels abundantly (Klemm &

Lang, 2002; Vanderwinden et al., 2002; Fujita et al., 2003; Iino & Nojyo, 2009). Thus,

FLC, not SMC, may mediate purinergic IJPs in colonic muscles. Our data demonstrate that when β-NAD and ATP are spritzed or bath applied to colonic muscles, activation of opposing conductances tends to antagonize hyperpolarization responses. 143

The present study demonstrates the complexity of the enteric purinergic signaling and suggests that the β-NAD system can be a new target for treating motility disorders of the large intestine.

144

3.6 Supplementary material

3.6.1 Supplemental Experimental Procedures

RNA isolation and RT-PCR

Total RNA was extracted using TRIZOL (Life Technologies, Gaithersburg. MD).

The manufacturer's instructions were followed, including use of polyinosinic acid (20 μg) as an RNA carrier. First-strand complementary DNA was prepared from the RNA using the Superscript II reverse transcriptase (Life Technologies, Rockville, MD). One microgram total RNA was reverse-transcribed with 200 U reverse transcriptase in a 20-

μL reaction containing 25 ng oligo dT(12-18) primer, 500 μmol/L each dNTP, 50 mmol/L Tris-HCl (pH 8.3), 75 mmol/L KCl, 3 mmol/L MgCl2, and 10 mmol/LDTT.

Polymerase chain reaction (PCR) was performed with gene-specific primers for human

P2Y1, P2Y2, P2Y4, P2Y6, and P2Y11 (Supplementary Table S3.1) on 2 μL complementary DNA using Advantage 2 polymerase mix reagents (Clontech, Mountain

View, CA). Human brain was used as a positive control. A 2-step PCR method (95°C for

10 minutes, then 40 cycles of 95°C for 15 seconds and 60°C for 1 minute) was used to amplify the primer pairs. After PCR, 2 μL RT-PCR product was analyzed on a 2.0% agarose gel.

145

3.6.2 Supplemental Results

Enteric Inhibitory Responses in the Colon

Spontaneous action potentials were superimposed upon resting membrane potentials in both species (Supplementary Figure S3.1). EFS (single pulses, 0.5 ms) evoked IJPs consisting of 2 components: a fast transient hyperpolarization followed by a sustained phase lasting for several seconds in monkey proximal colon. After relaxation of

IJPs, action potentials occurred at higher frequency and larger amplitude (rebound excitation). In the presence of NG-nitro-L-arginine (L-NNA), to block the nitrergic component of IJP, CM cells depolarized and action potential frequency increased

(Supplementary Figure S3.1A). After L-NNA, EFS evoked non-nitrergic responses consisting of a fast transient IJP and brief cessation of action potentials. Isometric contractions of monkey proximal colon were also measured. EFS produced transient relaxation and inhibition of phasic contractions. Marked rebound contraction was noted upon termination of EFS. L-NNA increased basal tone, reduced the amplitude of phasic contractions during EFS, and reduced the amplitude of rebound excitation after cessation of EFS. EFS also caused IJPs in human muscles, but the sustained nitrergic phase of IJPs was not apparent. L-NNA had little effect on IJPs in human colon, suggesting little contribution of nitric oxide in responses to brief EFS in human colon, as reported previously (Gallego et al., 2008a) EFS also relaxed human CM, and cessation of EFS was followed by a large rebound contraction. L-NNA caused little increase in basal tone in human colonic muscles, and had little effect on EFS-evoked relaxation or rebound contraction (Supplementary Figure S3.1B). β-NAD (0.5−10 mmol/L) caused relaxation 146 of monkey and human colonic smooth muscles; however, in some human colon preparations β-NAD caused transient contractions before relaxing the tissues

(Supplementary Figure S3.2).

IJPs were also recorded in the presence of nifedipine (1 μmol/L) to inhibit smooth muscle action potentials that mask postjunctional inhibitory responses. In monkey colon single pulses of EFS (0.5 ms) elicited IJPs with 2 distinct components (Supplementary

Figure S3.1C), a large amplitude transient component with amplitude averaging 22.8 ±

0.6 mV (n = 57) and half maximal duration of 1.21 ± 0.07 seconds, followed by sustained hyperpolarization (5.2 ± 0.5 mV amplitude; 7.8 ± 0.5 seconds duration). L-NNA (100

μmol/L) abolished the secondary component of IJP but had little or no effect on the primary component. Atropine (1 μmol/L) had no effect on the primary component of the

IJP. EFS of human CM evoked IJPs averaging 27.2 ± 0.8 mV (n = 41) with a duration of

1.41 ± 0.03 seconds (Supplementary Figure S3.1D). Human IJPs consisted of a single fast component with no distinct secondary phase. Addition of L-NNA (100 μmol/L) and atropine (1 μmol/L) did not affect IJPs, suggesting little or no contribution of nitric oxide or acetylcholine to responses evoked by brief pulses of EFS in human colon.

147

Fig S3.1

148

Supplementary Figure S3.1. Inhibitory neural regulation of colonic muscles. (A) Electrical (top trace) and contractile (bottom trace) activity from monkey colonic muscle (recordings not simultaneous). EFS (arrows) caused IJPs, consisting of transient purinergic phase and sustained, nitrergic phase. EFS (0.3ms; 5Hz; 30s; black bars) caused transient relaxation and inhibition of phasic contractions. Rebound followed stimulus. L-NNA increased basal tone (dashed line denotes break in record). With L-NNA, EFS evoked transient relaxation, but no rebound contraction. (B) Intracellular electrical (top traces) and contractile (bottom traces) activity from human colonic muscle before and after L-NNA. EFS (arrow) caused IJP followed by rebound excitation. L-NNA did not depolarize cells, as seen in monkey. IJP was not reduced or blocked by L-NNA. EFS (0.3ms; 5Hz, 30s) caused transient relaxation and inhibited phasic contractions. Pronounced rebound contraction followed stimulus (lower traces in B). L-NNA did not increase basal tone (dashed line) and did not affect relaxation responses or post-EFS rebound. (C) In monkey colonic CM EFS (0.5 ms; 1 pulse at solid circle, left trace) evoked bi-phasic IJPs, with fast primary component and smaller secondary component inhibited by L-NNA (100 µmol/L, middle trace). Atropine (1 µmol/L, right trace) had no effect on IJPs. (D) In human colonic muscles EFS (0.5ms; 1 pulse) evoked mono-phasic IJPs (left trace). L-NNA (center trace) and atropine (right trace) did not affect IJPs. Experiments performed in nifedipine (1 µmol/L).

149

Fig S3.2

150

Supplementary Figure S3.2. Concentration-dependent inhibition of monkey and human colonic muscles by β-NAD. Concentration-dependent inhibition of contractile activity of monkey (A) and human (B) colonic muscles by β-NAD. β-NAD was added at various concentrations at time points indicated by the black bars and removed to allow recovery of spontaneous contractions between applications. In some human colonic preparations β-NAD caused transient enhancement in contraction (B, lower trace, arrows) followed by sustained relaxation. (C) Monkey and (D) human. Summary dose- response curves for 7 experiments with monkey colons and 8 experiments with human colons. (E) and (F) Inhibition of spontaneous contractile activity by noncholinergic, non-nitrergic inhibitory neural inputs; experiment performed in the presence of L-NNA (100 μmol/L) and atropine (1 μmol/L) in monkey and human colonic muscles, respectively. Nerve responses were evoked by electrical field stimulation (EFS) using 5 Hz for 30 seconds (0.3-ms pulses at 5 Hz for 30 seconds, left traces in each panel). Neurally evoked inhibitory responses were blocked by MRS2179 (right traces in each panel).

151

Table S3.1

Supplementary Table S3.1. Oligonucleotide Primers for P2Y Receptors 152

Table S3.2

Total purines ADP AMP ADO

4 Hz 5.81 ± 0.57 (22) 3.31 ± 0.38 (22) 18.38 ± 2.92 (22) 44.87 ± 7.6 (22)

16 Hz 8.26 ± 1.7 (36) 8.26 ± 1.7 (36) 45.94 ± 13.36 (36) 98.02± 18.27 (36)o

TTX, 16 Hz 4.34 ± 0.45 (21) 2.76 ± 0.34 (21) 19.02 ± 3.2 (21) 37.95 ± 6.4 (21)*

ω-Ctx GVIA,

3.7 ± 0.47 (7) 2.92 ± 0.57 (7) 24.68 ± 4.91 (7) 40.07 ± 5.7 (7) 16 Hz

Supplementary Table S3.2. Human Proximal Colon. Electrical field stimulation (0.3 ms, 480 pulses)-evoked overflow of metabolic products of ATP and β-NAD and total purines (ATP+ADP+AMP+ADO+β-NAD+ADPR+cADPR) in fmol/mg tissue, number of experiments in parentheses, oP = .0321 vs 4 Hz. *P = .0174 vs 16 Hz. Note that each experiment is performed in a separate tissue. Therefore, experiments at 4 Hz and at 16 Hz controls or at 16 Hz controls and tetrodotoxin (TTX) at 16 Hz were all carried out in separate tissues.

153

Table S3.3

Total purines ADP AMP ADO

Circular Circular Circular Circular Whole Whole Whole Whole

4 Hz 9.7±2.8 7.6±1.7 3.6±0.6 7.6±3.1 29.1±6.9 42.6±20.2 61.7±9.8 74.4±21.5

(10) (4) (10) (4) (10) (4) (10) (4) 16 Hz 6.09±0.8 5.2±0.6 8.4±1.8 4.6±0.7 110.9±27 36.3±7.1 223.7±46.6 80.2±10.2

(34) (18) (34) (18) (34) (18) (34) (18)

TTX, 3.7±0.61 5.8±1.7 2.7±0.5 3.6±1.1 44.6±14 28.1±6.7 71.3±18.8 46.5±9.1

16Hz (11) (6) (11) (6) (11) (6) (11) (6)

ω- 2.5±0.26 4.11±1.6 3.38±.86 4.5±1.3 45.9±14.7 25.0±3.5 68.41±19.4 49.8±9.2 CtxG, (16)** (6) (16) (6) (16) (6) (16)* (6) 16Hz

Supplementary Table S3.3. Monkey Proximal Colon. Electrical field stimulation (EFS; 0.3 ms, 480 pulses)-evoked overflow of metabolic products of ATP and β-NAD and total purines (ATP+ADP+AMP+ADO+β-NAD+ADPR+cADPR) in fmol/mg tissue, number of experiments in parentheses. **P < .01 vs 16 Hz controls. Note that each experiment is performed in a separate tissue. 154

3.7 Reference List

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159

Chapter 4

Adenosine 5’-diphosphate-ribose is a neural regulator in primate and murine large intestine along with β-NAD+

Leonie Durnin*, Sung Jin Hwang*, Sean M. Ward, Kenton M. Sanders and Violeta N. Mutafova-Yambolieva Published in Journal of Physiology 2012 Apr; 590(Pt 8):1921-41

*These authors contributed equally to this work

160

4.1 Abstract

Adenosine 5′-triphosphate (ATP) has long been considered to be the purine inhibitory neurotransmitter in gastrointestinal (GI) muscles, but recent studies indicate that another purine nucleotide, β-nicotinamide adenine dinucleotide (β-NAD+), meets pre- and postsynaptic criteria for a neurotransmitter better than ATP in primate and murine colons. Using a small-volume superfusion assay and HPLC with fluorescence detection and intracellular microelectrode techniques we compared β-NAD+ and ATP metabolism and postjunctional effects of the primary extracellular metabolites of β-NAD+ and ATP, namely ADP-ribose (ADPR) and ADP in colonic muscles from cynomolgus monkeys and wild-type (CD38+/+) and CD38−/− mice. ADPR and ADP caused membrane hyperpolarization that, like nerve-evoked inhibitory junctional potentials (IJPs), were inhibited by apamin. IJPs and hyperpolarization responses to ADPR, but not ADP, were inhibited by the P2Y1 receptor antagonist (1R,2S,4S,5S)-4-[2-iodo-6-(methylamino)-9H- purin-9-yl]-2-(phosphonooxy)bicyclo[3.1.0]hexane-1-methanol dihydrogen phosphate ester tetraammonium salt (MRS2500). Degradation of β-NAD+ and ADPR was greater per unit mass in muscles containing only nerve processes than in muscles also containing myenteric ganglia. Thus, mechanisms for generation of ADPR from β-NAD+ and for termination of the action of ADPR are likely to be present near sites of neurotransmitter release. Degradation of β-NAD+ to ADPR and other metabolites appears to be mediated by pathways besides CD38, the main NAD-glycohydrolase in mammals. Degradation of

β-NAD+ and ATP were equal in colon. ADPR like its precursor, β-NAD+, mimicked the effects of the endogenous purine neurotransmitter in primate and murine colons. Taken 161 together, our observations support a novel hypothesis in which multiple purines contribute to enteric inhibitory regulation of gastrointestinal motility.

4.2 Introduction

Enteric inhibitory motor neurotransmission in the gastrointestinal (GI) tract is mediated largely by nitric oxide (Bult et al., 1990; Sanders & Ward, 1992) and a purine nucleotide, thought for many years to be ATP (Burnstock et al., 1970; Burnstock, 2008b).

Recent studies of murine colon (Mutafova-Yambolieva et al., 2007) and human and non- human primate (Macaca fascicularis) colons (Hwang et al., 2011) have shown that β- nicotinamide adenine dinucleotide (β-NAD+) better meets presynaptic and postsynaptic criteria for the inhibitory purine neurotransmitter than ATP. Frequency-dependent, neurotoxin-sensitive release of β-NAD+, but not of ATP, was demonstrated in colonic muscles, and exogenous β-NAD+ caused postsynaptic hyperpolarization that mimicked the pharmacology of inhibitory junction potentials (IJPs), generated by the endogenous purine neurotransmitter, better than ATP. In fact, most ATP appeared to originate from non-neuronal sources in colonic muscles, because a large part of the ATP release was insensitive to neurotoxins. Nevertheless, some portion of ATP may have been released from neurons, and the failure of exogenous ATP to mimic neural responses might have been due to activation of multiple postjunctional receptors, compartmentalization of postjunctional purine receptors, and/or rapid enzymatic degradation of ATP.

β-NAD+ released into the interstitium is degraded to nicotinamide and ADPR and to cADPR by NAD glycohydrolase and ADP-ribosyl cyclase, respectively, which in 162 mammals are mainly associated with the CD38 protein (Munshi et al., 2000; Lee, 2001;

Graeff et al., 2009). NAD glycohydrolase and ADP-ribosyl cyclase account for about

98% and 2% of the CD38 enzymatic activities (Lee, 2001). ADPR is further degraded to

AMP by ectonucleotide pyrophosphatases (E-NPPs, CD203 family) and AMP is degraded to adenosine (ADO) by 5′-nucleotidase, CD73 (Di Girolamo et al., 1997;

Zimmermann, 2000). ATP is degraded to ADP, AMP and ADO by ectonucleoside 5′- triphosphate diphosphohydrolases (E-NTPDases, CD39 family), nucleotide pyrophosphatases (NPPs), and 5′-nucleotidase, respectively (Zimmermann, 2000). These enzymes not only control the lifetime of purine nucleotides, they also generate agonists for additional purine receptors expressed by various cells within GI muscles (Abbracchio et al., 2009). We have demonstrated previously that tissue superfusates contain β-NAD+ and ATP, as well as their metabolites, ADP, AMP, ADPR, cADPR and ADO (Mutafova-

Yambolieva et al., 2007; Hwang et al., 2011). However, it is currently unknown whether the direct metabolites of β-NAD+ and ATP, ADPR and ADP, also contribute to the postjunctional responses elicited by purinergic inhibitory neurotransmission.

The effects of purinergic neurotransmitter(s) released from enteric inhibitory neurons are known to be mediated by postjunctional P2Y1 receptors (Gallego et al.,

2006; Gallego et al., 2011; Mutafova-Yambolieva et al., 2007; Hwang et al., 2011).

P2Y1 receptor-mediated effects can be activated by ADP and ATP (von Kugelgen, 2006;

Mutafova-Yambolieva et al., 2007) and by β-NAD+ (Mutafova-Yambolieva et al., 2007;

Klein et al., 2009; Hwang et al., 2011) and ADPR (Gustafsson et al., 2011). The effects of exogenous β-NAD+, but not ATP, are blocked by selective and specific P2Y1 receptor inhibitors (Mutafova-Yambolieva et al., 2007; Hwang et al., 2011). It is currently 163 unknown whether ADPR and ADP mimic the pharmacology of purinergic neurotransmission. In the current study we compared the degradation of β-NAD+ and

ATP in the tunica muscularis and in isolated circular muscles of monkey colon at rest and during nerve stimulation and examined whether CD38, the major extracellular β-NAD+- metabolizing protein in mammals, is responsible for the degradation of β-NAD+ in the large intestine. We also tested whether metabolites of β-NAD+ and ATP (ADPR and

ADP, respectively) display postjunctional effects that mimic the endogenous neurotransmitter(s). Our data suggest that ADPR, whether it is produced by metabolism of extracellular β-NAD+ or released as a primary neurotransmitter, may contribute to enteric inhibitory neurotransmission in GI muscles.

4.3 Methods

4.3.1 Ethical procedures

Cynomolgus monkeys (Macaca fascicularis), untreated with other drugs, were sedated with ketamine (10 mg kg−1) and 0.7 ml Beuthanasia-D (Schering-Plough AH,

Kenilworth, NJ, USA) and were exsangunated at Charles River Laboratories Preclinical

Services (Reno, NV, USA) for the purpose of clinical trials unrelated to this project. No cynomolgus monkeys were killed specifically for the experiments described in this paper.

Proximal colon was retrieved from untreated, control animals by the Charles River

Laboratories staff, placed in ice-cold Krebs–Ringer buffer (KRB) and transported to the

University of Nevada site of experimental work. Use of muscles from these animals and transport of tissues to laboratories at the University of Nevada was approved by the 164

Institutional Animal Care and Use Committee at University of Nevada (IACUC).

C57/BL6 wild-type and CD38−/− mice were killed by sedation with isoflurane followed by cervical dislocation and exsanguination. The entire gastrointestinal tract, from oesophagus to the internal anal sphincter, was removed and placed in oxygenated ice- cold KRB for further dissection. All experimental procedures were approved by the

IACUC.

4.3.2 Tissue preparation

Monkey circular muscles (CMs) were prepared by peeling away the longitudinal muscle with attached myenteric ganglia. Other experiments were carried out using whole tunica muscularis (whole muscle, WM). Proximal colons of C57BL/6 mice (wild-type,

CD38+/+, Charles River Laboratories, Wilmington, MA, USA) and CD38 knockout mice

(CD38−/−, The Jackson Laboratory, Bar Harbor, ME, USA) were prepared by peeling away the mucosa and submucosa.

4.3.3 Degradation of purine nucleotides in mouse and monkey colonic smooth muscle

To determine enzymatic degradation of β-NAD+, ATP, and ADPR colonic preparations were superfused with 1,N6-etheno-NAD (eNAD) and 1,N6-etheno-ATP

(eATP) substrates. 1,N6-Etheno-nucleotides have been used previously as exogenous substrates for nucleotidases (Secrist et al., 1972; Jamal et al., 1988; Todorov et al., 1997;

Mihaylova-Todorova et al., 2001; Bobalova & Mutafova-Yambolieva, 2003), since they are highly fluorescent and allow about 1,000,000-fold more sensitive detection of nucleotide metabolism compared to authentic nucleotides (Bobalova et al., 2002). Non- 165 derivatized purine nucleotides cannot be used in studies in small tissue preparations, because the authentic purine nucleotides generally have very low fluorescence coefficients, not allowing detection of small changes in substrate or product concentrations.

Smooth muscle tissue segments of mouse or monkey tunica muscularis or monkey circular muscle were placed in 200 μl water-jacket Brandel superfusion chambers equipped with platinum electrodes as used in neurotransmitter release studies and described previously (Bobalova & Mutafova-Yambolieva, 2001; Mutafova-

Yambolieva et al., 2007; Hwang et al., 2011). Chambers were mounted vertically and tissues were superfused with oxygenated Krebs solution (at 37°C) with the following composition (mM): 118.5 NaCl, 4.2 KCl, 1.2 MgCl2, 23.8 NaHCO3, 1.2 KH2PO4, 11.0 dextrose, 1.8 CaCl2 (pH 7.4). After a 45 min equilibration, tissues were superfused with either eATP or eNAD, at maximum saturating concentrations of 0.05 and 0.2 μM, respectively. Maximal saturating concentrations of eATP or eNAD were determined in preliminary concentration-metabolism experiments (data not shown). Three hundred microlitres of the superfusion solution were collected from the beaker containing the substrate (S0, (–) tissue). Typically the tissue samples were treated with the substrates for

30 or 60 s and the contents of the chamber were then collected in ice-cold Eppendorf tubes (S1, (+) tissue); the enzymatic reactions stopped by immersing the tubes in liquid nitrogen. In some experiments tissues were treated with substrate for 1 s and 5 s (30 min apart) to determine whether the same degradation of substrate occurs at shorter contact with tissue. In some experiments tissues were subjected to electrical field stimulation 166

(EFS; 480 pulses; 0.3 ms or 0.5 ms; 16 Hz) once the substrate had reached the chamber to evaluate potential ‘releasable' nucleotidase activity (S2, (+) tissue, (+) EFS).

The degradation of eNAD was evaluated by (1) the formation of its end product eADO, and (2) the sum of its products eADPR, eAMP and eADO. For the latter analysis we had to determine first the composition of the 12.5 min peak and evaluate the quantity of eAMP and eNAD as described in HPLC fraction analysis. Note that the formation of cADPR from eNAD cannot be measured in these experiments, because the N6 of the adenine moiety of eNAD is unavailable for cyclization and, therefore, 1,N6-etheno- cADPR cannot be formed (Smyth et al., 2006a). Therefore, to evaluate the ADP-ribosyl cyclase activity degrading β-NAD+ to cADPR we used nicotinamide guanine dinucleotide (NGD, 0.2 mM) as substrate (Graeff et al., 1998a) and measured the formation of cyclic guanosine diphosphate ribose (cGDPR) as shown previously (Smyth et al., 2006a). eATP metabolism was determined by the production of eADO as well as by the appearance of eADP + eAMP + eADO in superfusates collected during superfusion of tissues with eATP. eADPR degradation was determined by the production of eAMP and eADO. HPLC coupled with fluorescence detection was used to measure formation of 1,N6-etheno-nucleotides and nucleosides and cGDPR as described below.

The amount of products formed in S1 was compared with S0 to measure the spontaneous enzyme activity. The amount of products in S2 was compared with S1 to determine whether additional ‘releasable' activity (Todorov et al., 1997) is present in the large intestine. Endogenously released nucleotides were not derivatized in these experiments; thus, the formation of 1,N6-etheno-nucleotides and nucleosides corresponds to direct products of 1,N6-etheno-substrates without interference of endogenous nucleotides. 167

4.3.4 HPLC assay of etheno-nucleotides/nucleosides and cGDPR

A reverse-phase gradient Agilent Technologies 1200 liquid chromatography system (Agilent Technologies, Wilmington, DE, USA) was used to detect the 1,N6- etheno-products and cGDPR as described previously (Mutafova-Yambolieva et al., 2007;

Hwang et al., 2011). The mobile phase consisted of 0.1 M KH2PO4 (pH 6.0) as eluent A.

Eluent B consisted of 65% eluent A and 35% methanol. Gradient elution was employed according to the following linear programme: time 0, 0% eluent B; 18 min, 100% eluent

B. The flow rate was 1 ml min−1 and run time was 20 min. Column and autosampler temperatures were maintained at 25°C and 4°C, respectively. The fluorescence detector was set to record 1,N6-etheno-derivatized nucleotide and nucleoside signals at an excitation wavelength of 230 nm and emission wavelength of 420 nm. cGDPR was detected at an excitation wavelength of 270 nm and emission wavelength of 400 nm.

These are optimum conditions for detection of 1,N6-etheno-derivatized (Bobalova et al.,

2002) and non-etheno-derivatized (Smyth et al., 2004) nucleotides/nucleosides, respectively. The amount of nucleotide/nucleoside in each sample was calculated from calibration curves of nucleotide standards run simultaneously with each set of unknown samples. Results were normalized for sample volume and tissue weight and product formation was expressed in fmol (mg tissue)-1.

4.3.5 HPLC fraction analysis

eNAD elutes in the HPLC column at ∼12.5 min which is also the retention time of the eNAD product eAMP. Therefore, HPLC fraction analysis was carried out to determine the compounds present in the 12.5 min peak and the proportions of eAMP and 168 eNAD in this peak. An Agilent Technologies 1200 Analytical Fraction Collector was employed to collect the peak at 12.5 min. The sample was etheno-derivatized (see below) and re-injected in the HPLC system. Etheno-derivatization at high temperature caused eNAD to be converted to eADPR (Smyth et al., 2004) which elutes at 11.5 min in the

HPLC column. The peak that remained at 12.5 min was eAMP.

4.3.6 Preparation of etheno-substrates

ATP, ADO and ADPR were purchased from Sigma-Aldrich (St Louis, MO, USA) and etheno-derivatized as follows: 0.2 mM ATP, ADO or ADPR (dissolved in double distilled water) was acidified to pH 4.0 with citrate phosphate buffer. Chloroacetaldehyde

(1 M) was added and substrates were heated to 80°C for 40 min to form 1,N6- ethenoderivatives eATP, eADO and eADPR (Levitt et al., 1984; Bobalova et al., 2002). eNAD was purchased from BioLog Life Science Institute (Bremen, Germany). Substrates were further diluted in the superfusion solution to final concentration needed.

4.3.7 Preparation of cGDPR

cGDPR, used as a standard for the HPLC analysis of enzymatic degradation of

NGD, was prepared as previously described (Smyth et al., 2006a). Briefly, nicotinamide- guanine dinucleotide (NGD, 2 mM, Sigma-Aldrich, MO, USA) was incubated with 2.5 units of Aplysia californica cyclase (Sigma-Aldrich, MO, USA) in reaction solution containing Tris-HCl (20 mM) in double distilled water (total volume 1 ml) for 1 h at

37°C. Quantitative conversion of NGD to cGDPR was confirmed by HPLC analysis of

10 μl aliquots of the reaction mixture. 169

4.3.8 Intracellular electrical activity and force measurements

Circular muscles from monkey and mouse (approximately 10 mm × 10 mm) were pinned to the Sylgard elastomer-lined floor of a recording chamber with the myenteric plexus side of the circular muscle facing upward. The bath chamber was constantly perfused with oxygenated Krebs–Ringer buffer (KRB) of the following composition

(mM): 118.5 NaCl, 4.5 KCl, 1.2 MgCl2, 23.8 NaHCO3, 1.2 KH2PO4, 11.0 dextrose, 2.4

CaCl2, pH 7.4, at 37 ± 0.5°C. After 1 h equilibration circular muscle cells were impaled with glass microelectrodes with 80–100 MΩ resistance filled with 3 M KCl as described previously (Burns et al., 1996; Mutafova-Yambolieva et al., 2007). Experiments were performed in the presence of nifedipine (1 μM) to reduce contractions and facilitate impalements of cells for extended periods except where stated. Parallel platinum electrodes were placed on either side of the muscle strips and neural responses were evoked by square wave pulses of electrical field stimulation (EFS; 0.5 ms pulse duration) using a Grass S48 stimulator (Quincy, MA, USA). In each series of experiments agonists

(ATP, ADP or ADPR each at a concentration of 10 mM or 50 mM β-NAD+) were applied (10 ms pulses at 10 psi) via picospritz micropipettes (Picospritzer; General Valve,

East Hanover, NJ, USA) positioned close to the site of electrical recording before and after addition of purinergic antagonists onto tissues. Responses to picospritzed agonists were compared with responses evoked by electrical field stimulation of intrinsic nerves.

In one series of experiments concentration–response effects of ADP and ADPR and the effects of the P2Y1 antagonist, MRS2500, on these responses were compared. These experiments were performed by loading spritz pipettes with the same concentrations of

ADP and ADPR (10 mM) and then applying multiple spritzes, ranging in pulse duration 170 from 10 to 100 ms, while maintaining impalements in the same cell. Spritz pipette tips were positioned slightly farther from tissues in this series of experiments so impalements were not lost when longer duration spritzes (50–100 ms) were applied. In experiments in which mechanical activity was recorded, circular muscle strips of mouse colon (6 mm wide) were fixed at one end and attached to a Fort 10 isometric force transducer (World

Precision Instruments, Sarasota, FL, USA). Isometric force was measured as described previously (Burns et al., 1996).

4.3.9 RNA isolation and RT-PCR

Total RNA was isolated from colonic tunica muscularis of wild-type and CD38−/− mice using TRIzol reagent (Invitrogen, Carlsbad, CA, USA). RNA was treated with 1 U

μl−1 DNase I (Promega Corp., Madison, WI, USA) and cDNA was prepared using Oligo dT(12–18) primer and SuperScript II reverse transcriptase (Invitrogen). Polymerase chain reaction (PCR) was performed with gene-specific primers for CD38 (Kato et al., 1999), see below, on 2 μl complementary DNA using Advantage 2 polymerase mix reagents

(Clontech, Mountain View, CA, USA). A two-step PCR method (95°C for 10 min, then

40 cycles of 95°C for 15 s and 60°C for 1 min) was used to amplify the primer pairs.

After PCR, 2 μl RT-PCR products were analysed on a 2.0% agarose gel. The following primers were used in this assay: Cd38-F ACAGACCTGGCTGCCGCCTCTCTAG (Tm =

64°C) and Cd38-R, GGGGCGTAGTCTTCTCTTGTGATGT (Tm = 59°C); accession number NM-007646.

4.3.10 Western immunoblot analysis 171

Colons from monkey (WM and CM preparations) were frozen by immersion in liquid nitrogen. Frozen tissues were pulverized and total protein was extracted by glass– glass homogenization with a RIPA buffer composed of 20 mM Tris, 150 mM NaCl, 10% glycerol, 1% NP40, 2.5 mM NaF, 0.1 mM sodium orthovanadate, 1 mM benzamidine,

2.5 mM β-glycerophosphate, 100 μM AEBSF and 1 μM leupeptin. Insoluble material was pelleted by centrifugation at 15,000 g for 20 min at 4°C. Total protein concentration of the supernatant was determined by the BCA assay using bovine serum albumin for standards. Tissue homogenates were reduced with Laemmli reagent and equal amounts of total protein (10 μg) were resolved by SDS-PAGE (12% acrylamide) and transferred onto

PVDF membranes for 1.5 h at 100 V and 4°C (Bio-Rad, Hercules, CA, USA). HuT78 T lymphocyte lymphoma whole cell lysate (Santa Cruz Biotechnology, Santa Cruz, CA,

USA) was used as a positive control to identify the CD38 band. Membranes were blocked for 1 h with 2% non-fat dry milk + 0.2% Tween and probed for 18 h at 4°C with an anti-CD38 primary rabbit polyclonal antibody (Santa Cruz Biotechnology, cat no. sc-

15362), diluted 500-fold in the blocking solution. After removal of excess primary antibody, membranes were incubated for 1.5 h at room temperature with a secondary horseradish peroxidase-conjugated rabbit antibody. Immunostained protein bands were detected using ECL Advantage (GE HealthCare Biosciences, Piscataway, NJ, USA), and visualized with a CCD camera-based detection system (Epi Chem II, UVP Laboratory

Products, Upland, CA, USA).

4.3.11 Statistics 172

Data are presented as means ± SEM. Means were compared by Student's two- tailed, unpaired t test or by one-way ANOVA for comparison of more than two groups followed by a post hoc Bonferroni multiple comparison test (GraphPad Prism, v. 3

(GraphPad Software, Inc., San Diego, CA) or SigmaPlot (Systat Software Inc.,

Richmond, CA, USA)). A probability value of less than 0.05 was considered significant.

4.3.12 Drugs and chemicals

ATP, ADP, adenosine, ADP-ribose (ADPR), apamin, atropine, erythro-9-(2- hydroxy-3-nonyl)-adenine (EHNA), nicotinamide guanine dinucleotide (NGD), NG-nitro- l-arginine (l-NNA), nifedipine, pyridoxal-phosphate-6-azophenyl-2′,4′-disulfonate

(PPADS) and suramin were purchased from Sigma-Aldrich. 6-S-[(4-

Nitrophenly)methyl]-6-thioinosine (NBMPR or NBTI) and (1R,2S,4S,5S)-4-[2-iodo-6-

(methylamino)-9H-purin,-9-yl]-2-(phosphonooxy)bicyclo[3.1.0]hexane-1-methanol dihydrogen phosphate ester tetraammonium salt (MRS2500) were purchased from Tocris

Bioscience (Ellisville, MO, USA). β-Nicotinamide-1,N6- ethenoadenine dinucleotide

(eNAD) came from BioLog Life Science Institute (Bremen, Germany). All drugs were dissolved in deionized H2O, apart from nifedipine (dissolved in ethanol) and NBMPR

(dissolved in DMSO), and further diluted in perfusion solutions; final concentration of ethanol or DMSO in superfusion solution was 0.001%.

4.4 Results

4.4.1 β-NAD+ forms ADPR in primate colonic smooth muscles 173

We have previously shown that tissue superfusates collected during EFS of primate and murine colonic muscles contain ADPR, β-NAD+, ATP and ADP (Mutafova-

Yambolieva et al., 2007; Hwang et al., 2011). ADPR, in particular, comprised about 20–

30% of the β-NAD+ + ADPR + cADPR combination in primate colons (Hwang et al.,

2011). However, it is not clear whether ADPR is formed as a result of metabolism of β-

NAD+ released from nerves or whether it is a substance that is released upon activation of nerve terminals. Therefore, we first tested whether β-NAD+ is metabolized to ADPR in colonic muscles. Primate WM and CM colonic muscles were superfused with eNAD (0.2

μM), and the formation of eADPR and the end metabolite, eADO, was measured. To match neurotransmitter overflow studies (i.e. Hwang et al., 2011; Mutafova-Yambolieva et al., 2007), tissues were superfused with the substrate for 30–60 s, as this was the duration of EFS in previous transmitter overflow experiments. As shown in Fig. 4.1, eNAD substrate from BioLog Life Science Institute contains a small amount of eADPR

(Fig. 4.1A, (–) tissue). Clearly, eADPR is formed from eNAD in both WM and CM of the colon (Fig. 4.1A, middle and bottom chromatograms). Whereas eADO was not detected in the absence of tissue (Fig. 4.1A, top panel), eADO became detectable in the tissue superfusates when WM or CM was superfused with eNAD (Fig. 4.1A, middle and bottom panels). When normalized against tissue mass, the amount of eADO produced from eNAD was significantly higher in CM than in WM, P < 0.05 (Fig. 4.1B), suggesting that degradation of eNAD was greater in preparations containing a higher proportion of motor nerve processes and terminals per unit weight. Western immunoblot analysis of CD38, however, showed no significant differences in the levels of CD38 in WM and CM (Fig.

4.1C). No further degradation of eADO was observed when tissues were superfused with 174 eADO (0.05 μM) as a substrate (Table 4.1). Furthermore, the amounts of eADO remained unchanged in the presence of either the adenosine deaminase inhibitor (EHNA, 10 μM) or the adenosine uptake blocker NBMPR (10 μM) (Table 4.1). Similar results were obtained in murine colons (Table 4.1). Therefore, the formation of eADO appeared to be a relatively reliable measure for eNAD metabolism since no further degradation by adenosine deaminase (Agteresch et al., 1999) or adenosine transport (Zimmermann et al.,

1998) was detected in these preparations. Formation of eADPR and eADO from eNAD was observed even when the tissues were in contact with eNAD for 1 s or 5 s only. As discussed above eADO was not observed in the absence of tissue. However, eADO was

0.94 ± 0.17 (n = 8) and 1.34 ± 0.23 fmol (mg tissue)−1 (n = 7) after contact of eNAD with

WM for 1 s and 5 s, respectively. Likewise, in CM superfusates eADO was measured to be 1.45 ± 0.18 (n = 8) and 1.57 ± 0.21 fmol (mg tissue)-1 (n = 8) after contact with eNAD for 1 s and 5 s, respectively.

We determined the relative content of eAMP and eNAD in the 12.5 min (eNAD + eAMP) peak and analysed the degradation of eNAD by the formation of the total e- product eADPR + eAMP + eADO after 30–60 s superfusion with eNAD. For this, we first carried out an HPLC fractional analysis to determine the portion of eAMP in the

12.5 min chromatography peak: eAMP comprised ∼2.2% of the 12.5 min peak of the eNAD provided by BioLog Life Science Institute when no muscle was present. When muscles were superfused with eNAD, the proportion of eAMP in the 12.5 min peak was increased to 6.1% of the peak in WM and 3.12% of the peak in CM. In WM the sum of eADPR + eAMP + eADO was 7.37 ± 1.24 and 17.11 ± 1.64 fmol (mg tissue)−1 (n = 14, P

< 0.05) in the absence and presence of muscle, respectively. In experiments testing CM 175 eADPR + eAMP + eADO was 11.18 ± 1.37 and 24.28 ± 2.72 fmol (mg tissue)−1 (n = 20,

P < 0.001) in the absence and presence of muscle, respectively. Thus, the amount of eADPR + eAMP + eADO formed in CM was higher than in WM (P < 0.05), as also observed when eNAD degradation was evaluated by eADO formation.

We also tested whether degradation of eNAD increased in colonic muscles during nerve stimulation, as suggested for metabolism of purines in some smooth muscles innervated by autonomic neurons (Todorov et al., 1997). The degradation of eNAD in

WM and CM was not significantly increased during EFS. For example, in WM eADO was 4.39 ± 0.47 fmol (mg tissue)−1 in the absence of EFS (S1, n = 14) and 4.98 ± 0.67 fmol (mg tissue)−1 in the presence of EFS (S2, n = 14, P > 0.05). Likewise, in CM eADO was 7.37 ± 1.14 fmol (mg tissue)−1 in the absence of EFS (S1, n = 20) and 7.19 ± 1.09 fmol (mg tissue)−1 in the presence of EFS (S2, n = 20, P > 0.05). Therefore, conventional

HPLC analysis in conjunction with HPLC fractional analysis showed that: (i) eADPR is formed from eNAD in primate colonic muscles, (ii) degradation of eNAD occurs rapidly within muscles, (iii) degradation of eNAD is greater per unit mass in isolated CM than

WM, and (iv) no additional degradation could be resolved during nerve stimulation.

4.4.2 β-NAD+ is likely to be degraded by multiple enzymes in murine colon

To determine the role of CD38 and possibly other enzymes in the degradation of

β-NAD+ in the colon we next examined the degradation of eNAD and NGD in colonic preparations (tunica muscularis) isolated from strain-matched, wild-type (CD38+/+ mice) and from CD38−/− mice. We found equal amounts of eADO were generated from eNAD by muscles of wild-type and CD38−/− mice for 30 s contact with tissue (Fig. 4.2A and B). 176

Likewise, the amounts of total e-product eADPR + eAMP + eADO were comparable in colons from wild-type and CD38−/− mice: 27.28 ± 2.43 (n = 4) and 24.2 ± 2.78 fmol (mg tissue)−1 (n = 6) in wild-type and CD38−/− mice, respectively (P > 0.05). Therefore, either

NAD glycohydrolase activity is not attributed to CD38 in the murine colon or other enzymes, in addition to CD38, are also involved in the degradation of eNAD.

Importantly, eADO was also formed from eNAD after very brief contacts of the substrate eNAD with the tissue (i.e. for 1 s or 5 s, Fig. 4.3). Thus, in the absence of tissue eADO was not present (Fig. 4.3). However, in colonic preparations of wild-type mice eADO was 7.27 ± 1.76 (n = 6) and 10.49 ± 2.56 fmol (mg tissue)−1 (n = 6) after 1 s and 5 s contact of tissue with eNAD. Likewise, in colons isolated from CD38−/− mice the amount of eADO formed from eNAD after 1 s and 5 s of contact of tissues was 4.80 ±

1.2 (n = 6) and 6.92 ± 1.23 fmol (mg tissue)−1 (n = 6), respectively.

As in the monkey colons no further degradation of eNAD was observed during

EFS in wild-type and CD38−/− mice: eADO was 16.4 ± 3.74 fmol (mg tissue)−1 in S1 and

23.6 ± 4.47 fmol (mg tissue)−1 in S2 in wild-type mice (n = 4, P > 0.05) and 14.34 ± 1.69 fmol (mg tissue)−1 in S1 and 17.42 ± 1.56 fmol (mg tissue)−1 in S2 (n = 6, P > 0.05) in

CD38−/− mice. As shown in Fig. 4.2C RT-PCR analysis verified the lack of CD38 gene in the CD38−/− mice.

In contrast to eNAD, degradation of NGD to cGDPR occurred in colonic muscles from wild-type mice but not from CD38−/− mice (Fig. 4.4). These data demonstrate that the ADP-ribosyl cyclase activity in the murine colon is attributable to CD38.

4.4.3 Extracellular ATP is degraded in the colon 177

In a previous study we found that ATP was also released in colonic muscles during EFS, but ATP did not mimic the endogenous inhibitory neurotransmitter

(Mutafova-Yambolieva et al., 2007; Hwang et al., 2011). In the present study we investigated the metabolism of ATP by WM and CM preparations of monkey colon and by colonic muscles from wild-type and CD38−/− mice. As shown in Fig. 4.5, eATP, prepared from commercial ATP obtained from Sigma-Aldrich, contained a small amount of eADP, but no eAMP or eADO (Fig. 4.5A and C, (–) tissue). However, when WM or

CM was superfused with eATP, eADP appeared, along with eAMP and eADO (Fig.

4.5A, middle and bottom chromatograms), indicating that degradation of ATP is accomplished by colonic muscles. There was significant formation of eADO in both WM and CM preparations, and no significant difference in eATP metabolism was noted between WM and CM (Fig. 4.5A and B). eADO formed from eATP was comparable to eADO formed from eNAD. In WM eADO formed from eNAD was 4.39 ± 0.47 fmol (mg tissue)−1 (n = 14) and eADO formed from eATP was 7.13 ± 1.65 fmol (mg tissue)−1 (n =

15) (P > 0.05 comparison in eADO formation from eNAD and eATP). In CM eADO formed from eNAD was 7.37 ± 1.14 fmol (mg tissue)−1 (n = 20), and eADO formed from eATP was 10.57 ± 2.18 fmol (mg tissue)−1 (n = 22) (P > 0.05; comparison in eADO formation from eNAD and eATP).

eATP was also metabolized by murine colon (Fig. 4.5C and D). Interestingly, eADO formation from eATP appeared greater in colon from CD38−/− mice than in colons from wild-type mice. Analysis of eADP + eAMP + eADO also showed greater product formation in preparations from CD38−/− than wild-type mice: 24.15 ± 4.00 (n = 18), 38.3

± 4.14 (n = 10), and 47.55 ± 5.53 fmol (mg tissue)−1 (n = 8) in (–) tissue, wild-type and 178

CD38−/− mice, respectively (P > 0.05 vs. (–) tissue in wild-type animals and P < 0.01 vs.

(–) tissue in CD38−/− mice).

No further degradation of eATP occurred during EFS in all tissues studied. For example, the e-product (eADP + eAMP + eADO) formation was 47.85 ± 9.38 (n = 15) and 60.18 ± 12.16 fmol (mg tissue)−1 (n = 15) in the absence and presence of EFS in monkey WM, respectively (P > 0.05). The e-product was 53.06 ± 8.79 (n = 22) and 49.91

± 8.45 fmol (mg tissue)−1 (n = 22) in the absence and presence of EFS in monkey CM (P

> 0.05). Similar results were obtained using colonic muscles from wild-type and CD38−/− mice (n = 12 and n = 10, respectively, data not shown). Therefore, degradation of ATP occurs in monkey WM and CM colon and in the wild-type and CD38−/− murine colon; however, no ‘releasable' metabolic activity was observed in colonic muscles.

4.4.4 EFS-induced IJPs are sensitive to apamin and P2Y1 receptor inhibition in the monkey circular smooth muscle cells

The rapid appearance of metabolites of putative purine neurotransmitters raised the question of whether the metabolites might contribute to postjunctional responses.

Therefore, we investigated whether ADPR and/or ADP mimic responses to endogenous purine neurotransmitter(s) in monkey colonic preparations. In agreement with previous observations (Hwang et al., 2011), circular muscle cells of the monkey proximal colon had resting membrane potentials (i.e. most negative potentials between action potential complexes) averaging −49 ± 1 mV (n = 20). An ongoing noisy fluctuation in membrane potential, previously referred to as unitary potentials, was observed in these recordings

(Beckett et al., 2004; Hirst et al., 2004). IJPs were evoked in monkey colonic muscles by 179 electrical field stimulation (EFS; 1 pulse, 0.5 ms duration) and were reduced by apamin and the P2Y1 receptor antagonist MRS2500. Under control conditions (in the presence of atropine, 1 μM and l-NNA, 100 μM), IJPs were monophasic hyperpolarization responses,

24.3 ± 0.7 mV in amplitude and 0.77 ± 0.04 s in duration (n = 15). Apamin (0.3 μM) reduced the amplitudes of IJPs by 69 ± 4% (n = 7; P = 0.00001) and MRS2500 (1 μM) reduced the amplitude of the IJPs by 95 ± 2% (n = 8; P = 0.000001; Fig. 4.6A and B).

4.4.5 Hyperpolarization responses to localized application of exogenous ADPR and ADP

ADPR and ADP (10 mM loaded into picospritz pipettes) were applied (10 ms pulses) to circular muscles and electrical responses were recorded to evaluate the effects of ADPR and ADP on membrane potential. ADPR caused hyperpolarization of circular muscle cells, averaging 8 ± 1 mV (n = 10) that was sustained for several seconds before the membrane potential recovered to control levels (Fig. 4.6C and D, Control). ADP spritzes caused similar hyperpolarization responses, averaging 10 ± 0.5 mV (n = 10), which also persisted for several seconds (Fig. 4.6E and F, Control). The response to

ADPR was reduced 84 ± 3% (P = 0.006, n = 5) by apamin (0.3 μM) and 95 ± 3% (P =

0.0003; n = 5; bottom traces) by MRS2500 (1 μM). Examples are shown in Fig. 4.6C and

D. Apamin reduced the response to ADP by 78 ± 6% (P = 0.0004; n = 5), but the hyperpolarization to MRS2500 was reduced by only 38 ± 7%, and this did not reach a level of significance (P > 0.05; n = 5; Fig. 4.6E and F, bottom traces). Therefore, the pharmacology of ADPR responses better mimicked postjunctional responses to the endogenous purine neurotransmitter than ADP. 180

It is possible that if ADP is more potent than ADPR that the responses to ADPR might be more fully antagonized than responses to ADP. Therefore, we performed an additional series of experiments to test the effectiveness of MRS2500 in blocking responses over the full range of the concentration–response relationships of ADP and

ADPR. While maintaining single impalements, the pulse duration of spritzing was varied over the range of 10–100 ms and responses to ADPR and ADP were recorded before and after addition of MRS2500 (1 μM). Figure 4.7 shows that ADP and ADPR were approximately equally potent in producing hyperpolarization and that MRS2500 antagonized responses to ADPR to a much greater extent than to ADP.

Similar to previous results testing the effects of β-NAD+ (Mutafova-Yambolieva et al., 2007), ADPR also inhibited spontaneous contractions of colonic muscles cut parallel to the circular muscle fibres (Supplemental material, Fig. S4.1).

4.4.6 Electrophysiological evidence that degradation of β-NAD+ does not exclusively require CD38

We also tested whether postjunctional responses differ in colonic muscles isolated from wild-type and CD38−/− mice. In agreement with previous observations (Mutafova-

Yambolieva et al., 2007), robust IJPs were evoked by nerve stimulation in colonic muscles of wild-type mice, and the IJPs were blocked by apamin, suramin, PPADS and

MRS2500 (Fig. 4.8A–D and Table 4.2). Colonic muscles of wild-type mice were hyperpolarized by ADPR, and this response was also inhibited by apamin, suramin,

PPADS and MRS2500 (Fig. 4.8E–H and Table 4.2). Picospritzing of ADP also caused 181 hyperpolarization, and this response was significantly inhibited by apamin, but not by

MRS2500 (Fig. 4.8I and J and Table 4.2).

In muscles from CD38−/− mice, robust IJPs were evoked by nerve stimulation, and these responses were blocked by apamin and MRS2500 (Fig. 4.9A and B). There was no significant difference in IJPs or the pharmacology of IJPs in wild-type and CD38−/− mice.

Picospritzing β-NAD+ (Fig. 4.9C and D), ADPR (Fig. 4.9E and F), ATP (Fig. 4.9G and

H), or ADP (Fig. 4.9I and J) caused hyperpolarization, but the hyperpolarization responses were differentially blocked by apamin and MRS2500. For example, responses to β-NAD+ and ADPR were significantly reduced by apamin and MRS2500, but hyperpolarizations to ATP and ADP, although significantly reduced by apamin, were not affected by MRS2500 (Fig. 4.9 and Table 4.3). It appears, therefore, that the responses to endogenous neurotransmitter(s) and exogenous β-NAD+ are similar in colonic muscles from wild-type and CD38−/− mice and, therefore, pathways other than CD38 appear to be responsible for the degradation of β-NAD+ to ADPR. Moreover, the responses to locally applied β-NAD+ and ADPR, but not ATP and ADP, were attenuated by the P2Y1 receptor antagonist in colonic muscles from CD38−/− mice as shown for monkey WM and

CM (Fig. 4.6).

4.4.7 Mechanisms for ADPR degradation are present in the colon

As shown in Figs 4.6–4.9, localized application of ADPR caused hyperpolarization responses similar to the endogenous purine neurotransmitter. We investigated whether exogenous ADPR can be degraded by monkey and murine colonic muscles. Superfusion of monkey WM or CM tissues with eADPR resulted in formation 182 of eADO (Fig. 4.10A and B); eADO was 1.53 ± 0.24 (n = 10) and 3.53 ± 0.75 fmol (mg tissue)−1 (n = 19) in experiments with WM and CM, respectively. No eADO was resolved in superfusates in the absence of muscle tissues (P < 0.001 vs. superfusates exposed to muscles). Note that eADO formation was significantly greater per unit mass in CM preparations than in WM (P < 0.05). There was also significant formation of total products (eAMP + eADO) in WM (P < 0.05) and CM (P < 0.01), and total metabolite formation was greater per unit mass in CM than in WM, P < 0.05 (Fig. 4.10B).

In wild-type and CD38−/− colonic muscles, significant formation of eAMP and eADO occurred after exposure of eADPR to muscle tissues (Fig. 4.10C and D). In the absence of tissue the amount of eAMP was 2.79 ± 1.4 fmol (mg tissue)−1 (n = 27) and no eADO was resolved. eAMP formed was 15.36 ± 1.46 (n = 14, P < 0.001) and 12.74 ±

1.47 fmol (mg tissue)−1 (n = 13, P < 0.001) in wild-type and CD38−/− colons, respectively. eADO was 9.92 ± 0.96 and 9.57 ± 1.9 fmol (mg tissue)−1 in wild-type and

CD38−/−, respectively, P < 0.001 vs. no tissue control for both. The formation of total product (eAMP + eADO) was also significantly increased in the presence of wild-type and CD38−/− colons, respectively. Therefore, ADPR metabolism was similar in muscles of wild-type and CD38−/− mice.

183

Fig 4.1

Figure 4.1. Degradation of eNAD in monkey whole muscle (WM) and circular muscle (CM) colon preparations. A, original chromatograms of eNAD (0.2 μM) in the absence, (–) tissue, and presence, (+) tissue, of either WM or CM (30 s contact of substrate with tissue). The formation of eADPR and eADO was increased in the (+) tissue samples. No noticeable changes were observed in the peak of eNAD at 12.5 min, because this peak also contains eAMP formed from eNAD; LU, luminescence units. B, graphic representation of eADO formation in superfusate samples collected in the absence (–) or presence (+) of tissue. Note the increased formation of eADO in both WM and CM; the formation of eADO was greater in CM than WM. Asterisks denote significant differences from the amounts of eADO in (–) tissue samples (**P < 0.01, ***P < 0.001). There is a significant difference a significant increase in CM as compared to WM (†P < 0.05); number of experiments in parentheses. C, Western immunoblot analysis of CD38 showed no significant differences between the protein levels of CD38 in WM and CM. Density of each band is normalized to tubulin, which was used to control equal protein loading.

184

Fig 4.2

Figure 4.2. Degradation of eNAD in colon preparations isolated from wild-type and CD38−/− mice A, original chromatograms of eNAD (0.2 μM) in the absence, (–) tissue, and presence, (+) tissue, of wild-type and CD38−/− colons. Note the increase in eADO formation in the (+) tissue samples after 30 s contact of substrate with tissue. No noticeable changes were observed in the peak of eNAD at 12.5 min, because this peak also contains eAMP formed from eNAD. LU, luminescence units. B, graphic representation of eADO formation in superfusate samples collected in the absence (–) or presence (+) of tissue. eADO was formed in the presence of tissues. Note that the formation of eADO was comparable in preparations isolated from CD38+/+ and CD38−/− mice. Asterisks denote significant differences from the amounts of eADO in (–) tissue samples (**P < 0.01, ***P < 0.001); number of experiments in parentheses. C, genotyping confirms absence of CD38 from CD38−/− mice. The presence of CD38 (301 bp) was confirmed in the colons of wild- type controls but was absent in CD38−/− mice. The presence of CD38 was also confirmed in the brains of wild-type controls. RT control represents reverse transcriptase control and NTC represents non-template control.

185

Fig 4.3

Figure 4.3. Degradation of eNAD after brief contacts with murine colon A and C, original chromatograms of 0.2 μM eNAD in the absence of tissue, (–) tissue, and after 1 s and 5 s contact of eNAD with colon muscles isolated from wild-type mice (A) and colons isolated from CD38−/− mice (C). Note the appearance of the end product, eADO, in tissue superfusates after these brief exposures to eNAD. LU, luminescence units. B and D, graphic representation of eADO formation in superfusate samples collected in the absence (–) or presence (+) of tissue in colonic preparations isolated from wild-type (B) and CD38−/− (D) mice; number of experiments in parentheses. 186

Fig 4.4

Figure 4.4. Degradation of NGD in colon preparations isolated from wild-type and CD38−/− mice A, original chromatograms of 0.2 mM NGD in the absence (–) tissue and presence of wild-type and CD38−/− colons. Note the increase in cGDPR formation in the (+) tissue samples from colon preparations isolated from wild-type mice and lack of increase in cGDPR in colon preparations isolated from CD38−/− mice. LU, luminescence units. B, graphic representation of cGDPR formation in superfusate samples collected in the absence (–) or presence (+) of tissue. Asterisks denote significant differences from the amounts of cGDPR in (–) tissue samples (**P < 0.01); number of experiments in parentheses.

187

Fig 4.5

188

Figure 4.5. Degradation of eATP in colon preparations isolated from monkey and murine large intestine A, original chromatograms of eATP (0.05 μM) in the absence, (–) tissue, and presence, (+) tissue, of WM and CM of monkey colons. Note the increase in eADP, eAMP and eADO in the (+) tissue samples. LU, luminescence units. B, graphic representation of eADO formation in superfusate samples collected in the absence (–) or presence (+) of tissue. Asterisks denote significant differences vs. amounts in (–) tissue samples (**P < 0.01, ***P < 0.001); number of experiments in parentheses. C, original chromatograms of eATP (0.05 μM) in the absence, (–) tissue, and presence, (+) tissue, of colonic preparations isolated from wild-type and CD38−/− mice. eATP was decreased and eAMP and eADO were increased in both groups of preparations. LU, luminescence units. D, graphic representation of eADO formation in superfusate samples collected in the absence (–) or presence (+) of tissue. Asterisks denote significant differences from the amounts of eADO in (–) tissue samples (***P < 0.001). †Significant difference from CD38+/+ samples (P < 0.05); number of experiments in parentheses.

189

Fig 4.6

190

Figure 4.6. Inhibitory junction potentials and effects of ADPR and ADP on membrane potential of monkey colonic muscles A and B, electrical field stimulation (single pulse 0.5 ms duration; point of stimulation indicated by •), performed in the presence of atropine (1 μM) and l-NNA (100 μM) of monkey colonic circular muscles produced large inhibitory junction potentials (IJPs, control in both A and B) that were greatly attenuated or inhibited by apamin (0.3 μM; A) or by MRS2500 (1 μM; B). C and D, picospritzed ADPR (10 mM loaded in a spritz pipette, arrow, upper traces) produced robust and sustained membrane hyperpolarizations that were inhibited by apamin (0.3 μM) and MRS2500 (1 μM), respectively (lower traces in both panels). E and F, membrane hyperpolarizations induced by ADP (10 mM loaded in a spritz pipette, arrows upper traces) were also reduced by apamin (0.3 μM) and to a lesser extent by MRS2500 (1 μM; lower traces in each panel).

191

Fig 4.7

192

Figure 4.7. Concentration–response relationship for ADPR and ADP on membrane hyperpolarizations in murine colon A, a series of responses to spritzes of ADP (10 mM in spritz pipette) using pulse durations from 10–100 ms. Note the increase in hyperpolarization response as the spritz pulse is increased. B, responses to ADP after addition of MRS2500 (1 μM) to the bath solution. This P2Y1 antagonist only slightly decreased the area of the hyperpolarization responses to ADP. The fast voltage transients superimposed upon the record in B are due to static electricity. C, spritz responses to ADPR (10 mM in spritz pipette) using pulse durations from 10 to 100 ms. D, blockade of responses to ADPR after addition of MRS2500. The records in panels A–D were recorded during a single continuous impalement. E, summary of the results from 5 experiments using this protocol. Hyperpolarization responses were tabulated as areas under the response curves (mV s). *P < 0.01; **P < 0.001.

193

Fig 4.8

194

Figure 4.8. Effects of ADPR and ADP on membrane potential and inhibitory junction potentials of colonic circular muscle cells from wild-type CD38+/+ mice A–D show IJP evoked by EFS (single pulse 0.5 ms duration, •) in the presence of atropine (1 μM) and l-NNA (100 μM) (Control, upper traces in each panel) and after apamin (0.3 μM; A), suramin (100 μM; B), PPADS (30 μM; C) and MRS2500 (1 μM; D) (lower traces in each panel). Control responses were all recorded in the presence of atropine (1 μM) and l-NNA (100 μM). E–H shows the effects of ADPR (10 mM loaded in a spritz pipette, picospritzed onto circular muscles) at a time point indicated by arrow. Upper traces of each panel represent control responses recorded in the presence of atropine (1 μM) and l-NNA (100 μM). ADPR produced reproducible membrane hyperpolarizations which were sustained for several seconds before returning to pre-stimulus levels. ADPR-induced membrane hyperpolarizations were antagonized by apamin (0.3 μM; E), and the P2Y receptor antagonists, suramin (100 μM; F), PPADS (30 μM; G) and MRS2500 (1 μM; H). I and J shows the effects of ADP (10 mM, picospritzed) on murine colonic circular muscles before (upper traces) and in the presence of apamin (0.3 μM; I) and MRS2500 (1 μM; J). Apamin antagonized the membrane hyperpolarizations induced by ADP.

195

Fig 4.9

196

Figure 4.9. Effects of β-NAD+, ATP, ADPR and ADP on membrane potential and inhibitory junction potentials of colonic circular muscles from CD38−/− mice A and B, IJPs recorded from colonic circular muscles of CD38 null mice under control conditions (upper traces) and after apamin (A; lower trace) or MRS2500 (B; lower trace). Control conditions represent experiments that were performed in the presence of atropine (1 μM) and l-NNA (100 μM). C and D, β-NAD+ induced membrane hyperpolarizations (50 mM loaded in a spritz pipette, picospritzed at arrow) before (upper traces) and in the presence of apamin (0.3 μM; lower trace C) and MRS2500 (1 μM; lower trace D). E and F, membrane hyperpolarizations to ADPR (10 mM, picospritzed at arrow) before (upper traces) and in the presence of apamin (0.3 μM; lower trace E) or MRS2500 (1 μM; lower trace F). G and H, membrane hyperpolarizations in responses to picospritzed ATP (10 mM, upper traces in each panel) and in the presence of apamin (0.3 μM; lower trace G) or MRS2500 (1 μM; lower trace H). I and J, the effects of ADP on membrane potential under control conditions (10 mM, upper traces in each panel) and after apamin (0.3 μM; lower trace I) or MRS2500 (1 μM; lower trace J).

197

Fig 4.10

198

Figure 4.10. Degradation of eADPR in colon preparations isolated from monkey and murine large intestine A, original chromatograms of eADPR (0.05 μM) in the absence, (–) tissue, and presence, (+) tissue, of WM and CM of monkey colons. Note the decrease in eADPR and increase in eADO in the (+) tissue samples. LU, luminescence units. B, graphic representation of eAMP + eADO formation in superfusate samples collected in the absence (–) or presence (+) of tissue. Note that the formation of eAMP + eADO product was greater in CM than in WM. Asterisks denote significant differences from the amounts of eAMP + eADO in (–) tissue samples (*P < 0.05, **P < 0.01). †Significant difference from WM (P < 0.05); number of experiments in parentheses. C, original chromatograms of 0.05 μM eADPR in the absence, (–) tissue, and presence, (+) tissue, of colonic preparations isolated from wild-type and CD38−/− mice. eADPR was decreased and eAMP + eADO was increased in both groups of preparations. LU, luminescence units. D, graphic representation of eAMP + eADO formation in superfusate samples collected in the absence (–) or presence (+) of tissue. Asterisks denote significant differences from the amounts of eAMP + eADO in (–) tissue samples (***P < 0.001); number of experiments in parentheses.

199

Fig 4.11

Figure 4.11. Superposition of inhibitory junction potential (IJP) and hyperpolarization responses to exogenous ATP and β-NAD spritzed near the site of recording in a murine colonic preparation This experiment demonstrates the kinetic differences in responses to the endogenous purinergic neurotransmitter released from nerve terminals and hyperpolarization responses to exogenous transmitter candidates. Single pulses (0.5 ms pulse duration) of electrical field stimulation released neurotransmitter that resulted in fast inhibitory junction potentials (IJPs). The time constant of the upstroke of the IJPs, fitted by a single exponential (Clampfit; Molecular Devices, Sunnyvale, CA, USA), was 135 ± 8 ms (n = 41). In contrast responses to spritzed neurotransmitter candidates (10 psi; 25 ms pulses) developed more slowly (amplitudes scaled to approximate amplitude of the IJP); time constants for ATP (10 mM in spritz pipette) and β-NAD (50 mM in picospritz pipette) averaged 1217 ± 159 (n = 20) and 1001 ± 149 (n = 21), respectively. These data demonstrate that substantially more time is required for exogenous compounds to reach and bind receptors and for responses to develop than is required for the responses to neurotransmitters released from neurons. This implies that responses to neurotransmitters are transduced by postjunctional receptive fields quite close to the sites of release. These data also indicate that sites of neurotransmitter metabolism might be more accessible to transmitters released from neurons than to exogenous transmitter candidates. Thus, the kinetics of metabolism of exogenous substances may underestimate the kinetics of metabolism of endogenous transmitters by nearly an order of magnitude.

200

Table 4.1. Degradation of eADO (0.05 μM) by monkey WM, monkey CM and murine colon under control conditions or in the presence of either EHNA (10 μM), NBMPR (10 μM), or EHNA + NBMPR combined

(-) tissue (+) tissue, control (+) tissue, + EHNA (+) tissue, + (+) tissue, +EHNA NBMPR + NBMPR Monkey WM 60.0 ± 5.1 (10) 59.01 ± 10.9 (4) 63.51 ± 12.2 (2) 41.93 ± 9.1 (2) 57.15 ± 3.5 (2) Monkey CM 110.6 ± 4.9 (8) 108.87 ± 4.9 (2) 102.05 ± 7.4 (2) 61.79 ± 1.8 (2) 124.77 ± 5.5 (2) Mouse CD38+/+ 112.8 ± 13.9 (9) 104.2 ± 16.5 (7) – – 110.0 ± 34.7 (2)

Values are fmol (mg tissue)−1. No significant differences in (–) tissue versus (+) tissue, control for all groups (P > 0.05). No significant differences between (+) tissue, control and (+) tissue, + EHNA, or (+) tissue, + NBTI, or (+) tissue, + EHNA + NBTI for all groups (P > 0.05). Number of experiments in parentheses. 201

Table 4.2. Effects of antagonists to neural responses and exogenous ADPR and ADP in colons isolated from wild-type CD38+/+ mice.

IJP ADPR ADP % inhibition P value % inhibition P value % inhibition P value Apamin 65 ± 7 (9) 0.000001 90 ± 3 (5) 0.001987 71 ± 6 (4) 0.018600 Suramin 55 ± 6 (5) 0.000915 78 ± 2 (5) 0.000010 – – PPADS 65 ± 4 (5) 0.000134 73 ± 4 (5) 0.003722 – – MRS2500 93 ± 2 (12) 0.000001 96 ± 2 (6) 0.006596 30 ± 6 (6) 0.064000

202

Table 4.3. Effects of apamin and MRS2500 on neural responses and exogenous purines in colons isolated from CD38-/- mice

IJP β-NAD ADPR ATP ADP % inhibition P value % inhibition P value % inhibition P value % inhibition P value % inhibition P value Apamin 56 ± 3 (20) 0.000001 85 ± 5 (5) 0.00048 85 ± 4 (5) 0.0013 66 ± 9 (5) 0.0008 75 ± 8 (5) 0.013 MRS2500 99 ± 1 (21) 0.000001 82 ± 3 (6) 0.023 83 ± 4 (5) 0.0161 17 ± 7 (5) 0.3436 26 ± 4 (5) 0.2063

203

4.5 Discussion

In the present study we investigated tissue metabolism of purine neurotransmitter candidates in colonic muscles of monkeys and mice and postjunctional responses to direct metabolites. In confirmation of previous work, we found that colon muscle superfusates contained not only primary transmitter substances, β-NAD+ and ATP, but also their direct metabolites ADPR and ADP, respectively (Mutafova-Yambolieva et al.,

2007; Hwang et al., 2011). We provided evidence that ADPR and ADP are rapidly produced from β-NAD+ and ATP in colonic tunica muscularis and evaluated the role of

CD38 in β-NAD+ degradation. We tested whether ADPR and ADP mimic the effects of the endogenous purine neurotransmitter. In both monkey and murine colons, local application of ADPR and ADP evoked membrane hyperpolarization similar to β-NAD+ and ATP. However, only ADPR, like its precursor β-NAD+ (Mutafova-Yambolieva et al.,

2007; Hwang et al., 2011) mimicked the pharmacology of the endogenous purine neurotransmitter. These findings demonstrate that ADPR, either generated by metabolism of β-NAD+ released by enteric inhibitory neurons or released as a primary neurotransmitter, may contribute to postjunctional purinergic responses in GI muscles.

A component of nerve-evoked inhibitory responses in GI smooth muscles is blocked by antagonists of P2 purine receptors, indicating that a purine nucleotide, released from enteric motor neurons, mediates part of the inhibitory response in the GI tract. Since ATP is a common bioactive compound and a principle agonist of numerous

P2 receptors (reviewed in von Kugelgen, 2006), it has been assumed for many years that responses mediated by P2 receptors are elicited by ATP. In the context of GI physiology 204 this concept has amounted to ‘assigning' ATP to be the purine inhibitory neurotransmitter

(Burnstock et al., 1970; Burnstock, 2008b). However, ATP frequently failed to meet presynaptic criteria for a neurotransmitter substance or to mimic postjunctional responses elicited by the endogenous inhibitory neurotransmitter in GI muscles (Serio et al., 2003;

Mutafova-Yambolieva et al., 2007; Hwang et al., 2011). In contrast, β-NAD+, another endogenous purine nucleotide, is also released upon stimulation of enteric nerves and it mimics the effects and pharmacology of the inhibitory neurotransmitter in the murine and primate large intestine better than ATP (Mutafova-Yambolieva et al., 2007; Hwang et al.,

2011). These studies, although unable to formally rule out ATP as a neurotransmitter, raise the possibility that inhibitory purinergic neurotransmission is more complex than typically considered. The present study adds to the complexity by showing, that ADPR, the primary metabolite of extracellular β-NAD+, also mimics postjunctional responses to the endogenous inhibitory neurotransmitter.

Postjunctional electrophysiological responses (i.e. IJPs) attributed to release of purine neurotransmitter(s) are inhibited by P2Y1 receptor antagonists and by the small conductance K+ channel blocker apamin (Gallego et al., 2006; Gallego et al., 2008a;

Gallego et al., 2011; Mutafova-Yambolieva et al., 2007; Hwang et al., 2011). Similar to

IJPs, hyperpolarization responses to either β-NAD+ or ATP are inhibited by apamin

(Mutafova-Yambolieva et al., 2007; Hwang et al., 2011). β-NAD+ and ATP both bind to

P2Y1 receptors (von Kugelgen, 2006; Mutafova-Yambolieva et al., 2007; Kurahashi et al., 2011); however, only hyperpolarization responses to β-NAD+, and not ATP, were inhibited by P2Y1 receptor antagonists (Mutafova-Yambolieva et al., 2007; Hwang et al.,

2011). Therefore, in contrast to β-NAD+, the effects of ATP are mediated largely by 205 receptors in addition to P2Y1. With this observation, it is hard to imagine how ATP (or its metabolites) could be the purine neurotransmitter in these muscles if transmitters are released or metabolites are generated ‘in volume' within the interstitium. If this were the case, postjunctional responses (IJPs) would not be blocked by P2Y1 antagonists, just as responses to spritzed ATP and ADP are not blocked by P2Y1 antagonists. Our results, favour the concept that purine transmitters (or metabolites) are released (or generated) and bind receptors in a limited volume of the interstitium (e.g. in a synapse-like space), where P2Y1 receptors are the dominant receptors available for binding purines on postjunctional cells. In fact specialized cells, termed PDGFRα+ cells, have recently been described in colonic muscles. PDGFRα+ cells are closely associated with motor nerve terminals, form gap junctions with smooth muscle cells, and express the molecular apparatus to mediate P2Y1-dependent postjunctional responses (Kurahashi et al., 2011).

If purine transmitter(s), either by concentration or physical constraints, are limited to a receptive field, such as that provided by PDGFRα+ cells, then it is possible that both ATP and β-NAD+ could participate as co-transmitters. ADPR might be generated after release of β-NAD from enteric neurons, or ADPR could be stored and released as a primary neurotransmitter. It is also possible that ADP could be generated within junctional spaces and participate in P2Y1-dependent responses. At present we cannot clearly distinguish between these possibilities, but our data demonstrate that once released, β-NAD can be rapidly metabolized. Finding that ADPR is bioactive in GI muscles suggests that multiple purines could contribute to postjunctional purinergic responses in the gut.

We compared the degradation of extracellular β-NAD+, ADPR and ATP in whole tunica muscularis and in circular muscles (CM), containing nerve processes but no 206 ganglia, and we examined the role of CD38 in β-NAD+ metabolism. We first monitored metabolism of β-NAD+ and ATP in contact with muscles for 30–60 s to match neurotransmitter overflow studies performed with similar stimulation parameters (i.e.

Hwang et al., 2011; Mutafova-Yambolieva et al., 2007). Degradation of β-NAD+ was greater in monkey CM than in WM preparations. The increase in degradation of β-NAD+ in CM was not due to differences in ADO metabolism in WM versus CM as ADO did not undergo additional degradation during 60 s incubation with colonic tissues. Similar to β-

NAD+, ADPR was degraded more in CM than in WM of monkey colon. In contrast, degradation of ATP was similar in WM and CM preparations. Taken together these data suggest that the enzymatic activities responsible for the degradation of extracellular β-

NAD+ and ADPR are more prominent in parts of the colon wall containing nerve processes and varicosities than in parts containing myenteric ganglia. Thus, β-NAD+ and

ADPR may be degraded primarily near nerve terminals or close to the site of release.

ATP in contrast may be degraded at sites other than near sites of neurotransmitter release.

We found that a proportion of eNAD was degraded to eADO within a few seconds of contact with colonic muscles. One might argue on the basis of kinetics that 1 s or more is still slow compared to the kinetics of an IJP. However, this argument is too limited because metabolism of an exogenous neurotransmitter substance might have quite different kinetics than metabolism of neurotransmitters released from nerve terminals. In the former case, the exogenous neurotransmitter must first reach the site of metabolism, which might be very close to sites of transmitter release. It is not currently possible to simulate the delivery of endogenous neurotransmitter by application of an exogenous substance. Analogous to this, responses to exogenous neurotransmitter substances 207 develop with far slower kinetics than responses to neurotransmitters released from nerve terminals (Fig. 4.11). IJP hyperpolarization responses develop with a time constant that is nearly 10-fold less than the time constant of hyperpolarization responses to exogenous neurotransmitter (even applied locally with a spritz pipette). The same discrepancy might be expected in the kinetics of metabolism, and therefore, the breakdown in endogenous purines might occur 10-fold faster than metabolism of exogenous purines. It is likely that purinergic regulation of colonic muscles is the result of many IJPs, and previous studies have demonstrated tonic purinergic inhibition of colonic excitability (e.g. Spencer et al.,

1998a). Therefore, ample time exists for metabolism of purine neurotransmitters, and postjunctional responses are likely to be integrated responses to all primary transmitters and bioactive metabolites. The latter is also applicable to tissue pharmacology. Our results suggest that concentration–response studies of purines added to muscle baths are likely to be heavily contaminated by responses to bioactive metabolites. It should also be emphasized that quantitative analyses and comparisons of mechanisms and rates of degradation of purine nucleotides cannot be determined, at present, in whole tissue preparations due to the complexity of extracellular NAD+ metabolism (de Figueiredo et al., 2011), simultaneous processing by multiple enzymatic pathways, inability to simultaneously detect the formation of additional NAD metabolites (e.g. nicotinamide), and possible participation of other mechanisms in the elimination of β-NAD+ from the neuroeffector junction (e.g. neuronal uptake).

Since β-NAD+ meets pre- and postsynaptic criteria for a neurotransmitter in the colon better than ATP (Mutafova-Yambolieva et al., 2007; Hwang et al., 2011) and since the primary β-NAD+-metabolizing enzyme, CD38, is expressed in colon (Mutafova- 208

Yambolieva et al., 2007), we examined the functional role of CD38 in the degradation of extracellular β-NAD+ in the colon. As expected, the ADP-ribosyl cyclase activity, converting part of β-NAD+ into cADPR, is provided by CD38. However, NAD appeared to be degraded to ADO with equal efficiency in colonic muscles of wild-type and

CD38−/− mice. If CD38−/− colons lack the ADP-ribosyl cyclase activity they also probably lack the NAD-glycohydrolase activity that is associated with CD38, since the same active centre in the CD38 molecule carries both the ADP-ribosyl cyclase and NAD- glycohydrolase activities (Sauve et al., 1998). Therefore, enzymes other than CD38 are likely to be involved in the degradation of β-NAD+ in the colons isolated form CD38−/− mice. Moreover, amplitudes and durations of IJPs elicited by single stimuli were identical in colonic preparations isolated from wild-type and CD38−/− mice, in contrast to what would be expected if CD38 was the only enzyme degrading β-NAD+ released from motor neurons. If, on the other hand, ADPR contributed significantly to endogenous IJPs in the colon, reduced responses would be expected in CD38−/− mice, because ADPR would not be formed from β-NAD+ in the absence of CD38. Furthermore, responses to exogenous

β-NAD+ should be reduced in CD38−/− mice as ADPR would not be formed. However, our results show that both IJPs and hyperpolarization responses to exogenous β-NAD+ and ADPR were comparable in the presence and absence CD38. These are surprising observations, since CD38 is considered to be the main protein carrying NAD- glycohydrolase activity in mammals (Lee, 2001) and was established to be present in nerve terminals in vascular smooth muscles (Smyth et al., 2006a). The present study suggests, however, that additional enzymes with NAD-glycohydrolase activity are present in the GI muscles and might be upregulated to compensate for the CD38 absence 209 as can occur in animals with congenital gene deletions. For example, the GPI-anchored protein CD157 shares several characteristics with CD38, including a similar amino acid sequence and enzymatic functions involved in the metabolism of β-NAD+ (Ortolan et al.,

2002); no information is available about the potential role of CD157 in the degradation of

β-NAD+ in the colon. Further studies are warranted to determine the protein(s) responsible for the degradation of extracellular β-NAD+ in the colon.

In non-GI visceral smooth muscles (e.g. guinea-pig vas deferens), it has been reported that degradation of ATP increases during nerve stimulation due to ‘releasable’ enzymatic activity (Todorov et al., 1997). We investigated whether a similar mechanism for terminating transmitter activity exists in colonic muscles. Soluble (i.e. releasable) enzymatic activity was not observed in our experiments, as nerve stimulation did not enhance degradation of ATP and β-NAD+. This is in agreement with a previous study which also failed to detect soluble nucleotidase activity in enteric nerves of the guinea- pig taenia coli (Westfall et al., 2000), suggesting that soluble enzymatic activity is not generally a property of enteric motor neurons. Our data suggest that degradation of extracellular purines and purine neurotransmitter(s) released by nerve stimulation occurs via membrane-bound, ‘ecto-’, enzymes.

In summary, the present study demonstrates that: (i) bioactive metabolites of β-

NAD+ and ATP are rapidly formed in the tunica muscularis of the colon; (ii) ADPR, like its precursor β-NAD+, mimics the inhibitory purine neurotransmitter in the colon, (iii) more than a single purine substance may mediate purinergic neurotransmission in colonic muscles; (iv) degradation of extracellular β-NAD+ and ADPR is more pronounced in parts of the colon wall enriched in nerve terminals, and therefore, mechanisms exist to 210 degrade purine neurotransmitters and possibly limit their duration of action; (v) degradation of β-NAD+ appears to be only partially associated with CD38; degradation of

β-NAD+ seems to be accomplished by at least one additional pathway. Taken together, our data suggest that both primary neurotransmitters and metabolites are likely to contribute to purinergic signalling in GI muscles.

211

4.6 Supplementary material

Fig S4.1

Supplementary Figure S4.1. ADPR inhibited spontaneous contractions of colonic muscles cut parallel to the circular muscle fibers.

212

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Chapter 5

Differential release of β-NAD+ and ATP upon activation of enteric motor neurons in primate and murine colons

Leonie Durnin, Kenton M. Sanders and Violeta N. Mutafova-Yambolieva Published in Neurogastroenterology and Motility 2013 Mar;25(3):e194-204.

218

5.1 Abstract

Background: The purinergic component of enteric inhibitory neurotransmission is important for normal motility in the gastrointestinal (GI) tract. Controversies exist about the purine(s) responsible for inhibitory responses in GI muscles: ATP has been assumed to be the purinergic neurotransmitter released from enteric inhibitory motor neurons; however, recent studies demonstrate that β-nicotinamide adenine dinucleotide (β-NAD+) and ADP-ribose mimic the inhibitory neurotransmitter better than ATP in primate and murine colons. The study was designed to clarify the sources of purines in colons of

Cynomolgus monkeys and C57BL/6 mice. Methods: High-performance liquid chromatography with fluorescence detection was used to analyze purines released by stimulation of nicotinic acetylcholine receptors (nAChR) and serotonergic 5-HT3 receptors (5-HT3R), known to be present on cell bodies and dendrites of neurons within the myenteric plexus. Key Results: Nicotinic acetylcholine receptor or 5-HT3R agonists increased overflow of ATP and β-NAD+ from tunica muscularis of monkey and murine colon. The agonists did not release purines from circular muscles of monkey colon lacking myenteric ganglia. Agonist-evoked overflow of β-NAD+, but not ATP, was inhibited by tetrodotoxin (0.5 μmol L−1) or ω-conotoxin GVIA (50 nmol L−1), suggesting that β-NAD+ release requires nerve action potentials and junctional mechanisms known to be critical for neurotransmission. ATP was likely released from nerve cell bodies in myenteric ganglia and not from nerve terminals of motor neurons. Conclusions &

Inferences: These results support the conclusion that ATP is not a motor neurotransmitter 219 in the colon and are consistent with the hypothesis that β-NAD+, or its metabolites, serve as the purinergic inhibitory neurotransmitter.

5.2 Introduction

Gastrointestinal (GI) motility is regulated by enteric excitatory and inhibitory motor neurons which coordinate muscle contractions for propulsion of colonic contents

(Spencer & Smith, 2001). Muscles of the colon are under tonic neural inhibition (Lyster et al., 1995; Spencer et al., 1998b; Dickson et al., 2010a), and spontaneous inhibitory junction potentials (IJPs), mediated by release of neurogenic purines, contribute to tonic inhibition (Dickson et al., 2010b). Genetic deactivation of P2Y1 purine receptors delays transit through the colon (Hwang et al., 2012). Therefore, purinergic inhibitory neurotransmission is an important component in regulating colonic motility.

The identity of the purine(s) responsible for enteric inhibitory responses is controversial. ATP has been assumed to be the purine neurotransmitter (Burnstock et al.,

1970; Crist et al., 1992; Xue et al., 1999), but recent experiments on mouse (Mutafova-

Yambolieva et al., 2007) and primate colons (Hwang et al., 2011) showed that another purine, β-nicotinamide adenine dinucleotide (β-NAD+) and its bioactive metabolite, adenosine 5′-diphosphate ribose (ADPR) (Durnin et al., 2012b), mimic the endogenous purine neurotransmitter better than ATP. Measurements of purines released during stimulation of nerves by electrical field stimulation (EFS) demonstrated that both ATP and β-NAD+ are released in the colon, but these experiments were not capable of distinguishing the sites from which the two purines were released (Hwang et al., 2011). 220

Release of ATP and β-NAD+ from enteric neurons might occur from different sites by different mechanisms. For example, β-NAD+ (and ADPR) might be released from nerve varicosities at neuroeffector junctions, and ATP might be mainly released from extrajunctional sites, based on previous observations (Hwang et al., 2011). We tested this hypothesis by measuring purines released in response to stimulation of nicotinic acetylcholine receptors (nAChR) and serotonin (5-hydroxytryptamine) 5-HT3 receptors (5-HT3R) known to be expressed on cell bodies and dendrites of myenteric interneurons and muscle motor neurons (Dickson et al., 2010b; Kunze & Furness, 1999;

Steele et al., 1991; Zhou & Galligan, 1999; Galligan, 2002; Mazzia et al., 2003; Kapeller et al., 2011). For these stimuli to release neurotransmitters from nerve terminals of motor neurons, action potentials would need to develop and propagate to nerve varicosities, and

Ca2 + entry via Cacna1 family Ca2 + channels would need to occur in ‘active zones’ of varicosities to initiate vesicular fusion. Here, we demonstrate that the release of β-NAD+ in response to ganglionic stimulation is dependent upon these mechanisms, but release of

ATP is not. These findings are not consistent with ATP serving as a motor neurotransmitter in colonic muscles. Targeting β-NAD+ synthesis and/or metabolism may provide a novel rational for treating problems of colonic transit.

5.3 Methods

5.3.1 Tissue Preparation 221

Proximal colons of Cynomolgus monkeys (Macaca fascicularis) were obtained from Charles River Laboratories Preclinical Services (Reno, NV, USA). Monkeys, sedated with Ketamine (10 mg kg−1) and 0.7 mL Beuthanasia-D (Schering-Plough AH,

Kenilworth, NJ, USA), were exsanguinated [Charles River Laboratories Institutional

Animal Care and Use Committee (IACUC)] for reasons unrelated to this project.

C57BL/6 mice (Charles River Laboratories, Wilmington, MA, USA) were euthanized by sedation with isoflurane followed by cervical dislocation and exsanguination. The entire

GI tract was removed and placed in oxygenated cold Krebs solution for further dissection. All experimental procedures were approved by the IACUC at University of

Nevada.

Monkey proximal colon tunica muscularis (whole muscle, WM) tissues were prepared by dissecting away the mucosal layer. Monkey circular muscle (CM) tissues, containing only nerve terminals, were prepared by peeling away the longitudinal muscle with attached myenteric ganglia (Hwang et al., 2011). C57BL/6 mouse colons were prepared by removing the mucosa and submucosa.

5.3.2 Purine overflow

Colonic segments (40–70 mg) were placed in 200-μL superfusion chambers

(Hwang et al., 2011; Mutafova-Yambolieva et al., 2007) and superfused with oxygenated

−1 Krebs (37 °C; composition in mmol L ): 118.5 NaCl, 4.2 KCl, 1.2 MgCl2, 23.8

NaHCO3, 1.2 KH2PO4, 11.0 dextrose, 1.8 CaCl2 (pH 7.4). NG-nitro- l-arginine (l-NNA)

(100 μmol L−1) and atropine (1 μmol L−1) were present throughout. Superfusates were collected before and during stimulation of nAChR with epibatidine (500 μmol L−1, 30 s) 222

−1 −1 or DMPP (500 μmol L , 30 s) or 5-HT3R with SR57227 (500 μmol L , 30 s) and etheno-derivatized as described (Bobalova et al., 2002). Experiments were also

−1 performed with the nAChR antagonist, hexamethonium (500 μmol L ), or the 5-HT3R antagonist ondansetron (10 μmol L−1) for 30 min before stimulation with nAChR or 5-

HT3R agonists, respectively. In some experiments, tissues were superfused with tetrodotoxin (TTX, 0.5 μmol L−1) or ω-conotoxin GVIA (ω-Ctx GVIA, 50 nmol L−1) for

30 min before stimulation with nAChR or 5-HT3R agonists.

5.3.3HPLC assay of purines in tissue superfusates

A reverse-phased gradient Agilent Technologies 1200 liquid chromatography system equipped with a fluorescence detector (Agilent Technologies, Wilmington, DE,

USA) was used to detect 1,N6-etheno-derivatized nucleotides and nucleosides as described (Mutafova-Yambolieva et al., 2007; Hwang et al., 2011). Etheno-ATP is approximately 10-fold more fluorescent than the compound generated by etheno- derivatization of β-NAD+ (Breen et al., 2006). Areas of HPLC peaks were calculated and expressed in figures. Each peak was calibrated to individual etheno-derivatized purine standards. Results, normalized for sample volume and tissue weight, were expressed in femtomoles per milligram of tissue (fmol mg−1).

5.3.4 HPLC fraction analysis

1,N6-etheno derivatization forms 1,N6-etheno-ADPR by either β-NAD+, ADPR, or cyclic ADPR present in superfusates (Smyth et al., 2004). The compounds forming eADPR during nAChR or 5-HT3R activation were determined by HPLC fraction analysis 223

(Mutafova-Yambolieva et al., 2007; Hwang et al., 2011; Smyth et al., 2004).

Superfusates from 36 chambers were combined, concentrated, and analyzed by HPLC.

An Agilent Technologies 1200 Analytical Fraction Collector was employed to collect

400 μL fractions corresponding to retention times of cyclic ADPR (7.2-min fraction),

ADPR (8.5-min fraction), and β-NAD+ (10.5-min fraction). Fractions were subjected to etheno-derivatization and reanalyzed by HPLC for eADPR content.

5.3.5 Statistics

Data presented are means ± SEM. Means are compared by a two-tailed, unpaired t-test or by one-way anova for comparison of more than two groups followed by a post hoc Bonferroni multiple comparison test (GraphPadPrism, v. 3; GraphPad Software, Inc.,

San Diego, CA, USA). A probability value less than 0.05 was considered significant.

5.3.6 Drugs

(±)-exo-2-(6-Chloro-3-pyridinyl)-7-azabicyclo[2.2.1.]heptane (epibatidine) and 1-

(6-Chloro-2-pyridinyl)-4-piperidinamine hydrochloride (SR57227) were purchased from

Tocris Bioscience (Ellisville, MO, USA). Atropine, carbenoxolone, dimethylphenylpiperazinium (DMPP), hexamethonium bromide, l-NNA, ondansetron hydrochloride, and ω-Ctx GVIA were purchased from Sigma-Aldrich (St. Louis, MO,

USA). Tetrodotoxin was purchased from Ascent Scientific (Cambridge, MA, USA). All drugs were dissolved in deionized H2O, apart from epibatidine (dissolved in DMSO), and further diluted in perfusion solutions.

224

5.4 Results

5.4.1 Release of ATP and β-NAD+ elicited by activation of nAChR

We tested whether stimulation of nAChR evoked release of purines in monkey whole tunica muscularis (WM). As reported previously (Hwang et al., 2011), monkey

WM releases basal purines, including ATP, β-NAD+, and metabolites ADP, AMP, and adenosine (ADO) (Fig. 5.1A). Superfusion with nAChR agonist, epibatidine (Mandl &

Kiss, 2006) (500 μmol L−1 for 30 s), enhanced purine release (Fig. 5.1A,B). HPLC fraction analysis determined that the 11.2-min peak is composed of ∼92% β-NAD+, ∼5%

ADPR, and ∼3% cyclic ADPR; thus, β-NAD+ is the primary purine in the composite peaks. For simplicity, therefore, we refer to the purine eluted at 11.2 min as β-NAD+.

Hexamethonium (500 μmol L−1) inhibited the epibatidine-evoked release of ATP

(P < 0.05) and β-NAD+ (P < 0.001; Fig. 5.1B) verifying that the effect of epibatidine was mediated by nAChRs.

We tested whether release of purines in WM during nAChR activation is dependent upon nerve action potentials, which would be a requirement of release from motor neurons. The release of ATP in response to epibatidine was unaffected by TTX

(P > 0.05), but the release of β-NAD+ was significantly inhibited by TTX (P < 0.001)

(Fig. 5.1). These data suggest that release of β-NAD+ during activation of nAChR occurs at a site remote from the site of stimulation and requiring axonal action potentials for stimulus to couple to response. In contrast, the bulk of ATP released during activation of nAChR occurs from sites close to nicotinic receptors (e.g. nerve cell bodies). 225

We also tested the effects of an N-type voltage-dependent Ca2 + channel (VDCC) inhibitor, ω-Ctx GVIA, on purine release. ω-conotoxin GVIA significantly depressed epibatidine-evoked release of β-NAD+ (P < 0.001), but did not affect the release of ATP

(P > 0.05; Fig. 5.1). Therefore, release of β-NAD+ also required openings and influx of

Ca2 + through N-type Ca2 + channels, but this requirement was not present for release of

ATP during nAChR activation. Overflow of metabolites, AMP and ADO, followed the general changes in β-NAD+ overflow and release of total purines was significantly reduced by both TTX (P < 0.05) and ω-Ctx GVIA (P < 0.05), suggesting that most of the

AMP and ADO was due to metabolism of β-NAD+ (Fig. 5.1B) as previously suggested

(Hwang et al., 2011; Durnin et al., 2012b).

Dimethylphenylpiperazinium, another nAChR agonist, also evoked release of

ATP and β-NAD+ that was blocked by hexamethonium, but only the release of β-NAD+ was inhibited by TTX: release of β-NAD+ was reduced from 3.02 ± 1.1 fmol mg−1

(n = 12) to 0.50 ± 0.28 (n = 12, P < 0.05) and 0.54 ± 0.3 fmol mg−1 (n = 14, P < 0.05) in the presence of hexamethonium and TTX, respectively. The DMPP-evoked release of

ATP was reduced from 0.49 ± 0.16 (n = 12) to 0.10 ± 0.06 fmol mg−1 in the presence of hexamethonium (n = 12, P < 0.05). In the presence of TTX, the DMPP-evoked release of

ATP was 0.37 ± 0.09 (n = 14) which was not significantly different from control release

(P > 0.05).

There have been reports of presynaptic nAChR at nerve terminals of motor neurons regulating neurokinin release (Galligan, 2002; MacDermott et al., 1999).

Activation of these receptors might enhance Ca2 + entry at nerve terminals and facilitate transmitter release independent of axonal action potentials (Brain et al., 2001). We tested 226 the idea of presynaptic nAChR by stimulating CMs from monkey colon (i.e. muscles containing nerve terminals, but no cell bodies; see Hwang et al., 2011) with epibatidine.

In contrast to WM (Figs 5.1 and 5.2), epibatidine evoked no additional release of purines in pure CMs (Fig. 5.2), demonstrating that activation of prejunctional nAChR does not contribute to the epibatidine-mediated purine release in WM.

Finally, we examined the effects of epibatidine in mouse colon. Similar to monkey colon, epibatidine evoked release of ATP and β-NAD+ that was inhibited by hexamethonium (Fig. 5.3A,B). TTX significantly inhibited the epibatidine-evoked release of β-NAD+ (Fig. 5.3B), but did not affect ATP (Fig. 5.3A), suggesting that activation of nAChR in the mouse colon also evokes differential release of ATP and β-NAD+.

+ 5.4.2 Release of ATP and β-NAD elicited by activation of 5-HT3R

5-HT3Rs are also located on cell bodies of myenteric neurons and generate excitatory inputs to muscle motor neurons (Kapeller et al., 2011; Gershon, 2004).

Therefore, we also stimulated enteric neurons in monkey and murine colons with the

−1 selective 5-HT3R agonist, SR57227 (500 μmol L , 30 s) (Bachy et al., 1993). SR57227 stimulated release of ATP, β-NAD+, ADP, AMP, and ADO from monkey WM

−1 preparations (Fig. 5.4A,B). The 5-HT3R antagonist ondansetron (10 μmol L ) inhibited release of ATP and β-NAD+ evoked by SR57227 (P < 0.05; Fig. 5.4), confirming that the release was mediated by 5-HT3R. HPLC fraction analysis of tissue superfusates demonstrated that β-NAD+ was the dominant substance forming the 11.2-min 1,N6- etheno-ADPR peak (11.2 min fraction contained ∼83% β-NAD+, ∼15.5% ADPR, and

∼1.5% cyclic ADPR). Release of β-NAD+ evoked by SR57227 was inhibited by TTX 227 and ω-Ctx GVIA (P < 0.05 for both), but release of ATP was not affected (P > 0.05;

+ Fig. 5.4). Therefore, 5-HT3R-mediated release of β-NAD required axonal action potentials to convey information between sites of stimulation and transmitter release.

ATP release was unaffected by blocking Na+ channels and N-type VDCC. Release of

AMP, ADO, and total purines followed the same trend as β-NAD+, suggesting that most

AMP and ADO in superfusates were derived from released β-NAD+ than from ATP

(Fig. 5.4B).

In contrast to WM, SR57227 failed to evoke release of purines from monkey CM

(Fig. 5.5), indicating that SR57227-mediated release of ATP and β-NAD+ is not due to activation of receptors on nerve terminals within the CM.

As in monkey colon, SR57227 evoked release of β-NAD+ and ATP from mouse colons (Fig. 5.3C,D). TTX inhibited the SR57227-evoked release of β-NAD+ (P < 0.001;

Fig. 5.3B). The release of ATP showed a slight, but not significant, tendency of being reduced by TTX (P > 0.05; Fig. 5.3C).

We tested whether the release of ATP (or other purines) evoked by activation of

5-HT3R was dependent upon connexin or pannexin channels by exposing mouse colon muscles to carbenoxolone (100 μmol L−1 for 35–50 min) prior to stimulation with

SR57227. Carbenoxolone did not inhibit the release of ATP or β-NAD+; SR57227- evoked overflow of ATP was 1.91 ± 0.48 (n = 7) and 1.82 ± 0.57 fmol mg−1 (n = 3) before and in the presence of carbenoxolone, respectively (P > 0.05), and SR57227- evoked overflow of β-NAD+ was 5.32 ± 1.01 fmol mg−1 (n = 7) and 4.54 ± 0.82 fmol mg−1 (n = 3) before and in the presence of carbenoxolone, respectively (P > 0.05). In monkey colon, epibatidine-evoked overflow of ATP was 0.57 ± 0.08 (n = 22) and 228

0.46 ± 0.11 fmol mg−1 (n = 8) before and in the presence of carbenoxolone, respectively

(P > 0.05), and epibatidine-evoked overflow of β-NAD+ was 1.42 ± 0.3 (n = 22) and

0.97 ± 0.16 fmol mg−1 (n = 8) before and in the presence of carbenoxolone, respectively

(P > 0.05).

229

Fig 5.1

230

Figure 5.1. Stimulation of nicotinic acetylcholine receptors (nAChRs) causes release of purines in monkey colon whole muscle preparations. (A) Chromatograms of tissue superfusates collected before (control) and during stimulation of nAChRs with epibatidine (Epib, 500 μmol L−1, 30 s) in the absence and presence of hexamethonium (Hex, 500 μmol L−1 for 30 min), tetrodotoxin (TTX, 0.5 μmol L−1 for 30 min), or ω-conotoxin GVIA (ω-CtxG, 50 nmol L−1 for 30 min) in WM monkey colon. Small amounts of ATP, ADP, β-NAD+, AMP, and ADO were present in superfusates before stimulation, likely to cause tonic purinergic inhibition in colon. Epibatidine-evoked release of purines was inhibited by hexamethonium. Epibatidine-evoked release of β-NAD+, but not of ATP, was reduced by the neurotoxins TTX and ω-CtxG. Scale applies to all chromatograms. LU, luminescence units. (B) Averaged data are means ± SEM and summarize release of ATP, ADP, AMP, ADO, β-NAD+, and total purines (calculated as ATP+ADP+AMP+ADO+β-NAD+) during activation of nAChRs with epibatidine (Epib). Overflow (femtomoles per milligram of tissue) is the overflow during nAChR activation less spontaneous overflow. All purines were evaluated simultaneously in the same samples. Each peak was calibrated to individual etheno-derivatized purine standards. Asterisks denote significant differences from epibatidine-evoked release (*P < 0.05, **P < 0.01, ***P < 0.001); number of experiments in parenthesis.

231

Fig 5.2

Figure 5.2. Release of ATP and β-NAD+ during nicotinic acetylcholine receptor (nAChR) stimulation in whole muscle (WM) and circular muscle (CM) preparations of monkey colon. (A) Chromatograms of tissue superfusates collected before (control) and during nAChR stimulation with epibatidine (500 μmol L−1, 30 s) in WM and CM preparations of monkey colon. Small amounts of ATP and β-NAD+ were present in WM and CM superfusates in the absence of agonist. Epibatidine evoked release of ATP and β-NAD+ in WM preparations, but not in CM preparations. Scale applies to all chromatograms. LU, luminescence units. (B) Averaged data are means ± SEM, showing epibatidine-evoked release of ATP (left) and β-NAD+ (right) from monkey WM and CM preparations. Overflow (femtomoles per milligram of tissue) is the overflow during nAChR activation less spontaneous overflow. Overflow of ATP and β-NAD+ was significantly less in CM preparations. Asterisks denote significant differences from WM release (*P < 0.05, **P < 0.01); number of experiments in parenthesis.

232

Fig 5.3

A B

C D

Figure 5.3. ATP and β-NAD+ release during stimulation of nicotinic acetylcholine receptors (nAChRs) and 5-HT3Rs in mouse colon. A and B show release of (A) ATP and (B) β-NAD+ during activation of nAChRs with epibatidine (500 μmol L−1, 30 s) in the absence and presence of hexamethonium (Hex, 500 μmol L−1 for 30 min) or tetrodotoxin (TTX, 0.5 μmol L−1 for 30 min) in mouse colon. Averaged data are means ± SEM. Overflow (femtomoles per milligram of tissue) is the overflow during nAChR activation less spontaneous overflow. Activation of nAChRs with epibatidine evoked release of ATP and β- NAD+ that was inhibited by the nAChR antagonist hexamethonium. The epibatidine-evoked release of β-NAD+, but not ATP, was reduced by TTX. Asterisks denote significant differences from epibatidine-evoked release (*P < 0.05); number of experiments in parenthesis. C and D + show release of (C) ATP and (D) β-NAD during activation of 5-HT3 receptors with SR57227 (500 μmol L−1, 30 s) in the absence and presence of TTX (0.5 μmol L−1, 30 min) in mouse colon. Averaged data are means ± SEM. Overflow (femtomoles per milligram of tissue) is the overflow during 5-HT3 receptor activation less spontaneous overflow. The SR57227-evoked release of β- NAD+, but not ATP, was reduced by the neural blocker TTX (0.5 μmol L−1, 30-min perfusion). Asterisks denote significant differences from SR57227-evoked release (***P < 0.001); number of experiments in parenthesis.

233

Fig 5.4

234

Figure 5.4. Purines release by stimulation of 5-HT3 receptors in monkey colon whole muscle (WM) preparations. (A) Chromatograms of tissue superfusates collected before (control) and during activation of 5- −1 HT3 receptors with SR57227 (500 μmol L for 30 s) in the absence and presence of ondansetron (10 μmol L−1, 30 min), tetrodotoxin (TTX, 0.5 μmol L−1, 30-min superfusion), or ω-conotoxin GVIA (ω-CtxG, 50 nmol L−1, 30-min superfusion) in monkey WM. Small amounts of ATP, ADP, β-NAD+, AMP, and ADO were present in superfusate samples in the absence of agonist. Stimulation of 5-HT3 receptors with SR57227 evoked additional release of purines that was + inhibited by the 5-HT3 receptor antagonist ondansetron. SR57227-evoked release of β-NAD , but not ATP, was reduced by the neural blockers TTX and ω-CtxG. Scale applies to all chromatograms. LU, luminescence units. (B) Averaged data are means ± SEM and show release of ATP, ADP, AMP, ADO, β-NAD+, and total purines (calculated as ATP+ADP+AMP+ADO+β- + NAD ) during activation of 5-HT3 receptors with SR57227 (SR) and in the presence of ondansetron (Ond, 10 μmol L−1, 30 min), TTX (0.5 μmol L−1, 30 min), and ω-CtxG (50 nmol L−1, 30 min). Overflow (femtomoles per milligram of tissue) is the overflow during 5-HT3 receptor activation less spontaneous overflow. Each peak was calibrated to individual etheno-derivatized purine standards. Asterisks denote significant differences from SR57227-evoked release (i.e. control release) (*P < 0.05, **P < 0.01, ***P<.001); number of experiments in parenthesis.

235

Fig 5.5

+ Figure 5.5. Release of ATP and β-NAD during 5-HT3 receptor stimulation in whole muscle (WM) and circular muscle (CM) preparations of monkey colon. (A) Chromatograms of tissue superfusates collected before (control) and during 5-HT3 receptor activation with SR57227 (500 μmol L−1, 30 s) in WM and CM preparations of monkey colon. Small amounts of ATP and β-NAD+ were present in WM and CM superfusates in the absence of agonist. SR57227 evoked release of ATP and β-NAD+ in WM, but not in CM. Scale applies to all chromatograms. LU, luminescence units. (B) Averaged data are means ± SEM and show SR57227-evoked release of ATP (left) and β-NAD+ (right) from monkey WM and CM preparations. Overflow (femtomoles per milligram of tissue) is the overflow during 5-HT3 receptor activation less spontaneous overflow. Overflow of ATP and β-NAD+ was significantly less in CM preparations. Asterisks denote significant differences from WM release (*P < 0.05); number of experiments in parenthesis.

236

Fig 5.6

237

Figure 5.6. Sites of release of ATP and β-NAD+ in response to stimulation of nicotinic acetylcholine receptor (nAChR) and 5-HT3R in colonic muscles. ACh and 5HT released from descending interneurons in the myenteric plexus (MP) activate nAChR and 5-HT3R, respectively, on the cell bodies of inhibitory motor neurons and generates action potentials which propagate to nerve varicosities within muscle layers. β-nicotinamide adenine dinucleotide is released from nerve varicosities and serves as a primary enteric inhibitory motor neurotransmitter. ATP is released from sources (possibly nerve cell bodies) but not from nerve varicosities within muscle layers. LM, longitudinal muscle.

238

5.5 Discussion

We measured purines released in response to nAChR and 5-HT3R agonists. These receptors are localized on myenteric nerve cell bodies and dendritic projections of enteric neurons and mediate inputs to motor neurons from interneurons (Galligan, 2002). We reasoned that stimulation of motor neurons from ganglion targets might provide an opportunity to clarify the purine(s) involved in motor neurotransmission. Activation of

+ nAChRs and 5-HT3Rs evoked release of ATP and β-NAD in monkey and mouse colonic muscles, and release of β-NAD+, but not of ATP, was inhibited by the neurotoxins, TTX and ω-Ctx GVIA. It has been previously reported that purinergic neurotransmission in colon is blocked by TTX and ω-Ctx GVIA (Banks et al., 1979; Shuttleworth et al., 1997;

Rae et al., 1998; Gil et al., 2010; Bridgewater et al., 1995). Thus, enteric inhibitory neurotransmission attributable to purines requires nerve action potentials and Ca2 + entry via N-type Ca2 + channels, and the release of β-NAD+ is consistent with these properties of motor neurotransmission. In contrast, ATP release in response to ganglionic stimulants was not affected by TTX and ω-Ctx GVIA. Thus, it appears that ATP was released from ganglionic sources, possibly from the cell bodies of motor neurons. The source and mechanism of ATP was not investigated extensively because the goal of this study was not to elucidate the ganglionic sources of ATP, but to discriminate between sites of release for ATP and β-NAD+. Our findings make it very unlikely that ATP serves as a motor neurotransmitter in enteric inhibitory regulation of the colon and further clarify the role of β-NAD+ (Mutafova-Yambolieva et al., 2007; Hwang et al., 2011), or a metabolite

(Durnin et al., 2012b), as the purine motor neurotransmitter. 239

Previous studies have shown that the stimulation of ganglionic nAChR and 5-

HT3R elicits motor responses in GI muscles (Kadowaki et al., 1996; Borjesson et al.,

1997). Nicotinic acetylcholine receptor and 5-HT3R are ligand-gated ion channels on cell bodies of myenteric interneurons and motor neurons that mediate fast postsynaptic excitatory potentials (fEPSPs) and activation of action potential (Browning & Lees, 1996;

Obaid et al., 2005). Binding of appropriate ligands increases open probabilities of nAChR and 5-HT3R channels, facilitating entry of cations (inward current) that depolarizes neurons and initiates action potentials. Action potentials propagate down axons, depolarize nerve terminals, and activate N-type VDCC. Ca2 + entry into nerve terminals facilitates fusion of neuro-vesicles and transmitter release (Mandl & Kiss,

2006). Thus, neurotransmitter release initiated by somatodendritic ligand-gated ion channels would be expected to be inhibited by blockers of axonal action potentials and inhibitors of Ca2 + influx through VDCC at nerve terminals. Indeed, we found that the release of β-NAD+, but not ATP, satisfies these criteria for a motor neurotransmitter.

Thus, our findings support previous studies probing postjunctional mechanisms of purinergic motor neurotransmission and suggest that β-NAD+ is a primary purinergic neurotransmitter in the colon (as depicted in Fig. 5.6) (Mutafova-Yambolieva et al.,

2007; Hwang et al., 2011).

Acetylcholine binds to nAChRs on both excitatory and inhibitory motor neurons in the myenteric plexus (Browning & Lees, 1996; Obaid et al., 2005). Nicotinic acetylcholine receptor-mediated excitation of inhibitory motor neurons has also been demonstrated in human colon (Auli et al., 2008). We used epibatidine and DMPP in the present studies as ligands for nAChRs, and both agonists evoked release of ATP and β- 240

NAD+. Epibatidine in particular, is a potent and highly selective nAChR agonist (Mandl

& Kiss, 2006) that activates neurons, but not glia, within myenteric ganglia in the colon

(Gulbransen et al., 2010).

A subset of nAChRs has been reported at presynaptic nerve terminals where they modulate transmitter release (Mandl & Kiss, 2006; Wonnacott, 1997; Kirchgessner &

Liu, 1998). Direct stimulation of presynaptic nAChRs might cause transmitter release that is insensitive to TTX or ω-Ctx GVIA (Wonnacott, 1997). Therefore, we tested the effects of epibatidine on pure CM preparations from which the myenteric plexus was removed (Hwang et al., 2011). EFS has been shown to release β-NAD+ from CM

(Hwang et al., 2011), demonstrating that transmitter release mechanisms are intact in these muscles. Experiments on CM in the present study demonstrate that purine release is not coupled to prejunctional nAChRs.

Serotonergic descending interneurons release both ACh and 5-HT (Brookes,

2001; Galligan, 2002) and these interneurons appear to be important for exciting inhibitory motor neurons via nAChR and 5-HT3R and in producing tonic inhibition of

CM (Dickson et al., 2010b). 5-HT3Rs contribute to the inhibitory phase prior to propagation of a migrating motor complex. Antagonists of 5-HT3R and nAChR block spontaneous IJPs, suggesting that these receptors are expressed by inhibitory motor neurons that release purines (Dickson et al., 2010b). We examined this hypothesis directly in monkey and mouse colon by measuring purine release in response to a highly selective 5-HT3 receptor agonist, SR57227 (Bachy et al., 1993). SR57227 evoked release of β-NAD+, ATP, and their metabolites from monkey and mouse colonic WM, but failed 241

to release purines from CM preparations. Therefore, 5-HT3Rs are likely localized to the nerve cell bodies as shown by immunohistochemistry studies (Kapeller et al., 2011).

The assay techniques used in this study cannot determine the exact population of enteric neurons that release purines in response to nAChR and 5-HT3R agonists. nAChR and 5-HT3R are present on cell bodies of descending interneurons and inhibitory motor neurons in rodent and human colon (Dickson et al., 2010b; Mazzia et al., 2003; Kapeller et al., 2011; Auli et al., 2008; Kirchgessner & Liu, 1998; Torocsik et al., 1991).

Therefore, binding of these receptors could result in release of purines from either population of neurons. Interneurons release ATP as an excitatory neurotransmitter that can also mediate fEPSPs in motor neurons (Galligan et al., 2000), but if interneurons were the main population of neurons activated by nAChR and 5-HT3R, then ATP release would have been inhibited by TTX.

Nicotinic acetylcholine receptors are also expressed by a variety of non-neuronal cells (Wessler & Kirkpatrick, 2008). Therefore, nicotinic agonists might cause ATP release from non-neuronal sources. A non-neuronal source is unlikely for β-NAD+, because neurotoxins abolished its release. Just as the source of ATP is unknown, so is the mechanism responsible for ATP release in colon muscles. ATP can be released into the extracellular space via several mechanisms in addition to vesicle exocytosis, including connexin hemichannels, pannexin channels, ABC transporters, P2X7 receptor pores, and volume-regulated channels (Pankratov et al., 2006; Lazarowski, 2012; Lohman et al.,

2012; Mutafova-Yambolieva, 2012b). We tested carbenoxolone, a blocker of connexin/pannexin channels, but this compound did not block the release of ATP. Further 242 studies are needed to clarify the mechanisms of ATP release from nerve cell bodies or other cells in the GI tract.

In summary, activation of ligand-gated ion channels (nAChRs or 5-HT3R), expressed by myenteric neurons in mouse and monkey colon, evokes release of purine nucleotides. β-NAD+ release, but not ATP release, was blocked by neurotoxins, supporting the role of β-NAD+ as a primary enteric inhibitory motor neurotransmitter.

Our data suggest that ATP might function as a paracrine substance in ganglia, but does not function as a motor neurotransmitter in the colon. We suggest that further studies of

β-NAD+ synthesis, release, and metabolism may provide unique opportunities to exploit this pathway for therapeutic regulation of colonic motility.

243

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Chapter 6

Release, neuronal effects and removal of extracellular β-nicotinamide adenine dinucleotide (β-NAD+) in the rat brain

Leonie Durnin, Yanping Dai, Isamu Aiba, C. William Shuttleworth, Ilia A. Yamboliev and Violeta N. Mutafova-Yambolieva Published in European Journal of Neuroscience 2012 Feb;35(3):423-35.

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6.1 Abstract

Recent evidence supports an emerging role of β-nicotinamide adenine dinucleotide (β-NAD+) as a novel neurotransmitter and neuromodulator in the peripheral nervous system: β-NAD+ is released in nerve-smooth muscle preparations and adrenal chromaffin cells in a manner characteristic of a neurotransmitter. It is currently unclear whether this holds true for the central nervous system. Using a small-chamber superfusion assay and high-sensitivity HPLC techniques we demonstrate that high K+- stimulation of rat forebrain synaptosomes evokes overflow of β-NAD+, adenosine 5′- triphosphate (ATP), and their metabolites adenosine 5′-diphosphate (ADP), adenosine 5′- monophosphate (AMP), adenosine, ADP-ribose (ADPR), and cyclic ADPR. The high

K+-evoked overflow of β-NAD+ is attenuated by cleavage of SNAP-25 with botulinum neurotoxin A (BoNT/A), by inhibition of N-type voltage-dependent Ca2+ channels with

ω-conotxin GVIA, and by inhibition of the proton gradient of synaptic vesicles with bafilomycin A1, suggesting that β-NAD+ is likely released via vesicle exocytosis.

Western analysis demonstrates that CD38, a multifunctional protein that metabolizes β-

NAD+, is present on synaptosomal membranes and in the cytosol. Intact synaptosomes degrade β-NAD+. 1,N6-etheno-NAD, a fluorescent analogue of β-NAD+, is taken by synaptosomes and this uptake is attenuated by authentic β-NAD+, but not by the connexin

43 inhibitor Gap 27. In cortical neurons local applications of β-NAD+ cause rapid Ca2+ transients likely due to influx of extracellular Ca2+. Therefore, rat brain synaptosomes can actively release, degrade and uptake β-NAD+, and β-NAD+ can stimulate postsynaptic 250 neurons, all criteria needed for a substance to be considered a candidate neurotransmitter in the brain.

6.2 Introduction

Intracellular β-nicotinamide adenine dinucleotide (β-NAD+) serves as a coenzyme for cellular oxidation-reduction reactions, a donor of adenosine 5′-diphospho-ribose

(ADPR) in posttranslational modifications of proteins, and a precursor to cyclic ADPR

(cADPR) and other second messengers with Ca2+-releasing activity (Lee, 2001). In recent years β-NAD+ has also been recognized as an extracellular signaling molecule involved in cell-to-cell communication (Ziegler & Niere, 2004; Billington et al., 2006). In the peripheral nervous system β-NAD+ appears to serve as a novel neurotransmitter and neuromodulator (Smyth et al., 2004; Breen et al., 2006; Mutafova-Yambolieva et al.,

2007; Hwang et al., 2011). β-NAD+ is released from neurons in blood vessels, urinary bladder, and colon (Breen et al., 2006; Smyth et al., 2004; Smyth et al., 2006b;

Mutafova-Yambolieva et al., 2007; Hwang et al., 2011), and from nerve growth factor- differentiated rat PC12 cells (Yamboliev et al., 2009). Release of β-NAD+ correlates with neuronal activity and requires intact fast Na+ channels, N-type voltage-dependent Ca2+ channels, and the 25-kDa synaptosomal associated protein SNAP-25 (Smyth et al., 2004;

Smyth et al., 2006b; Breen et al., 2006; Mutafova-Yambolieva et al., 2007; Smyth et al.,

2009). β-NAD+ modulates the release of other neurotransmitters in blood vessels (Smyth et al., 2004), inhibits spontaneous smooth muscle contractions in urinary bladder (Breen et al., 2006) and has shown vasoactive effects in the mesenteric vasculature (Ralevic, 251

1995; Yamboliev et al., 2009). β-NAD+ also effectively causes membrane hyperpolarization and relaxation in the colon, consistent with an inhibitory neurotransmitter role in this tissue (Mutafova-Yambolieva et al., 2007; Hwang et al.,

2011). Therefore, evidence has been accumulating that β-NAD+ is a novel neurotransmitter in the peripheral nervous system and it remains to be determined whether this also holds true for the central nervous system.

In the brain β-NAD+ contributes to intracellular Ca2+ homeostasis, mitochondrial function, energy metabolism and complex processes such as aging and neuronal death

(Ying, 2007). In addition, β-NAD+ may participate in intercellular communication (De

Flora et al., 2004). For example, cultured hippocampal astrocytes release NAD+ and

+ 2+ 2+ respond to extracellular NAD with increased intracellular Ca ([Ca ]i) (Verderio et al.,

2001). Specific binding sites for NAD have been described and a role of NAD as a neuromodulator in rat and guinea-pig hippocampus has been suggested (Richards et al.,

1983; Snell et al., 1985; Galarreta et al., 1993). To the best of our knowledge, however, storage and release of β-NAD+ from central nervous system (CNS) neurons have not been reported. In the present study we have utilized high-sensitivity high pressure liquid chromatography (HPLC) methods to characterize candidate purine nucleotide transmitters released by depolarization of rat brain synaptosomes. We demonstrate that brain synaptosomes can actively release, degrade and uptake β-NAD+ and that exogenous application of β-NAD+ can generate robust Ca2+ transients in isolated cortical neurons.

Taken together, these findings suggest that β-NAD+ is a candidate neurotransmitter in the central nervous system. 252

6.3 Methods

6.3.1 Animals

For preparation of brain synaptosomes male Wistar-Kyoto or Wistar rats, 12–15 weeks of age, were euthanized by isoflurane inhalation. This method is approved by the

Institutional Animal Care and Use Committee (IACUC) of the University of Nevada.

After opening the scull the brain was quickly removed and placed in oxygenated Krebs- bicarbonate-HEPES (KBH) buffer (10°C) with the following composition

(mmol/L):NaCl 118, KCl 3.5, CaCl2 1.25, MgSO4 1.2, KH2PO4 1.2, NaCO3 25, HEPES-

NaOH 10, and D-glucose 11.5, pH 7.4 (Lonart & Sudhof, 2000). The forebrain was processed for preparation of synaptosomes as described below. For preparation of neuronal cultures, pregnant female Sprague–Dawley rats (16-day gestation) were obtained from Harlan Laboratories (Livermore, CA, USA). Primary neuronal cultures were established from day 16–17 embryos, as described below. The IACUC of the

University of New Mexico approved the experimental protocols involved in culture preparation.

6.3.2 Preparation of synaptosomes

Brain synaptosomes were prepared by differential centrifugation followed by

Percoll-gradient centrifugation as described in previous studies (Lonart & Sudhof, 2000;

Gualix et al., 2003; Gomez-Villafuertes et al., 2007; Wang, 2007) and shown in Figure

6.1. Brain tissue was homogenized in brain homogenizing buffer (mmol/L): HEPES 10, sucrose 320, EDTA 2, EGTA 2 (Gomez-Villafuertes et al., 2007; Wang, 2007) using a 253 glass-teflon homogenizer (0.25 mm clearance). Homogenate was then centrifuged for 5 min at 1,000 × g and 4°C to obtain pellet P1 (nuclei and cell debris) and supernatant S1

(containing synaptosomes). P1 was rewashed with homogenizing buffer, then centrifuged at 1,000 × g for 5 min and supernatant was combined with S1. Pooled S1 fraction was centrifuged at 17,000 × g and 4°C for 10 min to obtain P2 (synaptosome-rich pellet). P2 was resuspended in Percoll gradient buffer, prepared by dissolving 0.25 mmol/L dithiothreitol in brain homogenizing buffer (Gomez-Villafuertes et al., 2007; Wang,

2007). Resuspended P2 was applied onto a gradient of 23%, 10%, and 3% Percoll.

Gradient tubes were centrifuged at 25,000 × g for 10 min and 4°C, and synaptosomes were recovered from the 10%/23% Percoll interface. To remove residual Percoll, synaptosomes were diluted with 10 volumes of aerated (95% O2, 5% CO2) ice-cold KBH buffer and centrifuged at 20,000 × g for 5 min at 4°C. Synaptosome-rich pellet P3 was resuspended on ice in KBH buffer and aliquots of the P3 synaptosomal suspension were used for experiments as described below. In some experiments P3 suspension was centrifuged at 20,000 × g for 5 min and synaptosomes in pellet P4 were disrupted by placing in ice-cold hypotonic solution for 30 min. Supernatant S5 (intrasynaptosomal content) was obtained by a 5-min centrifugation at 20,000 × g and used to measure intrasynaptosomal concentrations of purines and catecholamines and uptake of 1,N6- etheno-NAD (eNAD).

6.3.3 HPLC assay of etheno-purines

Endogenous purines (e.g., ATP, ADP, AMP, ADO, β-NAD+, ADPR, and cADPR) in superfusates were measured after derivatization to 1,N6-etheno-derivatives to 254 increase detection sensitivity as described previously (Levitt et al., 1984; Bobalova et al.,

2002). One hundred μl of a citrate phosphate buffer (pH 4.0) was added to 200 μl of superfusate sample and purines were etheno-derivatized by adding 10 μl of 2- chloroacetaldehyde for 40 min at 80°C. Endogenous ATP, ADP, AMP, and ADO were derivatized to the 1,N6-derivatives eATP, eADP, eAMP, and eADO, respectively

(Bobalova et al., 2002). β-NAD+, ADPR and cADPR are all derivatized to eADPR

(Smyth et al., 2004). Etheno-purines were assayed by HPLC with fluorescence detection

(HPLC-FLD) as described previously (Bobalova et al., 2002). Agilent 1100 and 1200 liquid chromatography module systems (Agilent Technologies, Wilmington, DE) were used throughout this study. The mobile phase consisted of 0.1 mol/L KH2PO4 (pH 6.0) as eluent A; eluent B contained of 35 % methanol and 65 % eluent A. Gradient elution was employed according to the following linear program: time 0, 0 % eluent B; 18 min, 100

% eluent B at a flow rate of 1 ml/min and run time 20 min. Chromatography signals were recorded at Ex of 230 nm and Em of 420 nm. The amounts of purines in each sample were calculated from calibration curves of purine standards run simultaneously with every set of experimental samples. Results were normalized for sample volume and amount of protein loaded. The overflow of purines was expressed in nmol/mg protein.

6.3.4 Intrasynaptosomal contents of purines

One hundred μl of intrasynaptosomal fraction (described in Preparation of synaptosomes) were derivatized to 1,N6-etheno-purines and etheno purines were assayed by HPLC-FLD (Bobalova et al., 2002; Smyth et al., 2004).

255

6.3.5 Spontaneous secretion of purines by synaptosomes

Contents of purines were measured in samples collected after placing P3 synaptosomal suspension in a small-chamber superfusion system (described in High K+- evoked overflow) and superfused with KBH solution as described below. After 15-minute equilibration eight hundred μl of superfusate were collected in each protocol. Samples were analyzed for purines by a HPLC-FLD technique.

6.3.6 High K+-evoked overflow

Synaptosomes (600–800 μg protein) were applied under gentle suction onto

Whatman GF/B filter paper (3 mm diameter) which was placed in a 0.45 μm Cameo 3N syringe filter serving as a perfusion chamber as described previously (Yamboliev et al.,

2009). The chamber was perfused with a KBH solution (37°C) at 0.8 ml/min. After equilibration, 800 μl of superfusate were collected as pre-stimulation samples containing spontaneously released substances. Synaptosomes were then stimulated with KCl (25 mmol/L or 65 mmol/L). The high K+-solutions were prepared by substituting NaCl for equal amounts of KCl to maintain isotonicity. Eight hundred μl samples of superfusate were collected at 1, 2, 5 or 10 minutes of superfusion in ice-cold Eppendorf tubes. In some experiments synaptosomes were incubated with 300 nmol/L BoNT/A for 2 hours at

37°C and then high K+ overflow experiments were performed. In some experiments synaptosomes were superfused for 30 min with either bafilomycin A1 (0.1 μmol/L) or ω- conotoxin GVIA (ω-Ctx GVIA, 20–50 nmol/L) prior to treatment with a high-K+ (25 mmol/L) KBH solution.

256

6.3.7 HPLC fraction analysis

To identify the ratio of β-NAD+:ADPR:cADPR forming eADPR at 11.2 min we carried out an HPLC fraction analysis (Smyth et al., 2004) on samples from the intrasynaptosomal compartment as well as samples collected before and during high-K+ stimulation. Briefly, in overflow experiments 2.4-ml synaptosome superfusate samples were collected in tubes and immersed in liquid N2. Control samples from KBH or high

K+ solutions (no contact with synaptosomes) were also collected. The samples were concentrated by Speed Vacuum (Savant SVC100, Thermo Electron Corp., Westmont, IL) to 1 ml. Nine hundred μl of each concentrated sample were injected into the HPLC system and 400 μl-fractions were collected according to retention time (cADPR 7.0–7.4 min, ADPR 8.3–8.7 min, and β-NAD+ 10.3–10.7 min). Fractions were subjected to etheno-derivatization with 2-chloroacetaldehyde and were analyzed for eADPR content by HPLC-FLD.

6.3.8 Degradation of β-NAD+ and ATP

Figure 6.2 depicts enzymatic pathways for the degradation of β-NAD+ and ATP.

Six hundred μg of synaptosomes were loaded in a small-chamber superfusion system and superfused with KBH solution (see High K+-evoked overflow). Following equilibration, chambers were superfused with substrates eATP (50 nmol/L), eNAD (0.2 μmol/L) or

NGD (0.2 mmol/L) in KBH solution (Bobalova & Mutafova-Yambolieva, 2003; Smyth et al., 2006a) to assay ATP-ase, NAD-glycohydrolase, and ADP-ribosyl cyclase activities, respectively. Eight hundred μl-samples from substrate solution (no synaptosomes present, S1), and from superfusate after 1-min contact of synaptosomes 257

with substrate (S2) were collected in ice-cold tubes and immersed in liquid N2. Purines in samples were assayed by HPLC-FLD and enzymatic activities were determined by disappearance of substrate and appearance of products. Each purine was quantified against known standards and data were normalized for protein amount. Derivatized purines were detected at Ex of 230 nm and Em of 420 nm, whereas the non-modified compounds (i.e., NGD and cGDPR) were detected at Ex of 270 nm and Em of 400 nm

(Bobalova et al., 2002). cGDPR standard was prepared by incubation of NGD with

Aplysia californica as described previously (Smyth et al., 2006a).

6.3.9 Uptake of NAD

P3 synaptosomes were incubated with eNAD (1 mmol/L) at 37°C for 1, 5 or 60 minutes to measure the uptake of NAD. The reactions were stopped by inserting the tubes on ice and the tubes were centrifuged at 22,000 × g for 5 minutes at 4°C to produce synaptosome-containing pellet. The pellet fraction was then washed twice in 0.5–1 ml ice-cold KBH buffer and synaptosomes were hypotonically disrupted for 30 min on ice.

Suspension was then centrifuged again at 22,000 × g for 5 min at 4°C to produce supernatant (S5) as intrasynaptosomal fraction. Intrasynaptosomal samples were processed for purine detection by HPLC-FLD. Appearance of eNAD and its metabolites eADPR and eADO in intrasynaptosomal fractions was used as a measure of eNAD uptake. In some experiments synaptosomes were incubated with a mixture of eNAD (1 mmol/L) and β-NAD+ (10 mmol/L) to examine whether authentic β-NAD+ competes with eNAD for uptake mechanisms. In some experiments synaptosomes were pre- 258 incubated with Gap 27 (100 μmol/L) to examine whether connexin hemichannels are involved in the uptake of eNAD.

6.3.10 Western immunoblot analysis

Equal amounts of total protein from synaptosomal extracts or intrasynaptosomal fractions were resolved by SDS-PAGE and transferred onto nitrocellulose membranes for

1.5 hours at 24V and 4°C. Membranes were blocked for 1 hour with LI-COR blocking buffer (LI-COR, Inc., Lincoln, NE) and probed with the following primary antibodies diluted in LI-COR buffer: anti-synaptophysin (mouse monoclonal, Chemicon, dilution

1:1000), anti-chromogranin B (rabbit polyclonal, Abcam, dilution 1:300), anti-CD38

(goat polyclonal, Santa Cruz Biotechnology, dilution 1:200), and anti-SNAP25 (mouse monoclonal, Sternberger, dilution 1:1000). After removal of excess primary antibody, membranes were incubated for 45 min at room temperature with secondary antibodies coupled to infrared fluorescence markers with Em of 800 nm (IR800, Rockland

Immunochemicals, PA) or 680 nm (Alexa Fluor 680, Molecular Probes, OR) diluted

1:100,000 in LI-COR buffer. Images were obtained with an infrared Odyssey scanner

(LI-COR, Inc., Lincoln, NE).

6.3.11 Cell culture

Primary neuronal cultures were established from 16–17 day old rat embryos as described (Paul et al., 2003). Briefly, the cortex with the adjoining striatum were dissected out, dissociated mechanically and re-suspended in Dulbecco’s minimum essential medium/F12 (1:1) supplemented with 5% fetal calf serum. Cells (8×106 cells/60 259 mm dish) were plated on poly-d-lysine-coated tissue coverslips and grown for 12–14 days in vitro (DIV) at 37°C in a humidified atmosphere consisting of 5% CO2. To inhibit proliferation of non-neuronal cells, 10 μM cytosine d-arabinofuranoside was added to the cultures 72 h after plating. Thereafter neurons were maintained in Minimal Essential

Medium containing 5% fetal calf serum until the day of experiment.

6.3.12 Imaging of intracellular Ca2+ transients

Cytosolic Ca2+ levels were assessed using the high-affinity ratiometric indicator

Fura-2 as described previously (Tafoya et al., 2008). Cultures were loaded at room temperature with 3 μM Fura-AM for 20 min in HEPES buffer (mmol/L): 130 NaCl, 5

KCl, 2 CaCl2, 1 MgCl2, 11 Glucose, 10 HEPES, pH 7.6 and then rinsed for 20 min in

HEPES to allow for indicator de-esterification. Coverslips were then transferred to the recording chamber and superfused with HEPES (2 ml/min) at room temperature. β-

NAD+, ADPR or ATP (50 mmol/L loaded in picospritz pipettes) were applied locally using pressure ejection (500 ms, 40 psi, Picrospritzer II, General Valve Corp.) from a conventional patch recording electrode (5MΩ). This method allows rapid and reproducible ejections of very small volumes while avoiding the inherent desensitization of nerve cells which accompanies other methodologies (i.e., bath application or superfusion). Due to the very small volume of ejections (calculated at fractions of a nanoliter in the present study) the final concentration at neurons is likely very much lower than the agonist concentration loaded in the picospritz pipette. In some experiments the recording chamber was superfused with α,β-methylene-ATP (α,β-MeATP, 50

μmol/L) for 10 minutes prior to local application of β-NAD+. Cultures were allowed to 260 equilibrate for 20 min before recording began. Fura-2 excitation was achieved using

350/380 nm excitation pairs (40 ms each) delivered from a monochromator (TiLL

Photonics GmbH, Grafeling, Germany) via a 40×WI objective (Olympus, N.A. 0.8).

Fluorescence emission (510 nm) was detected using an interline transfer cooled CCD

(TiLL Imago). Image pairs were background-subtracted and then ratioed (Till Vision v

4.0). Uncertainties regarding final intracellular concentrations following loading of the

AM form of Fura-2 and Kd in the intracellular compartment limits reliable conversions to

Ca2+ concentrations, and measurements of Ca2+ are expressed as ratios of background- subtracted ratios of the 350/380nm excited fluorescence. Responses in each cell were compared with depolarization responses to 40 mmol/L KCl-containing buffer, obtained at the end of each experiment.

6.3.13 Statistics

Data are presented as means ± SE. One-way ANOVA with a post hoc

Bonferroni’s multiple comparison test was performed using GraphPadPrism v. 3 for

Windows (GraphPad Software, San Diego, CA) when three or more groups of data were compared. Paired or unpaired two-tailed Student’s t-test was performed using the same software when two groups of data were compared. A probability value of less than 0.05 was considered significant.

6.3.14 Drugs

ATP, ADP, α,βMeATP, AMP, ADO, ADPR, β-NAD+, NGD, and ω-conotoxin

GVIA were purchased form Sigma-Aldrich (St. Louis, MO). eNAD and cADPR came 261 from Biolog (Bremen, Germany). Bafilomycin A1 and Gap 27 came from Tocris

Bioscience (Ellisville, MO). BoNT/A was purchased from List Biological Laboratories

(Campbell, CA).

6.4 Results

6.4.1 Intra-synaptosomal content of purines and spontaneous release of endogenous purines from rat brain synaptosomes

To determine substances that have the potential to be released from synaptosomes upon membrane depolarization, we first assayed the intrasynaptosomal content of endogenous purines (Table 6.1). Note that the absolute amounts of individual purines might have been underestimated due to sample dilution and possible in vitro degradation during sample preparation. However, the use of identical methodologies to simultaneously measure intrasynaptosomal and extrasynaptosomal contents of multiple endogenous purine nucleotides allowed comparisons of the amounts of purines potentially available for release and purines released at rest and upon cell membrane depolarization. The amounts of purines spontaneously released from synaptosomes superfused with normal KBH buffer (5 mmol/L KCl) are shown in Table 6.1; approximately 0.2–0.5% of the intrasynaptosomal content of purines appeared to be released spontaneously from isolated synaptosomes.

An HPLC fraction analysis demonstrated that the β-NAD++ADPR+cADPR cocktail in the intra-synaptosomal compartment comprised about 71% β-NAD+, 20%

ADPR, and 9% cADPR. Identical analysis of the eADPR peak in superfusate samples 262 collected from unchallenged synaptosomes revealed 66.45±3.04% β-NAD+,

29.67±3.86% ADPR and 3.91±2.18% cADPR, n=7. Since β-NAD+ was the primary purine nucleotide in the mixture, the amounts of β-NAD++ADPR+cADPR were calculated based on β-NAD+ standards.

6.4.2 High-K+-evoked release of purines

Regulated vesicular release following membrane depolarization would be consistent with a neurotransmitter role of β-NAD+. We therefore examined whether depolarization with high K+ concentrations evoked additional release of purine in the superfusate. Superfusion with 25 mmol/L or 65 mmol/L KCl caused additional release of purines including β-NAD+. The release of most purines (e.g., β-NAD+, AMP and adenosine) appeared to be concentration-dependent (Fig. 6.3). Twenty five mmol/L KCl produced the highest overflow of ATP at 5 min (P<0.05, t=3.52 vs. 0 min control), whereas the overflow of β-NAD++ADPR+cADPR was significantly enhanced at 2 min

(P<0.05, t=3.488), 5 min (P<0.05, t=3.297), and 10 min (P<0.05, t=3.742), F=5.103 after stimulation with KCl. Likewise, the overflow of total purines was significantly increased at 5 min (P<0.05, t=3.153) and 10 min (P<0.05, t=3.637), F=4.431. The overflow of

ADP, AMP and ADO did not reach statistical significance at any time point. At 65 mmol/L KCl the overflow of AMP and ADO was increased at 5 min (P<0.05, t=3.026,

F=2.952 for AMP and P<0.05, t=3.048, F=2.883 for ADO). The overflow of β-

NAD++ADPR+cADPR and total purines was enhanced at 2 min (P<0.05, t= 2.954,

F=2.94 for β-NAD++ADPR+cADPR and P<0.05, t=3.128, F=4.132 for total purines) and

5 min (P<0.05, t=2.974, F=2.94 for β-NAD++ADPR+cADPR and P<0.05, t=3.625, 263

F=4.132 for total purines) of stimulation. The overflow of ATP and ADP did not reach statistical significance from 0 time; one-way analysis of variance and Bonferroni’s multiple comparison post hoc test. Time control experiments showed a fairly consistent evoked release of purines within the first two hours after isolation of synaptosomes.

Samples collected during 2-min stimulation with 65 mmol/L KCl contained 41.02±11.21 pmol/mg protein β-NAD+, 28.69±10.12 pmol/mg protein ADPR and 0.495±0.17 pmol/mg protein cADPR, which represents 62.72±3.38%, 36.42±3.52% and

0.862±0.317% of the pool, respectively, n=7.

We next examined whether disruption of synaptic vesicle release machinery affects spontaneous and K+ depolarization-evoked release of purines. Treatment with clostridium botulinum neurotoxin A (BoNT/A, 300 nmol/L), an inhibitor of the SNARE protein SNAP-25 (Blasi et al., 1993a), significantly attenuated the additional release of purines evoked by high-K+ solution (25 mmol/L) (Fig. 6.4B,E): the overflow of β-

NAD++ADPR+cADPR was increased in controls at 2 min (P<0.05, t=2.919) and 5 min

(P<0.05, t=3.275), F=4.365 after KCl stimulation, whereas in the BoNT/A-treated preparations the overflow of β-NAD++ADPR+cADPR remained unchanged by KCl

(P>0.05). Total purines were also increased in controls at 2 min (P<0.05, t=2.769) and 5 min (P<0.05, t=2.820), F=3.556 after KCl stimulation, but were not significantly increased in BoNT/A-treated preparations (P>0.05). Likewise bafilomycin A1 (0.1

μmol/L for 30 min), a drug blocking vesicular uptake of neurotransmitters by inhibiting membrane H+-ATPase (Moriyama & Futai, 1990) or ω-Ctx GVIA (0.02–0.05 μmol/L for

30 min), a specific inhibitor of N-type (CaV2.2) neuronal voltage-gated Ca2+ channels

(Olivera et al., 1984), attenuated the high K+ (25 mmol/L)-evoked release of β-NAD+ 264

(Fig. 6.4C,D,E), at 1, 2, and 5 min of stimulation P>0.05 vs. 0 min. The changes in total purine overflow followed closely the changes in β-NAD+ overflow (Fig. 6.4F).

6.4.3 Degradation of ATP and NAD

To evaluate whether enzymatic activities for termination of neurotransmitter action are present in isolated synaptosomes, synaptosomes were either superfused or incubated with etheno-derivatized purines as substrates (Jamal et al., 1988; Todorov et al., 1997; Bobalova & Mutafova-Yambolieva, 2003). Use of etheno-purines ensures

~1,000,000 times higher sensitivity (Bobalova et al., 2002) and allows for the detection of small changes in substrate or product concentrations. Importantly, spontaneously released endogenous purines collected in the superfusates were not subjected to derivatization prior to HPLC analysis and hence their fluorescence remained subthreshold. Superfusion of synaptosomes with eATP caused a decrease in levels of the substrate eATP and an increased levels of product eADP+eAMP+eADO (Fig. 6.5A,

P=0.0071, t=6.586, df=3, paired t test, two-tailed), suggesting the presence of enzymes such as E-NTPDases, NPPs and ecto-5′-nucleotidase that degrade ATP, ADP and AMP, respectively (see Fig. 6.2). Likewise, superfusion (Fig. 6.5B) or incubation (Fig. 6.6A) of synaptosomes with eNAD led to an increase of the eNAD metabolites eADPR and eADO

(P=0.0177, t=4.753, df=3, paired t test, two-tailed), indicating the presence of NAD- glycohydrolase, NPPs, and 5′-nucleotidase on synaptosome membranes. ADP-ribosyl cyclase was also present and responsible for the increased production of cGDPR from

NGD (Fig. 6.5C, P=0.0022, t=9.870, df=3, paired t test, two-tailed). NGD was used as a substrate instead of β-NAD+, because ADP-ribosyl cyclase also converts NGD into 265 cGDPR which is the end product of this reaction, whereas cADPR formed from β-NAD+ by ADP-ribosyl cyclase can be further processed to ADPR and other products (Graeff et al., 1994).

6.4.4 Expression of CD38 and synapse-associated proteins

We next examined whether CD38, a protein involved in the degradation of β-

NAD+, is expressed on intact synaptosomes and in intra-synaptosomal fractions. Western analysis demonstrated CD38 expression on synaptosomes, P3 (Fig. 6.6B) and in the intra- synaptosomal compartment, S5 (Fig. 6.6B), which also contained the markers for synaptic vesicles synaptophysin and chromogranin B (Fischer-Colbrie et al., 1987; Jahn

& Sudhof, 1994).

6.4.5 Uptake of NAD

To evaluate whether extracellular β-NAD+ can be taken up in brain synaptosomes, we incubated synaptosomes with 1 mmol/L eNAD, a highly fluorescent analogue of β-

NAD+. eNAD is not an endogenous substance and hence, is not present in the intrasynaptosomal compartment under control conditions (Fig. 6.7A,B 0′). Thus under these conditions, any appearance of eNAD in the intrasynaptosomal compartment after incubation of synaptosomes with eNAD would indicate eNAD uptake. One-, 5- and 60- minute incubation with eNAD led to the appearance of eNAD in the intrasynaptosomal compartment (Fig. 6.7A). The eNAD products eADPR and eADO also appeared intrasynaptosomally. The appearance of eNAD in the intrasynaptosomal compartment was not affected by incubation of synaptosomes with the connexin 43 hemichannel 266 inhibitor Gap 27, 100 μmol/L, (Fig. 6.7B, C), but was significantly reduced by incubation of synaptosomes with 10 mmol/L β-NAD+ (Fig. 6.7E, P=0.046, t=4.48, df=2, paired t test, two tailed). Likewise, the appearance of eNAD+eADPR+eADO in the intrasynaptosomal compartment remained unchanged by Gap 27 (Fig. 6.7D), but was reduced by authentic β-NAD+ (Fig. 6.7F, P=0.0177, t=7.144, df=2, paired t test, two- tailed). Incubation of synaptosomes with eADPR (1 mmol/L, 60 min) caused the appearance of 5.13±1.04 nmol/mg protein (n=2) eADPR, suggesting that some ADPR that is formed in the synaptic cleft can also be taken up by synaptosomes.

6.4.6 Neuronal Ca2+dynamics

To determine whether β-NAD+ affects target neurons, the effects of locally

+ 2+ applied β-NAD were examined on cultured rat cortical neurons. [Ca ]i levels were assessed in neurons loaded with the high-affinity Ca2+ indicator Fura-2. Localized

+ 2+ application of β-NAD caused a rapid, concentration-dependent [Ca ]i transients (Fig.

6.8A, Movie in Online Supplemental materials, http://onlinelibrary.wiley.com/doi/10.1111/j.1460-9568.2011.07957.x/suppinfo). β-NAD+

(50 mmol/L loaded in the picospritz pipette) produced ratio changes of 0.344±0.027

(n=9) which equated to a 59.1±7.7% increase in Ca2+ relative to responses to 40 mmol/L

KCl applied at the end of each experiment (Fig. 6.7D; n=9 cultures). Almost all neurons responded to β-NAD+. On average 93.2% of neurons (identified by both morphology and

2+ + sustained [Ca ]i elevations to KCl) located in front of the β-NAD application electrode

2+ responded with a stimulus-locked [Ca ]i transient (9 cultures, 148 total neurons). Control studies with identical localized pressure applications (500 ms) of either 1) solutions of 267 matched osmolarity (NaCl) or 2) low pH HEPES buffer (pH 3) showed no change in

2+ 2+ [Ca ]i levels (Figs. S6.1 and S6.2), indicating that the Ca transients were not triggered by activation of mechanosensitive channels or pH changes, but by the action of β-NAD+.

The rapid time course of Ca2+ transients suggested that transmembrane Ca2+ influx might be involved. To test this hypothesis, neurons were challenged with β-NAD+ in Ca2+-free buffer. Reproducible β-NAD+ responses were first generated in normal buffer (containing

2 mmol/L Ca2+) at 10 min intervals and then, after 10 min superfusion in nominally Ca2+- free buffer, responses to β-NAD+ were abolished (Fig. 6.8B, Movie in Online

Supplemental materials, http://onlinelibrary.wiley.com/doi/10.1111/j.1460-

9568.2011.07957.x/suppinfo). A partial recovery was observed after reintroduction of

Ca2+-containing solution (not shown). These results are consistent with β-NAD+ stimulation of Ca2+ influx, rather than mobilization of Ca2+ from intracellular stores.

The approach of localized application of millimolar concentrations of β-NAD+ has been utilized in other preparations, to mimic aspects of accumulation at sites of transmitter release (Mutafova-Yambolieva et al., 2007; Hwang et al., 2011). We next sought to determine whether localized application of another purine nucleotide, ATP, elicits comparable responses in rat brain neurons. As shown in Fig. 6.9A similar Ca2+ transients were observed with ATP challenges, using identical pipette concentrations and delivery pulses. Although individual neurons could respond preferentially to either ATP or β-NAD+, the population responses demonstrated rapid onset Ca2+ transients with no significant difference in initial rise times. A full pharmacological characterization of the receptor type(s) involved in β-NAD+ responses is outside the scope of the present study.

However, it is noted that activation of some P2X receptors is unlikely to explain the 268 postsynaptic Ca2+ increases observed here. Thus, bath exposures to αβ-MeATP at concentrations sufficient to fully desensitize P2X receptors (Evans et al., 1992) reduced but did not abolish β-NAD+ responses (Fig. 6.9B). Finally, we determined whether the direct metabolite of β-NAD+, ADPR (Lee, 2001; De Flora et al., 2004), could also stimulate postsynaptic neurons. As shown in Fig. 6.9C, robust Ca2+ transients were generated by localized ADPR applications. Like β-NAD+ responses, signals recovered quickly to baseline levels after termination of the stimulus, and the average peak amplitude of transients (Fura-2 ratio changes of 0.35±0.018; n=4 cultures), was very similar to those reported above for β-NAD+ (0.34±0.027; n=9 cultures). The rates of initial increase of Ca2+ signals in response to ADPR and β-NAD+ in Fig. 6.9C were 95.66

± 16.57 and 122.14 ± 14.19 (percent of peak response/second), respectively (P=0.167, t=1.616, df=5, paired t test, two tailed).

269

Fig 6.1

Figure 6.1. Protocol for isolation of synaptosomes from rat forebrain Different velocities of cold ultracentrifugation and Percoll gradient centrifugation were used to obtain intact synaptosomes (P3). In some experiments synaptosomal membranes were hypotonically disrupted and supernatant (S5) was used as intrasynaptosomal content as described in Methods.

270

Fig 6.2

Figure 6.2. Diagram of major enzymatic pathways for the degradation of β-NAD+ and ATP CD38 exhibits NAD glycohydrolase (NADase), ADP-ribosyl cyclase and cADPR hydrolase activities and degrades β-NAD+ to ADPR and cADPR. ADPR is degraded to AMP by nucleotide pyrophosphatases (NPPs). AMP in turn is degraded to ADO by 5′-nucleotidase. ATP is degraded by a family of ecto-nucleoside triphosphate diphosphohydrolases (ENTPDases) to ADP and AMP, and then to ADO by 5′-nucleotidases.

271

Fig 6.3

Figure 6.3. Spontaneous release and high K+-evoked release of adenine purines KCl 25 mmol/L (closed circles, n=7) or 65 mmol/L (open circles, n=13) evoked overflow of ATP (A), ADP (B), AMP (C), ADO (D), and β-NAD+ADPR+cADPR (E) in rat isolated forebrain synaptosomes. The sum of all purines (i.e., ATP+ADP+AMP+β-NAD++ADPR+cADPR+ADO) is also shown (F). Data are averaged for each individual purine as well as for the sum of all purines (total purines) and are expressed as the mean ± SE. *P<0.05 vs. controls at 0 min. One- way ANOVA followed by post-hoc Bonferroni’s multiple comparison tests. Note the sustained pattern of the overflow of β-NAD+ and total purines.

272

Fig 6.4

273

Figure 6.4. Effects of botulinum neurotoxin A (BoNT/A), bafilomycin A1 and ω-conotoxin GVIA (ω-Ctx GVIA) on spontaneous and high-K+-evoked release of adenine purines (A-D) Original chromatograms of superfusates from synaptosomes pretreated with either KBH solution (controls, n=13), BoNT/A (300 nmol/L for 2 h, n=6), bafilomycin (0.1 μmol/L for 30 min, n=8) or ω-Ctx GVIA (20–50 nmol/L for 30 min, n=6) and superfused with 25 mmol/L K+ for 5 minutes. Superfusate samples were collected at 0, 1, 2, and 5 minute of superfusion with high K+-solution. The peak eluting at ~11.5 minute is formed by β-NAD++ADPR+cADPR; this peak is labeled as β-NAD+ to reflect the primary compound in the β-NAD++ADPR+cADPR mixture. Scales apply to each groups of chromatograms. LU, luminescence units. (E, F) Data are averaged for β-NAD+ and total purines and are expressed as the mean ± SE. In contrast to controls the high K+ solution failed to evoke an increased release of purine nucleotides in the presence of BoNT/A, bafilomycin A1 or ω-CtxGVIA. *P<0.05 vs. controls at 0 minutes. One- way ANOVA followed by post-hoc Bonferroni’s multiple comparison tests. 274

Fig 6.5

275

Figure 6.5. Degradation of exogenous purine substrates during superfusion of synaptosomes (A) Ecto-ATPase activity. Left panel: Original chromatograms of 50 nmol/L eATP in the absence of synaptosomes (−s) and after 1-minute contact with synaptosomes (+s); LU, luminescence units. Note the decrease in eATP and the increase in eADP, eAMP, and eADO in the (+s) samples. Right panel: Graphic representation of e-product (eADP+eAMP+eADO) formation in superfusate samples collected before (−s) and during 1-min perfusion of synaptosomes with eATP (+s). (B) NAD glycohydrolase activity. Left panel: Original chromatograms of 0.2 μmol/L eNAD in the absence (−s) and presence of synaptosomes (+s). Note the decrease in eNAD and the increase in eADPR formation in the (+s) samples. Right panel: Graphic representation of eADPR formation in superfusate samples collected in the absence (−s) or presence (+s) of synaptosomes. (C) ADP-ribosyl cyclase activity. Left panel: Original chromatograms of 0.2 mmol/L NGD in the absence (−s) and presence of synaptosomes (+s). Note the increase in cGDPR formation in the (+s)-samples. Right panel: Graphic representation of cGDPR formation in superfusate samples collected in the absence (−s) or presence (+s) of synaptosomes. *P<0.05, **P<0.01. Data represent mean ± SE from 4 experiments with each substrate, paired Student’s t- test. 276

Fig 6.6

277

Figure 6.6. Degradation of eNAD and expression of CD38 (A) NAD glycohydrolase activity determined by a 60-minute incubation of synaptosomes with 1 mmol/L eNAD. Original chromatograms of eNAD in the absence (−s) and presence of synaptosomes (+s); LU, luminescence units. Note the increased formation of eADPR and eADO in contact with synaptosomes. (B) Western analysis showed expression of CD38 (45 kDa) in intact synaptosomes and in the intrasynaptosomal compartment. Intrasynaptosomal compartment also expressed the synaptic vesicle markers synaptophysin (42 kDa) and chromogranin B (78 kDa). 278

Fig 6.7

279

Figure 6.7. Uptake of eNAD Original chromatograms of intra-synaptosomal content of eNAD after 0, 1, 5, and 60-minute incubation of intact synaptosomes with 1 mmol/L eNAD alone (A) and in the presence of Gap 27 (100 μmol/L, n=2) (B). Incubation with eNAD caused appearance of eNAD and its metabolites eADPR and eADO in the intra-synaptosomal compartment. Gap 27 had no effect on the intrasynaptosomal levels of eNAD or eNAD++eADPR+eADO (C, D). Simultaneous incubation of β-NAD+ (10 mmol/L) and eNAD (1 mmol/L) abolished the appearance of eNAD in the intrasynaptosomal compartment and reduced the intrasynaptosomal levels of eNAD+eADPR+eADO after 60-minute incubation, n=3 (E, F). Data represent mean ± SE, P<0.05, unpaired Student’s t-test. 280

Fig 6.8

281

Figure 6.8. Intracellular Ca2+ increases in a cultured neurons following localized applications of β-NAD+ (A) Rapid Ca2+ increases in 6 neurons identified by number in the culture illustrated in the inset. β-NAD+ was applied via a brief (500 ms) pressure pulse from a microelectrode (E). Two consecutive responses are shown (2 min interstimulus interval), showing that responses in individual neurons are reproducible. (B) After 10 min of exposure to Ca2+-free buffer, responses to the same β-NAD+ stimuli were abolished. (C) For comparison with β-NAD+ responses, Ca2+ elevations are shown in the same neurons after restoration of extracellular Ca2+, and bath application of 40 mmol/L KCl. Scale bar: 30 μm. 282

Fig 6.9

283

2+ Figure 6.9. Similar neuronal [Ca ]i transients with ADPR and ATP 2+ + (A) [Ca ]i increases in the same neurons, following sequential applications of β-NAD and ATP. The delivery micropipette was changed between β-NAD+ and ATP trials, during a 10 min interval indicated by the axis break. (B) Desensitization of P2X receptors by pre-exposure to αβ- methylene ATP (50 μmol/L, 10 min) reduced, but did not abolish β-NAD+ responses. (C) Comparison of responses to β-NAD+ and its metabolite ADPR in the same neurons. As in A, the delivery micropipette was changed between challenges with the two agonists. Identical pipette concentrations (i.e., 50 mmol/L) and delivery pulses (500 ms) were used for β-NAD+, ATP and ADPR.

284

Table 6.1. Intra-synaptosomal content and spontaneous overflow of purines in rat brain synaptosomes (pmol/mg protein).

285

6.5 Discussion

Of major importance in this study are the findings that β-NAD+, a ubiquitous intracellular constituent, can be released by brain synaptosomes upon membrane depolarization via vesicle exocytosis and can also directly stimulate cortical neurons. β-

NAD+ was previously suggested as a novel neurotransmitter in the peripheral nervous system (i.e., Hwang et al., 2011; Mutafova-Yambolieva et al., 2007) and was shown to be released by cultured astrocytes (Verderio et al., 2001), but it has never been considered as a candidate for a neurotransmitter in brain nerve terminals. In the present study the high-K+-evoked release of β-NAD+ in isolated forebrain synaptosomes required intact synaptic vesicle exocytosis machinery. Mechanisms for removal of extracellular β-

NAD+, including enzymatic degradation and neuronal uptake, were also present.

Localized application of β-NAD+ produced Ca2+ transients in cortical neurons, suggesting that when released from presynaptic neurons, endogenous β-NAD+ could activate postsynaptic neurons. Taken together, these observations suggest that β-NAD+ qualifies as a candidate neurotransmitter in the brain.

Several lines of evidence in the present study support the notion that, like classical neurotransmitters, β-NAD+ is released upon membrane depolarization via vesicle exocytosis. Several lines of evidence in the present study support this notion: 1) membrane depolarization with high-K+ solutions caused overflow of β-NAD+ from viable synaptosomes used within 4–6 hours of fractionation (Gomez-Villafuertes et al., 2007),

2) the amount of released β-NAD+ appeared proportional to the membrane depolarization, 3) the high-K+-evoked release of β-NAD+ was abolished by inhibition of 286 neuronal N-type voltage-dependent Ca2+ channels, 4) the evoked release of β-NAD+ was inhibited after disruption of the proton gradient in synaptic vesicles, and 5) cleavage of the SNARE protein SNAP-25 inhibited the high-K+-evoked release of β-NAD+. Taken together these observations suggest that vesicle exocytosis was the primary mechanism of

β-NAD+ release in response to depolarization of the synaptosomal membrane.

After release of a neurotransmitter, mechanisms for recovery or local degradation of the substance are needed to terminate neural signals. β-NAD+ is the primary purine nucleotide in the β-NAD++ADPR+cADPR mixture in synaptosomal superfusates, but yet, a significant formation of ADPR also occurred. Therefore, membrane-bound β-

NAD+-degrading enzyme(s) are likely present on synaptosomal membranes. In agreement with previous studies in brain tissue (White, 1977; Sperlagh et al., 1998) we found that ATP was released upon high-K+-stimulation. The samples collected before and during stimulation also contained the direct ATP metabolite ADP, as well as AMP and adenosine as previously shown (Zimmermann, 1996). As depicted in Fig. 6.2, AMP and adenosine are degradation products of both β-NAD+ and ATP. Because the changes in total purines followed the changes in the β-NAD+ overflow better than those of the ATP overflow, it can be assumed that β-NAD+ is the primary source of AMP and adenosine in brain synaptosomes. Also, two approaches for evaluation of enzyme activities (i.e., small- chamber superfusion and at-rest-incubation) demonstrated that mechanisms for local degradation of β-NAD+ are available in brain synaptosomes. Finally, CD38, a type II integral membrane glycoprotein that exhibits ADP-ribosyl cyclase, cADPR hydrolase and

NAD glycohydrolase activities (Graeff et al., 1998a; Lee, 2001; De Flora et al., 2004) was present on synaptosomal membranes and in the intrasynaptosomal compartment. 287

This is in agreement with previous observations showing that CD38 is expressed on plasma membranes and cell organelles of neurons and astrocytes in rat brain (Yamada et al., 1997). The catalytic site of membrane-bound CD38 faces the extracellular space (Lee et al., 1993; Munshi et al., 2000), making this enzyme suitable as a regulator of extracellular β-NAD+ levels (Billington et al., 2006). Therefore, once released β-NAD+ can quickly be converted into ADPR and cADPR in the extracellular space.

Efficient control of synaptic transmission and rapid termination of the action of neurotransmitters may be achieved not only by the degradation of the neurotransmitter substances but also by their uptake into neural cells by specific transporters (reviewed in

Amara & Kuhar, 1993). Neurotransmitter uptake may also be an essential component in the recycling of neurotransmitters released during neural activity. To our knowledge the possibility that β-NAD+ can be imported across the plasma membrane to replenish intracellular or vesicle β-NAD+ pools in synaptosomes has not been addressed. However, information in other systems suggests that this may occur. For example, high concentrations of extracellularly applied NAD increase intracellular NAD levels in

NIH3T3 fibroblasts, cultured murine astrocytes, human neuroblastoma SH-SY5Y, and

HeLA cell types (Bruzzone et al., 2001b; Ying et al., 2003;Billington et al., 2008).

NADH, a close “relative” of β-NAD+, can be transported across the plasma membranes of astrocytes by a P2X7R-mediated mechanism (Lu et al., 2007). Bidirectional NAD transport in isolated connexin 43-expressing murine 3T3 fibroblasts has also been reported (Bruzzone et al., 2001a). In the present study we demonstrate that incubation of synaptosomes with the fluorescent analogue eNAD caused appearance of this substance in the intra-synaptosomal compartment, which otherwise does not contain eNAD. 288

Simultaneous incubation with 10-fold higher concentrations of authentic β-NAD+ inhibited the appearance of eNAD in the intracellular compartment, suggesting that β-

NAD and eNAD compete for the same transport mechanism(s). Inhibition of connexin 43 with Gap 27 did not significantly affect intrasynaptosomal uptake of β-NAD+, consistent with the notion that other transport mechanisms may be involved in rat brain synaptosomes. Defining the exact nature of β-NAD+ transporters warrants further studies.

It is also possible that some amount of the β-NAD+ metabolites ADPR and ADO can be taken up directly through the synaptosomal membrane.

We demonstrated that, if released, exogenous β-NAD+ could affect neuronal functions. A subpopulation of CNS neurons responded to β-NAD+ with rapid Ca2+ transients, suggesting that extracellular β-NAD+ could indeed participate in neuron-to- neuron communication. Picospritzing β-NAD+ close to target neurons, instead of superfusing the substance, was meant to simulate rapid direct application and avoid some of the factors that might dampen responses. Concentrations of β-NAD+ in synaptic vesicles or in the synaptic cleft have not been reported. However the concentrations of β-

NAD+ applied with picospritz pipettes were likely comparable to concentrations of neurotransmitters achieved near postsynaptic neurons following exocytosis, since in analogy with ATP and other neurotransmitters (Van der Kloot, 2003; Borycz et al., 2005;

Fields & Burnstock, 2006; Zimmermann, 2008) the concentration of β-NAD+ in synaptic vesicles could likely reach up to 150–1000 mmol/L. By analogy glutamate concentrations within the synaptic cleft rise to millimolar levels when glutamate is released from a presynaptic vesicle (Clements et al., 1992). Thus, localized application of very small volumes and rapid dilution of millimolar concentrations of exogenous neurotransmitter 289 substances appears to be a reasonable approach to determine whether exogenous β-NAD+ applications can indeed cause significant postsynaptic activation of central neurons. As shown in the present study equimolar concentrations of ATP produced Ca2+ responses with very similar amplitudes as observed for β-NAD in the same cell population. β-NAD+ failed to elicit Ca2+ transients in the absence of extracellular Ca2+ implying that the rapid neuronal responses are due to influx of Ca2+ from the extracellular space. These results differ from a recent report in rat cultured neurons (Fischer et al., 2009) which has demonstrated more modest Ca2+ increases following much longer exposures (10 seconds) to lower P2 receptor agonist concentrations (i.e., pressure application of 100 μmol/L

ATP). In that prior study, only metabotropic responses were detected, and it is likely that the robust Ca2+ influx responses observed in the present study with greater local concentrations of β-NAD+ or ATP were missed. It is also possible that during the longer period of ATP delivery some degradation of ATP might have occurred. Therefore, some of the observed effects in this prior study may in fact be due to bioactive ATP metabolites

(e.g. ADP).

Localized application of ADPR, a purine nucleotide that could be formed by the

NAD-glycohydrolase-mediated catabolism of β-NAD+, produced Ca2+ transients with similar rates of onset in neurons that also responded to β-NAD+. ADPR might be generated after release of β-NAD+ from brain neurons, or ADPR could be stored and released as a primary neurotransmitter. At present we cannot clearly distinguish between these possibilities, however our data demonstrate that both β-NAD+ and ADPR are bioactive in cortical neurons and suggests that multiple purines could contribute to postsynaptic responses in brain neurons. The dynamics of the responses to β-NAD+ and 290

ADPR appear comparable and β-NAD+ and ADPR likely function in concert in brain neurons. α,β-Me-ATP-sensitive receptors (i.e., P2X1 and P2X3 types, (North, 2002)) appeared to be only partially responsible for the Ca2+ transients in response to locally applied β-NAD+. Future studies should determine the mechanisms underlying the evoked influx of extracellular Ca2+ including possible activation of ionotropic receptors, non- selective cation channels or voltage-operated Ca2+ channels. Likewise, defining the subpopulation of β-NAD+-sensitive neurons should shed more light on the functional significance of β-NAD+ release in the brain.

In conclusion, our data provide the first evidence for the release of β-NAD+ by membrane depolarization of brain synaptosomes, for extracellular degradation and synaptosomal uptake of β-NAD+, as well as for β-NAD+ and ADPR-induced rapid activation of cortical neurons.

291

6.6 Supplementary material

Fig S6.1

Supplementary Figure S6.1. Control showing lack of effect of pressure ejection delivery method on Ca2+ dynamics in cultured neurons. (A) Position of delivery microelectrode, containing modified HEPES buffer with additional 50 mmol/L NaCl. (B) Plot showing intracellular Fura-2 ratios from four neurons selected from the field shown in (A), and lack of response to 500 ms NaCl ejection pulse (delivered at 10 min). (C) Same field, after switching delivery electrode to one containing 50 mmol/L β-NAD+ in HEPES, placed very close to the position in (A). Identical 500-ms pressure pulses generated robust Fura-2 transients in the four neurons, and this was followed by sustained Ca2+ elevations after depolarization generated by bath application of KCl (40 mmol/L).

292

Fig S6.2

Supplementary Figure S6.2. Control showing lack of effect of localized application of acidic solutions on Ca2+ dynamics in cultured neurons. The plot shows intracellular Fura-2 ratios from four neurons, with lack of response to a pair of 500-ms pH 3.0 ejection pulses. These same neurons responded robustly after switching delivery electrode to one containing 50 mmol/L β- NAD+, and this was followed by sustained Ca2+ elevations after depolarization generated by bath application of KCl (40 mmol/L).

293

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Chapter 7

Summary and Conclusions

300

The experiments described in this dissertation have investigated the role of extracellular purine nucleotides in the regulation of enteric and central nervous system functions. In particular, we have examined mechanisms of release, metabolism and action of adenosine 5’-triphosphate (ATP) and nicotinamide adenine dinucleotide (NAD+) in nerve growth factor (NGF)-differentiated rat pheochromocytoma PC12 cells, in murine and primate gastrointestinal tract and in isolated rat forebrain synaptosomes or cultured cortical neurons. We have provided evidence showing storage of NAD+ in synaptic vesicles and release via exocytosis (Yamboliev et al., 2009), we demonstrate novel neurotransmitter roles of NAD+ and/or its metabolite adenosine 5’-diphosphate ribose

(ADPR) in the enteric nervous system (Hwang et al., 2011; Durnin et al., 2012b; Durnin et al., 2013) and we identified NAD+ and/or ADPR as putative neurotransmitters in the brain (Durnin et al., 2012a). Interestingly, in the gut ATP does not appear to fulfill criteria for a motor neurotransmitter. As a result the purine hypothesis of neural regulation in the enteric and central nervous system is in need of review.

Over the last decade work in Mutafova-Yambolieva’s laboratory demonstrated using extremely sensitive high-pressure liquid chromatography (HPLC) techniques that

NAD+ is released into the extracellular compartment during neural stimulation in smooth muscles (Smyth et al., 2004; Smyth et al., 2006b; Breen et al., 2006; Smyth et al., 2009).

In blood vessels the release of NAD+ appeared to parallel that of the classical catecholamine neurotransmitter, norepinephrine (NE), as it depended on the level of nerve stimulation and was abolished by inhibitors of neural activity (Smyth et al., 2004;

Smyth et al., 2006b). These studies also demonstrated that, if released, NAD+ could elicit changes in the smooth muscle tone. For example, exogenous application of NAD+ caused 301 relaxation in blood vessels (Smyth et al., 2009) and reduced the frequency and amplitude of spontaneous contractions in the bladder (Breen et al., 2006). Furthermore, a mechanism for terminating the extracellular action of NAD+ was suggested in blood vessels by the neural localization and functional activity of the NAD+ degrading protein

CD38 (Smyth et al., 2006a). Thus NAD+ was revealed as a novel factor released during nerve stimulation in smooth muscles with a putative neurotransmitter or neuromodulator role. These were exciting findings in a time when mechanisms for regulated release of

NAD+ to the extracellular compartment were only beginning to emerge (Bruzzone et al.,

2001a). Furthermore, these findings had significant impact in the field of intracellular

NAD+ signaling in that novel intracellular storage sites of NAD+ in synaptic vesicles were revealed that had not before been identified (depicted in Billington et al., 2006).

Although these earlier studies demonstrated unequivocally that NAD+ can be released by Ca2+-dependent mechanisms during nerve stimulation, the complex organization of smooth muscle organs makes verification of vesicular NAD+ release a challenging task. Therefore in Chapter 2 we carried out experiments that aimed to verify presence of NAD+ in synaptic vesicles and release via exocytosis using a single cell model that overcomes some of the challenges associated with complex smooth muscle tissues (Yamboliev et al., 2009). Nerve growth factor (NGF)-differentiated rat pheochromocytoma PC12 cells are a single cell model that phenotypically resemble sympathetic neurons and synthesize and store catecholamine neurotransmitters (Greene

& Tischler, 1976) and ATP (Wagner, 1985). Using this model we demonstrated co- storage and release of NAD+ with catecholamines and ATP (Yamboliev et al., 2009).

These are important findings as they validate the presence of NAD+ in vesicles and 302 release via exocytotic mechanisms. This is the first study to identifiy storage of NAD+ in synaptic vesicles.

An important difference in the mechanisms releasing NAD+ and ATP was revealed in this study. Upon membrane depolarization release of ATP was not abolished after cleavage of the soluble N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE), SNAP-25, with botulinum neurotoxin A (BoNT/A) indicating that release occurred largely by non-exocytotic mechanisms (Yamboliev et al., 2009).

Previously in blood vessles ω-conotoxin GVIA-insensitive release of ATP was demonstrated (Smyth et al., 2009) suggesting neuronal N-type Ca2+ channels are not involved in ATP release in this smooth muscle. As a result these observations, in conjunction with previous uncertainties about the nature of the enteric purine transmitter

(e.g. Serio et al., 2003), led us to question the true identity of the purinergic inhibitory neurotransmitter in the gut where purine-mediated non-adrenergic non-cholinergic

(NANC) neurotransmission was originally described.

Normal gastrointestinal (GI) function is dependent on a multitude of mechanisms working in synergy and a large number of neurotransmitters and neuromodulators regulate the contractility of GI muscles. Purines are recognized as important inhibitory regulators of circular muscle contractions. In the 1970s a number of indirect observations led Geoffrey Burnstock and colleagues to assign ATP as the purinergic inhibitory motor transmitter in the guinea pig taenia coli (e.g. Burnstock et al., 1970; Burnstock & Wong,

1978). As a result, ATP is generally assumed to be the purine transmitter in smooth muscles, however presynaptic release of ATP is commonly overlooked in most studies. 303

For the first time we have revealed that upon rigorous examination of presynaptic release mechanisms and postsynaptic effects, ATP does not in fact fulfill criteria necessary to establish it as a neurotransmitter in the gut (Mutafova-Yambolieva et al.,

2007; Hwang et al., 2011; Durnin et al., 2013). In Chapter 3 (Hwang et al., 2011), we found that electrical field stimulation (EFS)-evoked release of ATP from primate colons was not released in a frequency-dependent manner and was not abolished by the neural toxins, tetrodotoxin (TTX) and ω-conotoxin GVIA, which have been demonstrated to inhibit enteric purine motor neurotransmission (Banks et al., 1979; Bridgewater et al.,

1995; Gil et al., 2010; Shuttleworth et al., 1997; Rae et al., 1998). In addition, the postjunctional membrane hyperpolarizations to exogenous ATP were not inhibited with purinergic P2Y1 receptor antagonists which have been shown to block purine-mediated inhibitory junction potentials (IJPs) in the gut (Gallego et al., 2006; Mutafova-

Yambolieva et al., 2007). On the other hand, EFS-evoked release of NAD+ was both frequency-dependent and neurotoxin-sensitive and postsynaptic hyperpolarizations to exogenous NAD+ were abolished by inhibition of P2Y1 receptors and small conductance

Ca2+-activated SK channels (Hwang et al., 2011). Thus the pharmacology of NAD+ better mimicked the endogenous purinergic motor transmitter than ATP in primate colons

(recently reviewed in Rodriguez-Tapia & Galligan, 2011). In fact NAD+ may represent an important target for treating motility disorders of the human large intestine as inhibitory regulation in human colonic muscles is mediated predominantly by purines

(Hwang et al., 2011).

In monkey colon close examination of electrical responses to NAD+ revealed transient depolarizations after blocking P2Y1 receptors and in human colon transient 304 contractions to NAD+ sometimes occurred before relaxation (Hwang et al., 2011). These findings suggested that NAD+ might activate opposing conductances in different cells. In isolated smooth muscle cells we demonstrated that exogenous NAD+ activated inward currents via non-selective cation channels that would lead to depolarization in the cell rather than the expected hyperpolarization. Therefore smooth muscle cells are unlikely to mediate purinergic IJPs in the gut. However Kurahashi et al., (2011) demonstrated that platelet-derived growth factor receptor α-positive (PDGFRα+) cells, which are in close proximity to nerve terminals and couple to smooth muscle via gap junctions, may be the cell type mediating purinergic IJPs as exogenous NAD+ activates apamin-sensitive large outward currents in these cells. These cells also abundantly express purinergic P2Y1 receptors (Kurahashi et al., 2011; Peri et al., 2013). Thus release of NAD+ from nerve terminals likely activates P2Y1 receptors on PDGFRα+ cells to cause hyperpolarization and relaxation in the smooth muscle.

Once released to the extracellular compartment, purines are quickly degraded by extracellular nucleotidases to terminate their activity. In primate colons we found that during nerve stimulation a substantial amount of ADPR was present in the extracellular compartment (Hwang et al., 2011). Previous studies have shown that ADPR can activate

P2Y1 receptors in INS-1E cells and pancreatic β-cells from rat and human (Gustafsson et al., 2011). Therefore we were interested in examining the potential contribution of ADPR to purinergic relaxation responses in the gut. In Chapter 4 (Durnin et al., 2012b) we found that ADPR could indeed be produced very rapidly from NAD+ in the colon.

Moreover, exogenous ADPR also produced membrane hyperpolarizations, the pharmacology of which matched the endogenous purine neurotransmitter. On the other 305 hand the ATP metabolite, ADP, failed to mimic the pharmacology of the endogenous neurotransmitter. Finding that ADPR is bioactive in GI muscle suggests that multiple purines could contribute to postjunctional inhibitory responses in the gut and purinergic regulation of smooth muscle contractility is likely to be much more complex than originally believed. In addition, our study reveals the inadequacy of many studies that utilize stable purine analogs to examine responses to purines (e.g. Gallego et al., 2006;

Gallego et al., 2008b); with the rapid degradation that inevitably occurs once purines are released into the extracellular compartment these studies overlook the contribution of metabolites to overall responses produced during nerve stimulation.

This study also revealed important differences in the preferential sites of degradation of extracellular NAD+ and ATP in colon. In monkey circular muscle preparations that have the ganglia removed there was more pronounced degradation of exogenous NAD+ (and ADPR) per unit mass compared to whole muscle preparations.

Thus degradation of NAD+ and ADPR appears to occur close to the nerve terminals (i.e. at the site of release of NAD+/ADPR) (Durnin et al., 2012b). In agreement with these observations, a recent study demonstrated high expression of the gene encoding CD38 in

PDGFRα+ cells (Peri et al., 2013); thus enzymes localized on PDGFRα+ cells would provide an effective mechanism for terminating responses elicited by NAD+ released from enteric nerve terminals. On the other hand, ATP was not degraded exclusively at nerve terminals as suggested by similar degradation in colonic preparations with and without myenteric ganglia (Durnin et al., 2012b); ATP may therefore be released as a paracrine mediator. Overall this study provides further evidence that NAD+/ADPR are better candidates than ATP (and ADP) as inhibitory enteric neurotransmitters. 306

Our studies have evoked controversy in the field of enteric purine neurotransmission and some investigators remain skeptical that NAD+/ADPR are involved in enteric inhibitory responses (Goyal, 2011). Therefore it was important for us to approach this issue using a methodology alternative to electrical field stimulation to measure presynaptic purine release. In Chapter 5 (Durnin et al., 2013), we examined release of purines upon activation of ligand-gated ion channel receptors localized on myenteric nerve cell bodies and dendrites in murine and primate colonic muscles.

Stimulation of ganglionic nicotinic acetylcholine (nACh) or serotonin 5-HT3 receptors elicits motor responses in GI muscles by mediating fast excitatory postsynaptic potentials

(fEPSPs) and activating action potentials which propagate down axons, depolarize nerve terminals and activate N-type voltage-dependent Ca2+ channels. Ca2+ entry facilitates fusion of vesicles and transmitter release.

In these studies we found that activation of either nACh or 5-HT3 receptors stimulated release of both ATP and NAD+ from colonic muscles, however release was differentially mediated and thus likely occurred from different sites within the myenteric plexus. Release of NAD+ was consistent with properties of motor neurotransmission and required action potential propagation and Ca2+ influx to nerve terminals via neural N-type voltage-dependent Ca2+ channels; thus NAD+ likely originated from nerve terminals as a neurotransmitter. On the other hand, the release of ATP might have originated from the nerve cell body due to release via TTX- and ω-conotoxin GVIA-insensitive mechanisms

(Durnin et al., 2013).

In addition to their neuronal localization, some nAChRs are expressed by non- neuronal cells in the gut (Wessler & Kirkpatrick, 2008). It is conceivable that some 307 release of ATP during activation of nAChRs occurs from non-neuronal sources due to the

TTX-insensitivity of release. Determining the mechanism of ATP release was outside the scope of these studies however we did reveal that connexin hemichannels are unlikely to be involved (Durnin et al., 2013). Regardless of the mechanism regulating release, our studies suggest that ATP might be released as a paracrine substance in ganglia or other non-neuronal source but does not function as a motor neurotransmitter in colon. These findings have been highlighted in a recent article in Nature Reviews Gastroenterology and Hepatology (Smith, 2013).

In summary, NAD+ and/or ADPR appear to be the primary neurotransmitters involved in purinergic inhibitory regulation of colonic motility; targeting NAD+/ADPR synthesis, release or metabolism may provide unique opportunities for therapeutically treating problems of altered colonic transit.

There is surprisingly little known about purinergic signaling in the central nervous system (CNS). Many investigators assume ATP to be the purine neurotransmitter involved in neuronal-neuronal signaling however accumulating evidence also demonstrates ATP release from non-neuronal cells, such as glial cells, in the CNS by mechanisms including release through connexin hemichannels (Stout et al., 2002), P2X7 receptor pores (Duan & Neary, 2006), volume regulated anion channels (Darby et al.,

2003) and via vesicle exocytosis (Coco et al., 2003; Montana et al., 2006). Thus in the

CNS ATP could be released from both neuronal and non-neuronal sources by both Ca2+- dependent and Ca2+-independent mechanisms. The question of whether ATP in fact functions as a neurotransmitter in the brain remains unanswered. These observations as 308 well as the strong evidence accumulated for possible neurotransmitter role of NAD+ in the peripheral nervous system led us to examine the potential role of NAD+ in the CNS.

Our studies demonstrate for the first time that NAD+ meets criteria to establish it as a potential neurotransmitter in the brain. In isolated rat forebrain synaptosomes release of NAD+ upon membrane depolarization was dependent on intact exocytosis machinery and was significantly reduced by neural toxins BoNT/A and ω-conotoxin GVIA and by inhibition of the proton gradient of synaptic vesicles with bafilomycin A1 (Durnin et al.,

2012a). In addition we demonstrated that localized application of exogenous NAD+ in cortical neurons caused rapid Ca2+ transients suggesting that endogenously released

NAD+ could participate in neuron-to-neuron communication in the brain. Exogenous

ADPR similarly produced Ca2+ transients in cortical neurons. Interestingly, we found that subpopulations of cortical neurons appeared to respond preferentially to either ATP or to

NAD+/ADPR which might suggest different mechanisms of action of these purines in the brain. Once again our studies highlight that neuronal excitability in the brain may also be regulated by multiple purine substances. While detailed characterization of the receptors involved in mediating responses to NAD+/ADPR was not the aim of this study, it appears that P2X1 and P2X3 receptors are only partially responsible for the Ca2+ transients elicited by NAD+ or ADPR (Durnin et al., 2012a).

Finally for neurotransmission to be effective a mechanism for terminating the extracellular action of NAD+ should exist in brain synaptosomes. We demonstrated that

CD38 was expressed by brain synaptosomes and exogenous NAD+ was degraded once in contact with synaptosomes. In addition we revealed a reuptake mechanism in brain synaptosomes probably as an alternative mechanism for terminating and recycling 309 released NAD+ (Durnin et al., 2012a). Previous studies have suggested NAD+ uptake through connexin hemichannels however our studies rule against the involvement of these channels in NAD+ transport in brain synaptosomes. Future studies may elucidate the specific mechanism involved in NAD+ uptake.

These experiments provide the first evidence for a putative neurotransmitter role of NAD+/ADPR in the CNS. While we do not exclude the possibility that ATP may function as a neurotransmitter in some CNS neurons, we emphasize the importance in considering purines other than ATP that could also participate in regulating neuronal excitability in the brain.

Overall the work described in this dissertation provides novel information on the storage, release, metabolism and extracellular action of NAD+/ADPR in the enteric and central nervous systems. Knowing the identity of the purinergic neurotransmitter is important for understanding critical mechanisms of cell-to-cell communication and may also have important translational implications. The NAD+/ADPR system may represent a novel therapeutic target for disorders associated with disrupted purinergic mechanisms in the central and peripheral nervous systems.

310

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