DISRUPTIONS IN NEUROMUSCULAR TRANSMISSION AND COLONIC MOTILITY IN DIET-INDUCED OBESITY By Yogesh Bhattarai A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Neuroscience – Doctor of Philosophy 2015   ABSTRACT DISRUPTIONS IN NEUROMUSCULAR TRANSMISSION AND COLONIC MOTILITY IN DIET-INDUCED OBESITY By Yogesh Bhattarai Diet-induced obesity (DIO) is primarily driven by the consumption of a high fat (HF) diet and is recognized as a major risk factor for the development of gastrointestinal (GI) motility complications, especially in the colon. Colonic motility is regulated by the division of the autonomic nervous system called the enteric nervous system (ENS). Despite high incidences of GI motility disorders in HF DIO, the role of enteric nervous system (ENS) in the development of these complications is unclear. The ultimate goal of this translational project was to understand how the consumption of a HF diet leads to impaired colon motility through effects on the cells controlling neuromuscular transmission. The etiology of obesity–induced colon motility disorders is multi-factorial and includes alterations to multiple cell types including enteric nerves and the GI smooth muscles. These cells communicate to control contraction and relaxation of smooth muscle via series of coordinated inhibitory and excitatory signaling mechanisms. Disturbances in these signaling mechanisms contribute to colon motility disorders. Therefore, it is crucial to identify in detail how neuromuscular transmission and smooth muscle excitability are regulated before the pathogenesis of HF diet associated colonic dysmotility can be studied. In the first part of this dissertation I studied the etiology of inhibitory neuromuscular transmission and smooth muscle excitability in the mouse distal colon. I found that inhibitory neuromuscular transmission is primarily mediated by ATP acting on P2Y1 receptors and nitric oxide (NO) diffusion on to the GI smooth muscle. These neurotransmitters act on the small   conductance Ca2+-activated K+ (SK) channels and the Ca2+-activated Cl- (CaCCs) channels to produce IJPs in the mouse distal colon. In addition, I discovered that the large conductance Ca2+-activated K+ channels (β1BK channel) play a crucial role in regulation smooth muscle excitability by controlling smooth muscle action potential firing activity in the mouse distal colon. Next, I sought to determine how HF diet alters neuromuscular signaling and smooth muscle cell physiology. My results show that HF diet specifically disrupts NO mediated inhibitory neuromuscular transmission, SM relaxation and excitability leading to impaired colonic propulsive motility. Genetic KO of β1BK channel closely recapitulates HF diet induced impairments in the mouse distal colon. This suggests that compromised BK channel β1-subunit function/expression might play a role in mediating colonic dysmotility in obesity.   ACKNOWLEDGMENTS I would like to express my sincere appreciation and gratitude to my advisor Dr. James Galligan for all the support and encouragement that he has provided over the years. He has been a phenomenal mentor who has stimulated with in me a sense of excitement for research, and provided critical insights and feedback when necessary. I would also to thank Dr. Brian Gulbransen and Dr. Mark Kadrofske for allowing me complete a significant portion of my dissertation work in their labs. I am sincerely grateful to Dr. Greg Swain for providing his guidance and support through out my PhD training. This dissertation would not have been possible without the constant supervision, guidance and constructive criticism of my committee members. It has been a real honor to work with all of them. I owe my sincere gratitude to Dr. Hui Xu, Roxanne Fernandes, and Beth Lockwood for teaching me new techniques and helping me with non-technical aspects of the research. I would also like to thank my lab members and friends Marion France, Nadine El-Ayache, Eileen Rodriguez-Tapia, Ryan Mui, David Fried, Prafulla Regmi and Ujjwal karki who made my stay in MSU all the more fun and an enjoyable experience. I am indebted to the Neuroscience Program at MSU for providing me with funding and all other logistics to help me complete my graduate education. I cannot thank my family enough for all their scarifies. They are inseparable part of my life and the greatest source of my inspiration. I would therefore like to dedicate this dissertation to my family members Manish Bhattarai (brother), Prerana Bastola (sister-in-law); Madhav Bhattarai (dad), and Pushpa Bhattarai (mom) for standing besides me and dealing with my absence for all these years.   iv     TABLE OF CONTENTS LIST OF TABLES……………………….……………………………………………………….ix LIST OF FIGURES.………………………………………….…………………………………...x KEY TO ABBREVIATIONS…………………………….…………………………………….xiii CHAPTER 1: GENERAL INTRODUCTION…….…………………………….………………..1 1.1 Overview of the nervous system………………………………...…………………………….2 1.2 Overview of the gastrointestinal (GI) tract……………………………...…………….4 1.2.1 GI histology……………………………...…………….……………………4 1.2.1.1 Mucosa…………………………….………………………………4 1.2.1.2 Submucosa…………………………….…………………………..6 1.2.1.3 Muscularis…………………………….…………………………...6 1.2.1.4 Serosa…………………………….………………………………..6 1.3 Intrinsic and Extrinsic innervations to the GI tract……………………………………8 1.3.1 Extrinsic innervations…………………………….…………………………8 1.3.2 Intrinsic innervations…………………………….………………………….9 1.4 Morphological classification of enteric neurons……………………………………..11 1.4.1 Dogiel type I…………………………….…………………………………11 1.4.2 Dogiel type II…………………………….………………………………...12 1.4.3 Dogiel type III…………………………….………………………………..13 1.5 Structural and physiological characteristics of the GI Smooth muscle cells………...14 1.5.1 Structure of the GI smooth muscle…………………………….…………..14 1.5.2 Smooth muscle contraction…………………………….…………………..14 1.5.3 Smooth muscle relaxation…………………………….……………………16 1.6 Interstitial cells of Cajal (ICCs)…………………..…………………..……………...18 1.6.1 Function of ICCs…………………..……………………………………….18 1.6.2 Generation of pacemaker current and slow wave production……………...19 1.7 Neuromuscular transmission in the GI tract…………………………………………20 1.7.1 Inhibitory neuromuscular transmission in the GI tract…………………….20 1.7.2 Excitatory neuromuscular transmission in the GI tract…………………….21 1.8 GI motility…………………………….……………………………………………...23 1.8.1 Rhythmic phasic contractions (RPCs)……………………………………..24 1.8.2 Giant migrating contractions (GMCs)……………………………………..24 1.8.3 Tonic contractions…………………………….……………………………25 1.9 Peristalsis…………………………….………………………………………………26 1.9.1 Peristaltic circuit …………………………….…………………………….26 1.10 Ca2+-activated K+ channels …………………………….…………………………..28 1.10.1 Large conductance Ca2+-activated K+ channels (BK channel)…………...28 1.10.1.1 β1 BK channels……………………….………………………...30 1.10.2 Small conductance Ca2+-activated K+ channels………………………......31 1.10.3 Intermediate conductance Ca2+-activated K+ channels…………………...32 1.11 Obesity and its associated disorders………………………….…………………….34   v     1.11.1 High fat (HF) diet-induced obesity (DIO) and disruption in colonic motility…………………………………………………………………...34 1.12 Oxidative stress…………………………….……………………………………….36 1.12.1 Effects of oxidative stress on biomolecules………………………………37 1.12.1.1 Nucleic acid…………………………….………………………37 1.12.1.2 Lipids…………………………….……………………………..37 1.12.1.3 Proteins…………………………….…………………………...37 1.12.2 Obesity and oxidative stress…………………………….………………..38 1.12.3 Impact of oxidative stress on inhibitory nitrergic signaling.……………..39 1.12.4 Impact of oxidative stress on β1BK channel function.…………………...41 1.13 BK channel and inhibitory nitric oxide signaling………………….……………….43 1.14 Hypothesis and specific aims…………………………….…………………………45 BIBLIOGRAPHY………………………………………………………………………………..46 CHAPTER 2 MECHANISM OF INHIBITORY NEUROMUSCULAR TRANSMISSION IN THE MOUSE DISTAL COLON………………………………………………………………...58 2.1 Abstract………………………………………………………………………………59 2.2 Introduction..…………………………….…………………………………………...60 2.3 Materials and methods…………………………….…………………………………62 2.3.1 Ethics statement…………………………….……………………………...62 2.3.2 Animals…………………………….………………………………………62 2.3.3 Electrophysiological recordings……………………………………………62 2.3.4 Local drug applications…………………………………………………….62 2.3.5 Solution and Drugs………………………………………………………...63 2.3.6 Statistics……………………………………………………………………64 2.4 Results……………………….……………………………………………………….65 2.4.1 Nifedipine application does not affect Resting membrane potential (RMP) in the mouse colon…………………………………………………….....................65 2.4.2 IJPs display both uniphasic and biphasic patterns…………………………65 2.4.3 IJPs are neurogenic……………………………………………...................67 2.4.4 Identification of receptors and neurotransmitters that mediate IJPs in the mouse colon……………………………………………………...............68 2.4.5 Identification of smooth muscle ion channels involved in mediating IJPs……………………………………………………………………….70 2.4.6 Identification of specific purinergic neurotransmitter involved in mediating the IJP…………………………………………………………………….72 2.5 Discussion…………………………….……………………………………………...76 BIBLIOGRAPHY…………………………….…………………………………………….........80 CHAPTER 3 β1BK CHANNEL REGULATES SMOOTH MUSCLE EXCITABILITY IN THE MOUSE DISTAL COLON…………………………………….………………………………...84 3.1 Abstract…………………………………………………………………………........85 3.2 Introduction..…………………………….…………………………………………...86 3.3 Materials and methods…………………………….…………………………………88 3.3.1 Ethics statement…………………………….……………………………...88 3.3.2 Animals…………………………….………………………………………88   vi     3.3.3 Electrophysiological recordings……………………………………………88 3.3.4 Drugs…………………………….…………………………………………89 3.3.5 Statistics……………………………………………………………………90 3.4 Results………………………….…………………………………………………….90 3.4.1 β1Bk channel does not mediate inhibitory neuromuscular transmission in the mouse distal colon……………………………………………………90 3.4.2 Tetrodotoxin (TTX) application does not block smooth muscle action potential in colonic smooth muscles……………………………………..92 3.4.3 Increased action potential activity in distal colonic smooth muscle from β1subunit KO mice…………………………………………………………93 3.4.4 Effects of nifedipine application in action potential generation and smooth muscle membrane potential in WT and β1-subunit KO……………...….95 3.4.5 Effects of Paxilline on membrane potential and action potential firing in distal colonic smooth muscle of WT mice………...…..............................97 3.5 Discussion……………………………………………………………………………99 BIBLIOGRAPHY……………………………………………………………………………....102 CHAPTER 4 HIGH FAT DIET DIRUPTS INHIBITORY NEUROMUSCULAR TRANSMISSION AND SMOOTH MUSCLE EXCITABILITY IN THE MOUSE DISTAL COLON………………………………….……………….……………………………………..106 4.1 Abstract……………………………………………………………………………..107 4.2 Introduction..…………………………….………………………………………….108 4.3 Materials and methods…………………………….………………………………..110 4.3.1 Ethics statement…………………………….…………………………….110 4.3.2 Animals…………………………….……………………………………..110 4.3.3 Oxidative stress measurements…………………………………………...110 4.3.4 Immunohistochemistry for nitric oxide synthase (NOS) containing neurons………………………………………………………………….111 4.3.5 Nitric oxide (NO) measurements ……………………………...................112 4.3.6 Electrophysiological recordings from colonic circular muscle SMCs.......................................................................................................112 4.3.7 Isometric tension measurement of muscle relaxation in-vitro……............113 4.3.8 In-vitro assessment of colonic motility…………………….……………..114 4.3.9 In-vivo assessment of colonic motility…………………………………...114 4.3.10 Western blot for α subunit protein expression………………..................115 4.3.11 Real time rt-PCR for α and β1 subunits mRNA expression…………….116 4.3.12 Statistics…………………………….………………………...................116 4.4 Results…………………………….………………………………………………...117 4.4.1 HF diet increases body weight and abdominal fat pad weight…………...117 4.4.2 WT HF mice exhibit increased neuronal oxidative stress, decreased NOS neuron density and lower NO availability……………………..……….117 4.4.3 HF diet impairs inhibitory nitrergic neuromuscular transmission………..119 4.4.4 HF diet impairs neurogenic smooth muscle relaxation……………….......120 4.4.5 HF diet mimics the effect of β1BK KO on SM electrical activity……….122 4.4.6 NO availability affects SM excitability in WT and β1BK KO mice……..122   vii     4.4.7 WT HF mice exhibit disruptions in colonic migrating motor complex (CMMC) activity and CMMCs are disrupted in β1BK KO mice regardless of diet…………………………………………………………………...124 4.4.8 Consumption of a HF diet increases fecal pellet output in WT mice…….126 4.4.9 α-subunit protein expression of a β1BK channel is not altered in HF mice……..................................................................................................127 4.4.10 β1-subunit mRNA expression is not changed in HF mice……………....128 4.5 Discussion…………………………….…………………………………………….129 BIBLIOGRAPHY…………………………….………………………………………………...134 CHAPTER 5 WESTERN BLOT ANALYSIS OF BK CHANNEL β1- SUBUNIT EXPRESSION SHOULD BE INTERPRETED CAUTIOUSLY WHEN USING COMMERCIALLY AVAILABLE ANTIBODIES……………….………………...…………140 5.1 Abstract……………………………………………………………………………..141 5.2 Introduction..…………………………….………………………………………….142 5.3 Materials and methods…………………………….………………………………..144 5.3.1 Ethics statement…………………………………………………………..144 5.3.2 Animals…………………….……………………………………………..144 5.3.3 Western blot…………………………….………………….......................144 5.3.4 Real time RT-PCR………………………………………………………..147 5.4 Results…………………………….………………………………………………...149 5.4.1 Western blots in MA and colons from WT and BK β1-KO mice………..149 5.4.2 BK β1-subunit mRNA expression in tissues from BK β1-KO mice …….150 5.4.3 Expression of BK α-subunit in the colons…………………..……………151 5.5 Discussion…………………………….…………………………………………….153 BIBLIOGRAPHY…….…………………………….…………………………………………..157 CHAPTER 6 KEY FINDINGS, SIGNIFICANCE AND FUTURE DIRECTIONS…………...162 6.1 Key findings…..…………………………….………………………….…………...163 6.2 Significance…………………………….…………………………….……………..166 6.3 Future directions……………………………………………………………………167 BIBLIOGRAPHY………………………………………………………………………………169   viii     LIST OF TABLES Table 1.1 Electrophysiological and pharmacological characteristics of Ca2+-activated K+ channels…………………………………………………………………………………………..33 Table 5.1 List of commercially available anti-BK β1-subunit antibodies……………………..146 Table 5.2 Publications that performed BK β1-subunit western blot analysis using commercially available antibodies……………………………………………………………………………..154   ix     LIST OF FIGURES Figure 1.1 Organization of the nervous system…….…………………………………………….3 Figure 1.2 Histological layers of the gastrointestinal (GI) tract………………………………….7 Figure 1.3 Organization of the enteric nervous system (ENS)…..……………………………...10 Figure 1.4 Schematic diagram of Dogiel type I neuron as classified by Dogiel………………...11 Figure 1.5 Schematic diagram of Dogiel type II neuron as classified by Dogiel……………….12 Figure 1.6 Schematic diagram of Dogiel type III neuron as classified by Dogiel………………13 Figure 1.7 Schematic diagram illustrating smooth muscle contraction…………………………16 Figure 1.8 Schematic diagram illustrating smooth muscle relaxation…………….…………….17 Figure 1.9 Excitatory and inhibitory neuromuscular transmission in the GI tract. ……………..22 Figure 1.10 Large conductance Ca2+-activated K+ (β1BK) channel structure…………………..30 Figure 1.11 Small conductance Ca2+-activated K+ (SK) channel structure……………………..31 Figure 1.12 Model describing mechanisms of oxidation of biomolecules.……………………..38 Figure 1.13 Model indicating how accumulated fat in obesity leads to obesity associated disorders………………………………………………………………………………………….39 Figure 1.14 Mechanism of peroxynitrite (ONOO-) production in the inhibitory nitric oxide (NO) neurons …………………………………………………………………………………………..41 Figure 1.15 Mechanism of disruption of β1BK channel function via oxidative stress………….42 Figure 1.16 Mechanism of NO production and β1BK channel activation in the neuromuscular junction.………………………………………………………………………………………….44 Figure 2.1 Effect of nifedipine on colonic smooth muscle resting membrane potential..............65 Figure 2.2 IJPs have both a uniphasic and biphasic patterns……………………………………66 Figure 2.3 Schematic diagram illustrating neuromuscular transmission in the mouse colon…...67 Figure 2.4 The effect of P2Y1 receptor blocker on evoked IJPs………………………………..68   x     Figure 2.5 The effect of NOS inhibitor on evoked IJPs……………………………………..…..69 Figure 2.6 Effect of MRS 2179 and L-NNA co-application…………...…….……………….....69 Figure 2.7 The Effects of SK channel blocker on evoked IJP……………...…………………...70 Figure 2.8 Effects of CaCC blocker on evoked IJP……………………………………………..71 Figure 2.9 Effects of apamin and niflumic acid (NFA) co-application…………………………71 Figure 2.10 ATP and β-NAD induced hyperpolarizing response in the mouse distal colon.…...73 Figure 2.11 Effects of POM-1 on the amplitude and area of IJP in the mouse distal colon…….74 Figure 2.12 Effect of POM-1 on ATP response..……………………………………...………...75 Figure 2.13 Effect of POM-1 on exogenous Adenosine 5′-[β-thio]diphosphate (ADP-β-S)…...75 Figure 2.14 Effect of MRS 2179 on exogenous ADP-β-S……………………………………....75 Figure 3.1 IJP measurements from WT and β1-subunit KO.…………………………………...90 Figure 3.2 Effects of paxilline on IJPs…………………………………………………………..91 Figure 3.3 Effect of co-application of apamin and paxilline on IJPs……………………………91 Figure 3.4 Effect of tetrodotoxin (TTX) on smooth muscle action potential.…………………..92 Figure 3.5 Action potential firing in WT and β1-subunit KO…………………………………..94 Figure 3.6 Nifedipine blocks action potential firing in WT and BK β1-subunit KOs…………..96 Figure 3.7 Effect of paxilline on smooth muscle action potential firing ……………………….97 Figure 3.8 Effect of paxilline on the smooth muscle membrane potential……………………...98 Figure 4.1 Glutathione fluorescence in the myenteric plexus of the mouse distal colon……...118 Figure 4.2 Immunohistochemical labeling for neuronal nitric oxide synthase (nNOS) containing neurons from control and high fat (HF) mice…………………………………………………..119 Figure 4.3 IJP recordings from control and HF mice.…………………………………………120 Figure 4.4 Measurement of neurogenic smooth muscle relaxation in the mouse distal colon...121 Figure 4.5 NO modulates smooth muscle excitability…………………………………………123   xi     Figure 4.6 In-vitro colonic migrating motor complex (CMMC) recordings from WT and β1subunit KO mice placed on control and HF diet.……………………………………………….125 Figure 4.7 In-vivo colonic motility measurements from WT and β1-subunit KO mice……….126 Figure 4.8 BK channel α-subunit western blot ………………………………………………..127 Figure 4.9 rt-PCR measurements for the α and the β1subunit mRNA expression…………….128 Figure 5.1 Representative western blot obtained using commercially available anti-BK β1 subunit antibodies..……………………………………………………………………………..150 Figure 5.2 Real-time RT-PCR analysis of BK β1-subunit and GAPDH in MA and colons from WT and BK β1-KO mice……………………………………………………………………….151 Figure 5.3 BK α‐subunit western blot in colonic tissues from WT and BK β1‐KO mice……......................................................................................................................................152 Figure 6.1 Mechanism of IJP production in the mouse distal colon.…………………………..163 Figure 6.2 Regulation of colonic smooth muscle excitability via β1BK channel function……164 Figure 6.3 Mechanisms of colonic dysmotility due to HF diet consumption………………….165   xii     KEY TO ABBREVIATIONS ACh Acetylcholine ADP Adenosine diphsphate ADP-β-S Adenosine 5′-[β-thio]diphosphate trilithium salt AMP Adenosine monophosphate Ano1 Anoctamin-1 ANS Autonomic nervous system ATP Adenosine triphosphate β1BK Large conductance Ca2+-activated K+ channels CaCC Ca2+-activated Cl- channels CaCM Ca2+-calmodulin complex CMMC Colonic migrating motor complex CNS Central nervous system DAF-FM-DA Diaminofluorophore 4-amino-5-methylamino-2′-7′difluorofluorescein diacetate DIO Diet-induced obesity DMSO Dimethyl sulfoxide EFS Electrical field stimulation ENS Enteric nervous system fIJP Fast inhibitory junction potential GI Gastrointestinal GSH Reduced glutathione GSSG Glutathione disulphide   xiii     5-HT 5-Hydroxytryptamine HF High fat H2 O2 Hydrogen peroxide IJP Inhibitory junction potential ICC Interstitial cells of Cajal IK Intermediate conductance Ca2+-activated K+ channels IPANS Intrinsic primary afferent neurons KO Knock out L-NNA Nitro-L-arginine LMMP Longitudinal muscle myenteric plexus MLCK Myosin light chain kinase MRS2179 2′-deoxy-N6-methyladenosine 3′, 5′-bisphosphate tetrasodium salt NANC Non-adrenergic, non-cholinergic NEM N-ethylmaleamide; NO Nitric oxide nNOS Neuronal nitric oxide synthase NFA Niflumic acid NTPDase Ecto-nucleotidases β-NAD β–nicotinamide adenine dinucleotide NMT Neuromuscular transmission O2 - Superoxide anion ONOO- Peroxynitrite PA Palmitic acid   xiv     PFA Paraformaldehyde PKG Protein kinase g PNS Peripheral nervous system POM-1 Sodium polyoxotungstate 1 PSNS Parasympathetic nervous system P2Y1 Purinergic receptor RMP Resting Membrane potential ROS Reactive oxygen species SERCA Smooth endoplasmic reticulum Ca2+ ATPase sIJP Slow inhibitory junction potential SK Small conductance Ca2+-activated K+ channels SMC Smooth muscle cell SNP Sodium nitroprusside SNS Sympathetic nervous system SOD Superoxide dismutase TTX Tetrodotoxin TEA Tetra ethyl-ammonia T2D Type 2 diabetes VIP Vasoactive intestinal peptide WB Western blot WT Wild type   xv     CHAPTER 1 GENERAL INTRODUCTION   1     1.1 Overview of the nervous system The nervous system is a complex network of nerve cells and fibers that communicate electrochemical information back and forth between various regions of the body. The nervous system is divided into the central nervous system (CNS) and the peripheral nervous system (PNS). The CNS consists of the brain and the spinal cord, while the PNS consist of nerves and ganglia that lie outside the CNS. The PNS is further subdivided into the somatic and autonomic nervous systems. The somatic nervous system carries out various voluntary body functions like the movement of limbs and processing of sensory stimuli including hear, touch and sight, while the autonomic nervous system (ANS) maintains involuntary functions like controlling heart rate and digestion. The ANS comprises the enteric nervous system (ENS), the sympathetic nervous system (SNS) and the parasympathetic nervous system (PSNS) (Kandel et al., 2000) The ENS is an autonomous entity, which regulates the gastrointestinal (GI) function (Furness, 2006). The SNS and the PSNS on the other hand have a modulatory role and are distributed mostly to internal organs (heart, lungs, and intestine), secretory glands (lachrymal, salivary gland sweat gland) and blood vessels. The SNS arises from the thoraco-lumbar region of the spinal cord and prepares the body for the “fight and flight” response by increasing blood supply to the skeletal muscles, increasing the heart rate, inhibiting GI functions and dilating the pupils. The PSNS arises from the cranio-sacral region of the spinal cord and prepares the body for “rest and digest” response by slowing heart rate, constricting pupils and promoting digestion (Kandel et al., 2000).   2     Figure 1.1 Organization of the nervous system.   3     1.2 Overview of the gastrointestinal (GI) tract The GI tract is an organ system that extends from the mouth to the anus. The primary function of the GI tract is to release digestive enzymes, absorb nutrients and excrete undigested waste. The GI tract is mesodermal in origin and is developmentally divided into the foregut, midgut and hindgut. The foregut consists of the esophagus, stomach and duodenum. Ligament of Treitz, a duodenojejunal flexure demarcates the boundary between the foregut and the midgut. The midgut consists of lower duodenum, jejunum, ileum, caecum, ascending colon and first twothird of the transverse colon. The hindgut consists of the remaining one third of the transverse colon, descending colon, rectum and the anal canal. Although different regions of the GI tract have structural and functional differences, the basic histology of the GI tract remains the same. 1.2.1 GI histology Histologically, the GI tract is divided into four major layers namely: the mucosa, submucosa, muscularis and serosa. The architecture and the function of these layers are described below. 1.2.1.1 Mucosa The surface architecture of the mucosa varies depending on the location and the function of the mucosa within the GI tract. The mucosa in the stomach and the small intestine increases the surface area by forming “rugae” (gastric folds) and “plicae circulares” (intestinal folds) respectively to help facilitate release of digestive enzymes and absorption of nutrients. Unlike the stomach and the small intestine, the mucosa in the large intestine are not involved in nutrient absorption and therefore does not contain any folds comparable to rugae or plicae (Sarna, 2010). The mucosa consists of an epithelium and underlying lamina propia with the muscularis mucosa providing the demarcation between mucosa and submucosa. The epithelium in the   4     stomach consists of columnar epithelial cells including the glandular and the endocrine cells, which are responsible for secretion of alkaline mucous, gastric acids and enzymes. In the small intestine, the intestinal mucosa protrudes from the epithelial lining to form finger like projections called the “intestinal villi.” The villus contains endocrine cells that release gastrin, somatostatin, cholecystokinin and secretin into the lumen of the small intestine. The major function of the villi is to increase the surface area of the intestine and serve as a site of absorption for fluids and nutrients. The mucosa of the large intestine on the other hand contains thin brush border layer of columnar epithelium with numerous ‘mucin’ producing goblet cells to help lubricate the stool (Sarna, 2010). The middle layer of lamina propria contains loose connective tissues, nerves, blood vessels, fibroblasts and immune cells. In the small intestine, however, lymphocytes often invade the epithelium and form lymphoid nodules called the “peyer's patch.” Peyer’s patch in the lamina propria facilitates mucosal immune response. Finally, a thin muscle layer of muscularis mucosa is found in the innermost portion of mucosa. Muscularis mucosa is more prominent in the large intestine compared to the small intestine where it influences absorptive and secretory functions of the epithelium via smooth muscle contractions and relaxations (Uchida & Kamikawa, 2007).   5     1.2.1.2 Submucosa The submucosa of the large and the small intestine contains blood vessels, lymph vessels, connective tissues and the submucosa plexus. Besides this, the duodenal submucosa also contains ‘Brunner's glands,’ which release alkaline mucous secretion to protect small intestinal mucosa against the acidic environment. The submucosa plexus, also known as Meisssner’s plexus, is located above the muscularis layer and contains nerves that control gastric and intestinal secretions. 1.2.1.3 Muscularis The muscularis is a muscle layer that is located between the serosa and the submucosa. In the large and the small intestine, the muscularis contains the circular and the longitudinal muscle layer. In the stomach, however, an additional inner oblique muscle layer is also found. Between the circular and the longitudinal muscle layer, a layer longitudinal muscle myenteric plexus (LMMP) also known as the “Aurebach’s plexus” is found. LMMP contains intrinsic primary afferent neurons (IPANS), interneurons and the excitatory and the inhibitory motor neurons which form local circuits to mediate GI motility (Furness, 2007c). 1.2.1.4 Serosa The serosa consists of secretory epithelial cells, blood vessels and adipose tissues. The epithelial cells form a thin membranous sheath and encapsulate the GI tract. These epithelial cells secrete a serous fluid, which provides lubrication to reduce friction generated by the muscularis layer.   6     Figure 1.2 Histological layers of the gastrointestinal (GI) tract.   7     1.3 Intrinsic and Extrinsic innervations to the GI tract 1.3.1 Extrinsic innervations The extrinsic innervations to the GI tract are provided via the SNS and the PSNS. In the SNS, the pre-ganglionic nerve fibers arise from the thoraco-lumbar region (T5-L2) of the spinal chord. The pre-ganglionic fibers from the thoracic region of spinal chord make a synaptic connection with the post-ganglionic nerve fibers in the celiac and the superior mesenteric ganglion to innervate the proximal colon, while the pre-ganglionic fibers from lumbar region extend to the inferior mesenteric ganglion and innervate the distal colon. The sympathetic preganglionic nerve fibers releases acetylcholine (ACh), while the post-ganglionic nerve fibers release norepinephrine (NE) as the neurotransmitter. The sympathetic innervations through these pre-vertebral ganglia serves as an integration center between CNS and the ENS to decreases peristaltic speed (Gribovskaja-Rupp et al., 2014), blood flow, and secretion during a “fight or flight” response (Anuras, 1992). The PSNS on the other hand consists of the vagus and the pelvic nerves. These nerves arise from the cervical and the sacral (S2-S4) region of the spinal chord respectively. They have long pre-ganglionic fibers and make synaptic connections in or near the target organs. The vagus nerve provides parasympathetic innervations to the upper GI tract including the stomach, small intestine and proximal colon, while the pelvic nerves provide parasympathetic innervations to the distal colon, rectum and anus. Both the pre- and post-ganglionic neurons of the PSNS release ACh as the neurotransmitter. The PSNS modulates intrinsic innervations to increase peristaltic speed (Gribovskaja-Rupp et al., 2014), blood flow and absorption during “rest and digest” response (Anuras, 1992).   8     1.3.2 Intrinsic innervations The ENS mediates intrinsic innervations to the GI tract. The ENS is an semi-independent neural network that is found with in the walls of the GI tract, and controls GI motility and secretion (Costa, 2000). Although the sympathetic and the parasympathetic input to the GI tract plays a crucial role in modulating GI functions by acting as a relay center between ENS and the CNS, the ENS in itself is capable of integrating and processing GI functions (Anuras, 1992). It is therefore referred to as the “second brain” (M Costa, 2000). The ENS comprises of neurons and glia. Developmentally, enteric neurons and glias are derived from vagal neural crest cells (Young & Newgreen, 2001) and ablation of these cells during development leads to loss of enteric ganglia (Yntema & Hammond, 1954). The ENS in an adult contains more than 400 million neurons (Furness, 2012), whereas enteric glia outnumber enteric neurons by a factor of 4-10 (Neunlist et al., 2007). The enteric glia are irregular stellate shaped cells, which ensheath the neuronal cell bodies and provide regulatory signals for the development, survival and function of enteric neurons in the GI tract (Yu & Li, 2014). Besides enteric neurons and glia, the ENS also contains dense network of interstitial cells of Cajal (ICC). ICC are pacemaker cells of the GI tract that are closely associated with the enteric neurons and the smooth muscles, where they not only modulate the rhythmic contractile activity via slow wave production, but also contributes to transportation, absorption and secretion of fluids in the GI tract (Anuras, 1992; Ordög et al., 1999; Kunisawa & Komuro, 2008). The majority of enteric neurons, glia and ICC in the ENS are arranged among submucosal (Meisssner’s) and myenteric (Aurebach’s) plexus (Furness, 2012), where they work together to regulate GI motility, local blood flow to the smooth muscles, gut barrier function and transmucosal movement of fluids for secretion of digestive enzymes and absorption of nutrients   9     (Furness, 2012; Yu & Li, 2014). Failure of either one of these component of the ENS to properly carry out it role leads to enteric neuropathies such as IBS, slow transit constipation, diabetic gastroparesis, achalasia, Chagas disease, pathogen induced diarrhea, Hirschsprung disease e.t.c (Furness, 2012). Figure 1.3 Organization of the enteric nervous system (ENS). The ENS comprises of two major nerve plexuses, namely, the submucosal plexus (SMP) and the longitudinal muscle myenteric plexus (LMMP). The SMP controls GI secretion while the LMMP controls GI motility. Reprinted from Nature Reviews: Gastroenterol. Hepatology, volume 9, John B. Furness, The enteric nervous system and Neurogastroenterology, p286-294, copyright (2012) with permission from Macmillan. Abbreviations: ENS, enteric nervous system; SMP, submucosal plexus; LMMP, longitudinal muscle myenteric plexus.   10     1.4 Morphological classification of enteric neurons The enteric neurons have varying shapes and can be therefore be classified into three major types based on their morphology namely: Dogiel type I, Dogiel type II and Dogiel type III neurons (Brehmer et al., 1999; Furness, 2007b). 1.4.1 Dogiel type I Dogiel type I neurons have short, broad, and ~ 4-20 laminar dendrites. These dendrites are flattened in myenteric plane and have narrow field of influence, such that, they branch and end within the ganglion of origin (Brehmer et al., 1999). Dogiel type I neurons are 13-35 µm long and 9-22 µm wide (Furness, 2006). They make up most of the inhibitory and excitatory motor neurons in the ENS. Dogiel type I neurons are mono-axonal and the axons continue up to four other ganglia before entering the circular muscle coat (Furness, 2006). Figure 1.4 Schematic diagrams of Dogiel type I neurons example as classified by Dogiel. Reprinted: The enteric nervous system (Book) John B. Furness, copyright (2006).   11     1.4.2 Dogiel type II Dogiel type II neurons are the most prominent neurons in the myenteric and the submucosal plexus of the colon and the small intestine but are absent in the stomach (Brehmer et al., 1999; Furness, 2006). They are multi-axonal and have large cell bodies that are either round or oval in shape. The major diameter and the minor diameter of type II neurons are ≥ 22-47 µm and 13-22 µm respectively (Furness, 2006). These neurons contains ~3-30 dendrites that leave ganglion of origin and have a broader field of influence (Furness, 2006; Sarna, 2010). These cells make up majority of intrinsic primary afferent neurons (IPANs) of the submucosal and the myenteric plexus and show electrophysiological after-hyperpolarization (AH) phenomenon following an action potential (Hodgkiss & Lees, 1983; Tamura, 1992; Brehmer et al., 1999). Figure 1.5 Schematic diagrams of Dogiel type II neurons example as classified by Dogiel. Reprinted: The enteric nervous system (Book) John B. Furness, copyright (2006)   12     1.4.3 Dogiel type III Dogiel type III neurons consist of ~2-10 smooth dendrites that gets thinner with branching distance (Brehmer et al., 1999). Although Dogiel type III neurons end within the ganglion of origin, their dendritic length are comparatively longer than that type I (Brehmer et al., 1999). The axon of a Dogiel type III neuron begins either from small conical protrusion of the cell body, or from a dendrite. Dogiel type III neurons make up a majority of descending interneurons (Furness, 2007b). Figure 1.6 Schematic diagrams of Dogiel type III neurons example as classified by Dogiel. Reprinted: The enteric nervous system (Book) John B. Furness, copyright (2006)   13     1.5 Structural and physiological characteristics of the GI Smooth muscle cells Smooth muscles make up a majority of contractile tissues in the GI tract. In the GI tract smooth muscle cells are functionally coupled via gap junctions. This causes GI smooth muscle to produce synchronous contraction and relaxation (Anuras, 1992). The synchronicity of smooth muscles helps in propulsion and mixing of luminal content, modulation of reservoir capacity (notably in the stomach and colon), and expulsion of fecal matter, pathogens and noxious chemicals from the GI tract (Furness, 2012). 1.5.1 Structure of the GI smooth muscle GI smooth muscles are small (<10µm in diameter) spindle shaped cells that are arranged in bundles or ‘fascicae’. GI smooth muscles contain the contractile actin and myosin proteins to mediate contractions and relaxations (Anuras, 1992). In the smooth muscles these proteins are not arranged in sarcomeres, and therefore appear unstriated. In the smooth muscles, actin filament bundles are anchored to “dense bodies” (α- actinin, an actin binding protein) that are present either on the sarcolemma or on the cytoskeleton of the cell. The smooth muscle cell also contains membranous tubules called sarcoplasmic reticulum, which is a major intracellular calcium (Ca2+) storage site. In the surface of the smooth muscle cell there are small flask shaped involutions called “Caveolae” that protrude ~100 nm into the cells. Caveolae helps to increase the surface area of the plasma membrane and are smooth muscle analogue of skeletal muscle Ttubule system. 1.5.2 Smooth muscle contraction Smooth muscle contractility is primarily mediated via an increase in free intracellular Ca2+ concentrations. At a lower intracellular Ca2+ concentration (i.e < 10-7 M), the smooth muscle fails to contract.   However after depolarization, an increase in intracellular Ca2+ 14     concentration causes smooth muscle contractile actin and myosin proteins to interact with one another resulting in contraction. Depolarizing stimulus leads to an increase in intracellular Ca2+ concentration via an influx of Ca2+ through voltage gated Ca2+ channels or via agonist (neurotransmitters, hormones) induced stimulation of G-proteins (Somlyo & Somlyo, 1994). Stimulation of G proteins increases phospholipase C activity which produces two potent second messengers from the membrane lipid phosphatidylinositol 4,5-bisphosphate, namely: diacylglycerol (DG) and inositol 1,4,5-trisphosphate (IP3) (Webb, 2003). DG in presence of Ca2+ activates protein kinase C (PKC), which later phosphorylates Ca2+ channels and other proteins, to regulate cross-bridge cycling. IP3 on the other hand binds to IP3 receptors on the sarcoplasmic reticulum, causing release of Ca2+ via Ca2+ induced Ca2+ release (CICR) mechanisms (Anuras, 1992). At higher intracellular Ca2+ concentration, Ca2+ ions bind to Ca2+ binding protein called ‘calmodulin’. The Ca2+-calmodulin (CaCM) complex activates Myosin Light Chain Kinase (MLCK). MLCK phosphorylates the myosin head specifically at Ser19 residue of the 20 kDa myosin protein (Somlyo & Somlyo, 1994; Borman et al., 2002). Phophorylation of myosin head causes the myosin head to interact with the actin filament, which initiates cross-bridge cycle, and results in smooth muscle contraction. Since, high intracellular Ca2+ concentration are toxic to the cells, the increase in intracellular Ca2+ concentration is transient. Under low Ca2+ concentration however, the contractile response is maintained by a Ca2+-sensitizing mechanism bought about via activation of G protein ‘Rho A’. Rho A activates Rho kinase which in turn inhibits the activity of a heterotrimeric protein called ‘myosin phosphatase’ (MP)(Webb, 2003).   15   Inhibition of MP   activity causes myosin head to stay phosphorylated, causing prolonged contraction even at lower intracellular Ca2+ levels (Webb, 2003). Figure 1.7 Schematic diagram illustrating smooth muscle contraction. Increase in intracellular Ca2+ directly via opening of voltage gated Ca2+ channels or indirectly via agonist induced G-protein activation leads to formation of CaCM complex. CaCM complex activates MLCK causing myosin head to bind to actin filament resulting in smooth muscle contraction. Rho A activation of Rho-Kinase prevents myosin phosphatase activity causing prolonged contraction. Reprinted from Advances in Physiology Education, volume 27, R. Clinton Webb, Smooth Muscle Contraction and Relaxation, p201-206, copyright (2003). Abbreviations: DG, diacylglycerol; PKC, protein kinase C; MLC, myosin light chain kinase. 1.5.3 Smooth muscle relaxation Removal of a contractile stimulus (agonist stimulation) causes decrease in intracellular Ca2+ concentration via closure of voltage-gated Ca2+ channels and decrease in IP3 mediated Ca2+ release from the intracellular stores. At lower intracellular Ca2+ concentration, CaCM complex is down regulated leading to subsequent down regulation of MLCK activity. Down regulated MLCK is unable to phosphorylate the regulatory head of the myosin light chain (Webb, 2003).   16     Myosin phosphatase (MP) on the other hand catalyzes dephosphorylation of phosphate from the myosin light chain (LC20) and prevents the interaction between the actin and myosin filament causing cessation of contraction (Borman et al., 2002). The Ca2+ concentration is returned back to the normal levels via Ca, Mg-ATPases and Na+/Ca2+ exchangers that are located either on the surface of the SR or on the plasma membrane. Figure 1.8 Schematic diagram illustrating smooth muscle relaxation. Decrease in intracellular Ca2+ directly via closure of voltage gated Ca2+ channels or indirectly via removal of agonist induced G-protein activation leads to disintegration of CaCM complex and activates MP. MP prevents interaction of actin and myosin causing smooth muscle relaxation. The resting Ca2+ concentration is bought back to normal levels via Ca, Mg-ATPase and Na+/Ca2+ exchangers located on the SR and the plasma membrane. Reprinted from Advances in Physiology Education, volume 27, R. Clinton Webb, Smooth Muscle Contraction and Relaxation, p201-206, copyright (2003). Abbreviations: MLC, myosin light chain.   17     1.6 Interstitial cells of Cajal (ICCs) ICC are the “pacemaker cells” of the GI tract, which produce rhythmic slow waves to modulate smooth muscle contraction. Although ICCs were first identified by Santiago Ramon y Cajal in 1893 as neuron like cells, ultrastructural studies later revealed that ICCs shared characteristics that resemble primitive muscle cells or fibroblast like cells (Faussone Pellegrini et al., 1977; Komuro, 1989). For example, they contained numerous elongated processes, rough and the smooth endoplasmic reticulum, made close contact with enteric nerve terminals, smooth muscles, and also formed numerous gap junctions both with each other, and with the smooth muscle cells (Al-Shboul, 2013). ICCs are now known to be mesenchymal cells, which share common developmental pathway with the smooth muscle cells (Fintl & Hudson, 2010). ICCs in the GI tract express receptor tyrosine kinase (c-Kit), which is essential for the function and development of these cells (Sanders et al., 2006). Blockade of c-Kit prevents binding of ligand steel factor stem cells factors (SCF) and impairs ICC development in mouse GI tract (Sanders et al., 1999). 1.6.1 Function of ICCs ICCs are electrically coupled to smooth muscle cells (Cousins et al., 2003). Therefore, currents generated in the ICCs are continuously transmitted to smooth muscle cells to set up the frequency of GI contractions (Sanders et al., 2006). ICC organizes smooth muscle synchronicity and contractile behavior via ‘slow wave’ generation (Sanders et al., 2006; Fintl & Hudson, 2010; Al-Shboul, 2013). Slow waves are continuous rhythmic pacemaker currents that cause oscillation of the membrane potential and alternating periods of high and low smooth muscle excitability. A slow waves consist of a rapid upstroke, longer plateau phase and repolarization of the cell membrane potential (Al-Shboul, 2013). In periods of high excitability the plateau phase   18     of slow waves rises above the threshold for L-type Ca2+ channel activation and leads to action potential generation and smooth muscle contraction (Sanders et al., 2006). Thus, the rhythmicity of slow wave activity in the GI smooth muscles modulates smooth muscle contractility and subsequent GI motility patterns (Fintl & Hudson, 2010; Drumm et al., 2014). 1.6.2 Generation of pacemaker current and slow wave production Mechanisms involved in generation of slow waves are still not entirely understood and the identity of ion channels that generates pacemaker currents in the ICC is controversial (Huizinga et al., 2002; Zhu et al., 2003). A most plausible hypothesis proposes that pacemaker current is initiated at least in part via non-selective cation channels (NSCCs) in the ICCs (Sanders et al., 2006). According to this hypothesis, reduction in intracellular Ca2+ concentration activates NSCCs in the plasma membrane resulting in inward pacemaker currents and cellular depolarization. Cellular depolarization then leads to opening of voltage gated Ca2+ channels (VGCC) and subsequent discharge of IP3 receptors. Ca2+ induced Ca2+ release from the IP3 receptors also stimulates Ca2+ reuptake into the mitochondria. This effectively reduces cytoplasmic Ca2+ concentration causing activation of NSCC and the generation of pacemaker currents (Sanders et al., 2006). Propagation of slow waves into the neighboring ICCs and smooth muscles are achieved via dihydropyridine resistant voltage gated Ca2+ channels (Sanders et al., 2006; Bayguinov et al., 2007). It is now known that besides VGCC and NSCC, Ca2+ activated Cl- channels (CaCCs) are also activated by intracellular Ca2+ and play an important role to generate pacemaker slow wave currents (Zhu et al., 2009). Ca2+ level in the cytoplasmic volume is regulated via smooth endoplasmic reticulum Ca2+ ATPase (SERCA) pumps and mitochondrial Na+/Ca2+ exchangers (Sanders et al., 2006).   19     1.7 Neuromuscular transmission in the GI tract Neuromuscular transmission in the GI tract occurs in the neuromuscular junction. Unlike skeletal muscles, smooth muscle neuromuscular junction lack well defined structural specializations comparable with motor end plate (Hoyle & Burnstock, 2010). Instead, smooth muscle neuromuscular junctions are labile structures with varicosities that are able to move along the length of an axon (Hoyle & Burnstock, 2010). One aspect of this is that autonomic nerves that participate in neuromuscular transmission do not release transmitter solely from terminals, but also from varicosities that occur at intervals of 5-15 µm along axons (Hoyle & Burnstock, 2010). Because of such variations smooth muscle neuromuscular junctions are susceptible to neuromodulatory influence in both pre and post junctional sites leading to exacerbated or diminished neurotransmitter release and uptake (Hoyle & Burnstock, 2010). Neuromuscular transmission in the GI tract plays a major role in regulating GI motility. Two major class of motor neurons that regulate GI motility are the inhibitory and the excitatory motor neurons. These motor neurons make junctional connections with GI smooth muscle cells that are electrotonically coupled by gap junctions to mediate smooth muscle contractions and relaxation. Besides smooth muscles, ICCs also play a major role in mediating neuromuscular transmission (Ward et al., 2006) and smooth muscle excitability (Fintl & Hudson, 2010). Absence of ICC results in loss of excitatory (cholinergic) and inhibitory (nitrergic) neural responses (Ward et al., 2000; Ward et al., 2006). 1.7.1 Inhibitory neuromuscular transmission in the GI tract The current model of inhibitory neuromuscular transmission contains inhibitory motor neurons that release purines (ATP/ β-NAD), (Xue et al., 1999; Mutafova-Yambolieva et al., 2007), nitric oxide (NO) (El-Mahmoudy et al., 2006), vasoactive intestinal peptide (VIP) (El-   20     Mahmoudy et al., 2006) and pituitary activating cyclic AMP (Katsoulis et al., 1996) peptide. These inhibitory neurotransmitter cause transient hyperpolarization of the post junctional smooth muscle cells and takes the membrane potential away from threshold for action potential generation resulting in smooth muscle relaxation. This is referred to as inhibitory junction potential (IJP). The neurotransmitter, receptors and ion channels involved in mediating inhibitory junction potentials are described in greater detail in chapter 2. 1.7.2 Excitatory neuromuscular transmission in the GI tract Excitatory neuromuscular transmission occurs via the actions of excitatory motor neurons. These neurons release excitatory neurotransmitters such as ACh and tachykinins (such as substance P) that either act directly on smooth muscles or indirectly through ICCs (M Costa, 2000) to cause smooth muscle depolarization. This depolarization is referred to as the excitatory junction potentials (EJPs). If the depolarization caused by the EJP is sufficient to overcome the threshold for action potential generation, action potentials will be generated and propagated across individual smooth muscle cells via gap junction causing smooth muscle contraction (Furness, 2007a).   21     Figure 1.9 Excitatory and inhibitory neuromuscular transmission in the GI tract. Balance between excitatory and inhibitory neuromuscular communication is crucial to maintain proper GI motility. Abbreviations: ENS, enteric nervous system; ATP, adenosine triphosphate; β-NAD, β nicotinamide adenine dinucleotide; ACh, acetyl choline; SP, substance p.   22     1.8 GI motility GI motility refers to the process by which the luminal contents are moved across the length of the GI tract. It was earlier shown that segments of the intestine that are either removed from the animal and placed in artificial nutrient solution (Mall, 1896), or have connections to the CNS severed still displayed propulsive motility (Bayliss & Starling, 1899). This indicated that the nerve circuit necessary to produce propulsive motility is present with in the gut wall (Bayliss & Starling, 1899; Langley & Magnus, 1905). Since then, intrinsic peristaltic circuits have been studied in detail in different animals and different regions of the GI tract (Langley & Magnus, 1905; Bülbring et al., 1958; Crema et al., 1970; Thomas & Baldwin, 1971). These studies conclude that the requirement for GI motility is different in a region specific manner and requires varying effort of the enteric neurons, smooth muscles and the interstitial cells of Cajal (ICCs) (Tsuji et al., 1992; Sanders et al., 2006; Shiina et al., 2010; Gudsoorkar & Quigley, 2014; Mañé et al., 2014). For example, in the small intestine extensive stirring and sluggish movement of luminal contents is required for nutrients absorption, while in the colon extensive stirring is not required. Colonic contents instead needs to be moved relatively rapidly than the small intestine so that proper amount of water is absorbed, and the stool is not either over-solidified, or too loose for defecation. To achieve such diverse motility requirements, the enteric neurons, smooth muscles and the ICCs participate differentially to generate three kinds of contractions namely rhythmic phasic contractions, giant Migrating contractions and tonic contractions whose function, spatio-temporal patterns, and regulatory mechanisms vary depending upon the location in the GI tract (Sarna, 2010).   23     1.8.1 Rhythmic phasic contractions (RPCs) RPCs are randomly occurring contraction that do not activate descending relaxation circuitry and therefore propagate only over a short distance. These contractions are regulated via ICC dependent generation of smaller amplitude slow wave, and concurrent activation of excitatory cholinergic motor neurons (Sarna, 2010). The slow waves in the stomach and the small intestine are locked in a phase, which facilitates slow propagation of contractions in these regions. However, the slow waves do not show phase locking in the colon, and therefore exhibit little or no propagation (Sarna, 2010). RPCs are therefore involved in mixing and stirring of GI content for absorption of water and nutrients, but they do not to play a role in propulsive colonic motility (Sarna, 2010). 1.8.2 Giant migrating contractions (GMCs) GMCs have larger amplitude and travel over longer distances to produce propulsion of content. In humans these contractions occur ~2-5 times a day, and helps expel feces by rapidly propagating from the colon to the rectum. GMCs are achieved by activating descending relaxation circuitry, which results in relaxation of the smooth muscles distal to the site of stimulation (Sarna, 2010). Increased GMC frequency results in rapid propagation of colonic content and lower absorption of water and electrolytes by the colonic mucosa, while the lower GMC frequency leads to prolonged contact of feces with colonic mucosa, and higher absorption of water (Bueno et al., 1980). Therefore, disruptions in GMCs have been associated with colonic motor dysfunctions such as diarrhea and constipation. Unlike humans, in rodents, GMCs can easily be recorded from ex-vivo colonic circular smooth muscle strips and rings in organ bath experiments. Rodents therefore serve as an excellent model to study cellular pathways involved in generating GMCs (Fida et al., 1997; Sarna, 2010).   24     1.8.3 Tonic contractions Tonic contractions increase the tone of the smooth muscle cell. The duration of tone depends upon the volume of food ingested. Tonic contractions mostly occur in the small intestine and work with RPCs to narrow lumen size and help in effective propulsion and mixing behaviors (Ford et al., 1995; Sarna, 2010).   25     1.9 Peristalsis The term “Persitalsis” describes the process by which intestinal contents are moved along the length of the GI tract. Bayliss and Starling in 1899 postulated that intraluminal bolus at any point on the GI tract activates a series of reflex circuitry that causes smooth muscle contractions and relaxations rostral (oral) and caudal (aboral) to the bolus (Bayliss & Starling, 1899). Although descending relaxation circuitry is also activated during GMCs, GMCs occur infrequently and travel over longer distances unlike peristalsis, which occurs frequently and occur over shorter distances (Sarna, 2010). 1.9.1 Peristaltic circuit Peristaltic circuit, also known the “myenteric reflex circuit ”, is activated when mechanical and chemical stimuli stimulate the mucosal surface (Hukuhara & Miyake, 1959; Smith & Furness, 1988). Stimulation of the mucosal surface triggers release of serotonin (5-HT) from enterochromaffin (EC) cells (Nozawa et al., 2009). 5-HT release in turn activates the mucosal endings of the IPANs (Bulbring & Crema, 1958; Foxx-Orenstein et al., 1996; Nozawa et al., 2009). IPANS have a distinct Dogiel type II shape and exhibit a long after hyperpolarization after an action potential, which confers directionality to the peristaltic circuit (Furness et al., 1998; Furness, 2007c). IPANs form synaptic connections with interneurons in the myenteric plexus. IPANs release acetylcholine (ACh) onto the interneurons (Furness et al., 1998), which in turn activates myenteric excitatory and inhibitory motoneurons of the LMMP (Costa & Furness, 1976). Finally, excitatory and inhibitory motor neurons release excitatory and inhibitory neurotransmitter in the neuromuscular junction to causes smooth muscle contraction in the oral direction, and relaxation in the aboral direction. Since myenteric plexus wraps around the entire length of the GI tract, the myenteric reflux causes propulsive contraction/relaxation   26     pattern to move along the entire length of the GI tract propelling food from upper esophagus to the internal anal sphincter but only over short segments at a time (M Costa, 2000).   27     1.10 Ca2+-activated potassium (K+) channels Ca2+-activated K+ channels are fundamental regulators of cellular excitability (Petkov et al., 2001; Sah & Faber, 2002; France et al., 2012). Ca2+-activated K+ channels are activated via an increase in free intracellular Ca2+ concentrations, which is bought about via influx of Ca2+ through L-type Ca2+ channels, and CICR mechanisms. Activation of Ca2+-activated K+ channels leads to efflux of K+ ions down the electrochemical gradient causing cellular hyperpolarization (Gardos, 1958). Based on molecular, pharmacological and electrophysiological properties, three distinct classes of Ca2+-activated K+ channels have been identified to date (Vergara et al., 1998). These are: Large conductance Ca2+-activated K+ channels Intermediate conductance Ca2+-activated K+ channels Small conductance Ca2+-activated K+ channels 1.10.1 Large conductance Ca2+-activated K+ channels (BK channel) BK channels also known as SLO1or Maxi-K channels are derived from a single channel gene containing 27 exons, which is found in the 10th and 14th chromosome in humans and mice respectively (Ge et al., 2014). The single channel conductance of BK channels is significantly higher than other Ca2+ activated K+ channels and ranges from 200 to 400 pS (Marty, 1981). BK channels are the only class of Ca2+-activated K+ channels that are voltage sensitive. This is evident by non-linear I-V relationship, which suggests that that the amount of current passing through these channels depends on both the driving force and the probability of channel being open at a particular potential. During an action potential spike, BK channels are activated by depolarization induced opening of voltage gated Ca2+ channels and subsequent activation of CICR mechanisms (Faber & Sah, 2003). At intracellular Ca2+ concentrations ≥ 1µM, BK   28     channels are activated, leading to efflux of K+ from the cell. Since BK channels are activated with a delay from peak of action potential spike, they contribute to spike repolarization (the falling phase of action potential), and generation of fast after-hyperpolarization (fAHP) (Storm, 1987a, b). Thus, BK channels not only control the duration of action potential but also the action potential firing frequency. Structurally, BK channels are composed of four alpha (α) and four 4 beta (β) subunits. The α-subunit of a BK channel is encoded by a single ‘slo’ gene. The α subunit contains eleven hydrophobic domains (S0-S10) (Ledoux et al., 2006). Out of eleven hydrophobic domains, S0S6 domains are transmembrane domains (TMD) that make up the core of the channel. The S4 domain contains several positively charged arginine residues that confer voltage sensitivity and channel gating. The TMD S5-S6 forms the pore of the channel. Site directed mutagenesis studies have revealed that intracellular domain of the α subunit (S7-S10) contains a negatively charged aspartate amino acids residues. These residues form a Ca2+ bowl and regulator of conductance for K+ (RCK) regions to act as a Ca2+ binding site (Vergara et al., 1998; Ledoux et al., 2006). The β-subunit in the BK channel on the other hand contains two TMD with intracellular amino and carboxy-terminus. Although α subunits determine Ca2+ sensitivity, β subunits is responsible for translation of physiologic Ca2+ signals (Ledoux et al., 2006). Thus, the β-subunit serves to modulate the Ca2+ sensitivity of the α-subunits. Genetic knockout of β subunits reduces Ca2+ sensitivity of the channel and disrupts BK channel function in smooth muscle and kidneys (Brenner et al., 2000; Grimm & Sansom, 2010). Until now four different β-subunit genes have been identified (β1- β4). BK β1-subunit is found in the smooth muscle cell (SMC) and in the distal nephron of murine kidney (Grimm et   29     al., 2007), BK β2- and β3-subunits are expressed in endocrine cells (Xia et al., 1999; Braun et al., 2008), while BK β4-subunits are expressed in neurons (Meera et al., 2000) and in the distal nephron of the kidney (Grimm et al., 2007). The pharmacologic antagonists for BK channels include iberiotoxin, charbydotoxin and paxilline (Galvez et al., 1990; Knaus et al., 1994) 1.10.1.1 β1 BK channels The β1BK channels are mostly smooth muscle cell specific (Brenner et al., 2000). β1subunit expression in the BK channel is essential to regulate the Ca2+ sensitivity of the α-subunit. β1 subunit therefore plays a crucial role in regulating resting membrane potential and the excitability of smooth muscle cells (Vergara et al., 1998; Ledoux et al., 2006). Loss or downregulation of BK β1-subunit expression causes membrane depolarization and increased muscle contractility which leads to various disorders including hypertension (Yang et al., 2013), heart failure (Wan et al., 2013), asthma (Semenov et al., 2011), urinary bladder overactivity (Petkov et al., 2001) and gastrointestinal motility disorders (France et al., 2012). Figure 1.10 Large conductance Ca2+-activated K+ (β1BK) channel structure. (A) β1BK channel consists of the α and the β1 subunit. (B) The α subunit contains eleven hydrophobic domain among which seven (S0-S6) are transmembrane domain. The β1 subunit consists of two transmembrane domain with intracellular amino and carboxy terminus. Reprinted from Physiology, volume 21, Ledoux et. al., Ca2+-Activated Potassium Channels and the Regulation of Vascular Tone, p69-78, copyright (2006). Abbreviation: RCK, regulator of conductance for K+ .   30     1.10.2 Small conductance Ca2+-activated K+ channels Unlike the BK channels, the SK channels are purely Ca2+-gated channels (voltageindependent). Structurally, SK channels comprises of four α-subunit (Vergara et al., 1998; Ledoux et al., 2006). Each α-subunit contains six (S1-S6) TMD among which TMD S5-S6 forms the pore of the channel. SK channels do not have a Ca2+ bowl region, instead they contain ‘calmodulin’, a Ca2+-sensitive protein, which confers higher intracellular Ca2+-sensitivity than BK channels (Ledoux et al., 2006). Although positively charged S4 TMD is present in SK channels, the I-V relationship for SK channel is linear, which suggests that gating properties of SK channel is exclusively controlled by Ca2+-bound calmodulin proteins (Sah & Faber, 2002). SK channels have a smaller single channel conductance than BK channels (4-20pS). SK channels contribute to slow after hyperpolarization (sAHP), and limits action potential firing (Ishii et al., 1997). SK channels are expressed almost ubiquitously in CNS, nonvascular smooth muscle cell, and the endothelium. Suppression of SK channels inhibits vasodilation and increase blood pressure (Ledoux et al., 2006). Figure 1.11 Small conductance Ca2+-activated K+ (SK) channel structure. (A) SK channel contains six transmembrane domains. TMD S5-S6 acts as a pore of a channel. A Ca2+ sensitive protein, calmodulin, interacts with the channel’s intracellular COOH terminus. (B) Binding of Ca2+ to calmodulin opens the gate of the SK channel. Reprinted from Physiology, volume 21, Ledoux et. al., Ca2+-Activated Potassium Channels and the Regulation of Vascular Tone, p69-78, copyright (2006).   31     1.10.3 Intermediate conductance Ca2+-activated K+ channels The IK channel was first identified in the red blood cells as the Gardos channel (Gardos, 1958). Since then the IK channels is found to be expressed in multiple cell types including the immune cells, endothelial cells, and neuron of the CNS and PNS (Ishii et al., 1997; Turner et al., 2014). The IK channels are hetero-tetrameric channels. The IK and the SK channels arise from the same gene family but only share ~50% amino acid sequence homology (Ishii et al., 1997). Like the SK channels, the IK channels contain calmodulin, and is activated only by increase in intracellular Ca2+ concentration (Turner et al., 2014). The single channel conductance of IK channels are, however, higher than SK channels (20-85 pS) (Ishii et al., 1997). Pharmacologically, IK channels differ completely from SK and BK channels as they are reversibly blocked by charybdotoxin and clotrimazole but not by apamin and iberiotoxin (Ishii et al., 1997). The major characteristics feature of these channels is summarized in table below.   32     Table 1.1 Electrophysiological and pharmacological characteristics of Ca2+-activated K+ channels   33     1.11 Obesity and its associated disorders Accumulation of body fat leads to obesity. In humans, obesity is defined as a body mass index (BMI) of 30 kg/m2 (body weight in kilograms divided by height in meters squared). High fat (HF) diet consumption is a major cause of obesity in humans (Y Lee, 2013). Diet-induced obesity (DIO) in turn is a risk factor for Type 2 diabetes (T2D) (Colditz et al., 1995), cardiovascular diseases (Huang et al., 1998), gall bladder diseases (Stampfer et al., 1992), degenerative arthritis (Heliövaara et al., 1993), cancer (Garfinkel, 1985; De Nunzio et al., 2011) and GI motility disorders (Pecora et al., 1981; vd Baan-Slootweg et al., 2011). In 2011, ≥ 284 million people worldwide were obese with T2D (Farag & Gaballa, 2011). By 2030, ~440 million people (8% of the world population) are expected to be affected driving health care expenditures to exceed $490 billion (Shamseddeen et al., 2011). DIO therefore presents a significant challenge as it is a major health care concern worldwide. 1.11.1 High fat (HF) diet-induced obesity (DIO) and disruption in colonic motility HF DIO and T2D patients exhibit elevated free fatty acids level and increased susceptibility to GI motility complications (Mushref & Srinivasan, 2013; Voss et al., 2013). Approximately, 75% of patients suffering from obesity and obesity-associated disorders such as T2D experience nausea, bloating, diarrhea, constipation, gastroparesis, abdominal pain, and heartburn (Delgado-Aros et al., 2004; Xing & Chen, 2004). DIO mice that were fed 12 weeks of HF diet showed accelerated colonic transit time and enhanced cholinergic and serotonergic signaling in the ENS (Reichardt et al., 2013), while 4-6 months of HF feeding caused substantial loss of myenteric neurons and increased intramuscular lipid accumulation in the ileum and the colon (Voss et al., 2013; Beraldi et al., 2014). Among myenteric neurons, inhibitory motor neurons are significantly impaired in mouse model of HF DIO and T2D (Chandrasekharan &   34     Srinivasan, 2007; Rivera et al., 2011; Mushref & Srinivasan, 2013). Colon specimens from patients with T2D exhibit oxidative stress concurrent with loss of myenteric neurons (Chandrasekharan et al., 2011a). However, how HF diet translates to colonic dysmotility in obesity is not yet entirely understood. In chapter 3, I studied the pathogenesis of HF diet associated colonic dysmotility in great details.   35     1.12 Oxidative stress Oxidative stress is an imbalance between production of free radicals and antioxidants such that the production of free radicals overwhelms the production of antioxidants. Important sources of oxidative stress include the mitochondria respiratory cycles, nicotinamide adenine dinucleotide phosphate oxidase (NADPH oxidase) formed in immune cells, and enzymes such as fatty acyl CoA oxidase, monoamine oxidase and xanthine oxidase (Baggiolini & Wymann, 1990; Simonson et al., 1993; Fridovich, 1995; Murphy et al., 1998; Marikovsky et al., 2003; Thayer et al., 2011). In the mitochondria ~4% of the electrons that are released from energy molecules such as NADH and FADH2 lead to the formation of superoxides (O2-) and do not participate in ATP production (Fridovich, 1995). Under normal circumstance, reactive oxygen species (ROS) such as O2- levels are tightly regulated via antioxidants such as superoxide dismutase (SOD), catalase and reduced glutathione (GSH). In the cell, SOD catalyzes dismutation of O2-, converting O2into hydrogen peroxide (H2O2) while GSH further reduces H2O2 to H2O. However, when the electron transport chain is chronically or acutely overloaded in order to break down glucose and fats, ROS production is increased either through mitochondrial uncoupling or via β oxidation. This causes oxidation of inner mitochondrial membrane and disruption of ATP synthesis leading to even further increase in ROS generation (Vincent et al., 2004). In addition to increased ROS in obesity and T2D, antioxidants such as catalase and GSH are reduced under hyperglycemic conditions causing increased H2O2 levels in cells (Lu et al., 2006). ROS including H2O2 not only affects the host cell, but also diffuses across the plasma membrane to affect the neighboring cells (Vincent et al., 2004).   36     1.12.1 Effects of oxidative stress on biomolecules 1.12.1.1 Nucleic acids ROS induced oxidation of DNA nucleotides gives rise to 8-hydroxydeoxyguanosine, whereas oxidation of RNA nucleotides forms 8-hydroxyguanosine. In the DNA, 8- hydroxydeoxyguanosine is mutagenic and therefore carcinogenic while in the RNA, 8hydroxyguanosine causes reduced protein translation capacity (Bar-Or et al., 2015). 1.12.1.2 Lipids ROS oxidizes long-chain fatty acids in the cell membrane to form peroxidation products such as malondialdehyde (MDA). MDA react with amines (that are present in the proteins) to generate another toxic compound called aldehyde, 4-hydroxynonenal (HNE) (Bar-Or et al., 2015). These products are not only toxic to neuronal membrane and impair neurotransmitter release and uptake but they also impair mitochondrial function at the synapse (Keller et al., 1997). 1.12.1.3 Proteins Proteins are readily oxidized by free radicals at the amino terminus, carboxy terminus or at side chains to produce peroxides, keto-acids, and other oxidation derivatives (Bar-Or et al., 2015). This causes protein degradation and misfolding which is commonly observed in Alzheimer's disease and Amyotrophic lateral sclerosis (ALS). Protein oxidation is also a major signal for facilitation of ubiquitin attachment. Ubiquitin, a 8.5 kDa regulatory protein, is a major signal for protein degradation by proteosomes (Iwai et al., 1998).   37     Figure 1.12 Model describing mechanisms of oxidation of biomolecules. 1.12.2 Obesity and oxidative stress Obesity is identified as a major risk factor for development of various metabolic disorders including T2D, hyperglycemia, dyslipidemia, and hypertension. Although the mechanistic relationship between obesity and its associated disorders is not completely understood, oxidative stress induced free radical formation is regarded as key player in pathogenesis of obesity-associated disorders. Emerging studies indicate that increased oxidative stress in accumulated fat is at least in part the underlying cause of development of GI dysmotility in obesity (Furukawa et al., 2004). In obese mice, H2O2 generation is selectively increased in adipose tissue accompanied by increased NADPH oxidase mRNA expression, and decreased expression of SOD, catalases and GSH (Furukawa et al., 2004). Treatment with NADPH oxidase inhibitor in obese mice reduces   38     ROS production in adipose tissue and improved diabetes and hyperlipidemia suggesting that increased ROS generation in accumulated fat is an early instigator of metabolic syndrome in obesity (Furukawa et al., 2004). Figure 1.13 Model indicating how accumulated fat in obesity leads to obesity associated disorders. 1.12.3 Impact of oxidative stress on inhibitory nitrergic signaling in the GI tract ROS such as O2•- and H2O2 are formed continually through biological reactions, however, they are not reactive enough to cause large scale tissue damage (Halliwell & Gutteridge, 1989). Early on, most oxidative damage was ascribed to the highly reactive hydroxyl radical (•OH), which is formed as a result of Fenton reaction from H2O2 (Halliwell & Gutteridge, 1989). However this hypothesis is controversial for two reasons. First, •OH radical is extremely reactive and therefore can only diffuse few angstrom before reacting with a target tissues, and second, a much higher concentration of free metal (iron) and H2O2 than that is available in an invivo condition is required to cause large scale oxidative damage and subsequent biological disorders (Beckman, 1994; Crow & Beckman, 1996).   39     In GI tissues another highly reactive free radical called peroxynitrite anion (ONOO-) is involved in causing oxidative stress (Murphy et al., 1998; Rivera et al., 2011). ONOO- is more potent oxidant than H2O2 or O2- and can together oxidize proteins, lipids and DNA (Chandrasekharan et al., 2011b). ONOO- anions are produced by reaction between NO and O2•anion (Murphy et al., 1998). In the human GI tract, NO is synthesized in inhibitory NO neurons via action of Ca2+ sensitive enzyme called neuronal nitric oxide synthase (nNOS). nNOS catalyzes conversion of arginine into NO, a diffusible inhibitory neurotransmitter. NO once released into the neuromuscular junction can diffuse into the post junctional smooth muscle cells to control inhibitory neuromuscular transmission and cause smooth muscle relaxation (Matsuda & Miller, 2010). Since NO passively diffuses from nNOS neurons to smooth muscle cells both nNOS neurons and smooth muscle cells are vulnerable to oxidative stress. NO and ONOOanions oxidizes proteins, and cause post-translation modification, either by binding specifically to cysteine residue (to form nitrosothiols) or by reacting with tyrosine residues in the protein to produce 3-nitroso derivatives (tyr3 nitrosylation) (Rivera et al., 2011).   40     Figure 1.14 Mechanism of peroxynitrite (ONOO-) production in the inhibitory nitric oxide (NO) neurons. NO scavenges superoxide anions in the inhibitory nNOS neurons to produce peroxynitrite (ONOO-). ONOO- is extremely potent than NO or O2- and causes oxidation of biological molecules including (1) nucleic acids (2) lipids and (3) proteins. 1.12.4 Impact of oxidative stress on β1BK channel function The β1BK channel is present throughout the smooth muscle and plays a crucial role in regulating the excitability of the vascular, GI and urinary bladder smooth muscle cells (Vergara et al., 1998; Petkov et al., 2001; France et al., 2012). The β1 subunit achieves this function and modulates BK channel activity by enhancing apparent Ca2+ sensitivity of the pore forming α subunit and also by slowing activation / deactivation kinetics (Cox & Aldrich, 2000; Qian & Magleby, 2003). In vascular smooth muscle cell, ROS preferentially attacks Ca2+ sensing cysteine residue leading to disruption of Ca2+ sensing mechanisms and causing the channel to behave as a β1 subunit KO (Tang et al., 2004). Furthermore, β1 subunit is also amplifies oxidative regulation of the β1BK channel (Santarelli et al., 2004). In rat cerebral arteries ONOO-   41     decreases BK channel open probability in a dose dependent fashion, and this effect is reversed by addition of GSH (Brzezinska et al., 2000). Cerebral arteries of diabetic mice also exhibit decreased BK channel β1/α subunit ratio, and altered Ca2+ spark generation (Rueda et al., 2013). This results is similar to the results from cerebral artery of β1KO mice which exhibits altered Ca2+ spark/spontaneous transient outward current coupling (Plüger et al., 2000) suggesting that β1BK channels from diabetic mice behaves similar to β1KO mice. Besides this, low antioxidants production as seen in HF mouse model (Furukawa et al., 2004), also exacerbates oxidation of BK channels (Tang et al., 2004; Li et al., 2011). These studies suggest that oxidative stress is a key modulator of β1BK channel expression and function. Figure 1.15 Mechanism of disruption of β1BK channel function via oxidative stress. Neuronal and non-neuronal NO (produced via action of enzymes like xanthanine oxidoreductase and reaction of H2O2 with arginine) production results in increased ONOO- formation in the smooth muscles. ONOO-, H2O2 and O2- oxidizes amino acids within the β1BK channel and affects its function and expression.   42     1.13 BK channel and Inhibitory nitric oxide signaling Exogenous NO activates BK channels in the smooth muscle cells (Mistry & Garland, 1998; L'Heureux et al., 2010). Activation of BK channel by NO occurs via cyclic GMPdependent protein kinase G (PKG) activation, and via S-nitrosylation of cysteine sulfhydryl residue of the BK channel (George & Shibata, 1995; Mistry & Garland, 1998; Jaffrey et al., 2001; Hess et al., 2005). In the rat hippocampus NO donor, SNP, restored BK channel activity in nNOS deficient neurons caused by intermittent hypoxia induced oxidative stress (Tjong et al., 2008). Besides this, NO is also involved in rapid anterograde trafficking of β1 subunit from the endosomes to the cell membrane in order to helps associate itself with BKα subunit. NO thus causes elevated Ca2+ sensitivity and activity in myocytes (Leo et al., 2014). My studies (discussed in chapter 3) also show that lower NO availability in DIO translate to altered smooth muscle excitability in DIO. Taken together these results suggest that NO and BK channel function are intricately linked.   43     Figure 1.16 Mechanism of NO production and BK channel activation in the neuromuscular junction. Activation of arginine via nNOS leads to production of NO. NO passively diffuses in the cell and causes activation of soluble guanylyl cyclase (GC), leading to increased cyclic GMP production and activation of protein kinase G (PKG). (1) PKG phosphorylates BK channel and affect channel function. (2) In addition, NO also oxidizes protein thiol to affect BK channel function.   44     1.14 Hypothesis and specific aims The overall aim of this research was to identify the pathogenesis of colonic dysmotility in obesity. My overall hypothesis is that HF diet leads to oxidative stress induced loss of inhibitory nNOs neurons, and impaired β1BK channel function/expression. This causes impaired neuromuscular transmission and smooth muscle excitability and ultimately leads to colonic dysmotility. 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I studied inhibitory junction potentials (IJPs) in the mouse colon caused by electric field stimulation (EFS; 0.2 s train, 10 Hz, 90-100 V). IJPs were biphasic consisting of fast initial hyperpolarization (fIJP) followed by sustained slow hyperpolarization (sIJP). MRS 2179 (10 µM), a purinergic P2Y1 receptor antagonist, blocked fIJPs but not sIJPs. L-NNA (100 µM), a nitric oxide synthase inhibitor, blocked sIJPs but not fIJPs. MRS 2179 plus L-NNA blocked fIJPs and sIJPs. Fast and sIJPs were inhibited by apamin and niflumic acid (NFA), a SK channel and Ca2+ activated Cl(CaCCs) channel blockers respectively. ATP, β–nicotinamide adenine dinucleotide (β–NAD) and adenosine 5′-[β-thio]diphosphate (ADP-β-S) caused smooth muscle hyperpolarization. ATP produced larger hyperpolarizations than β–NAD. Polyoxotungstate 1 (POM-1), an ecto- nucleotidases inhibitor, blocked fIJPs and ATP response but not ADP-β-S responses. ADP-β-S responses were blocked by MRS 2179. These data suggests that IJPs are mediated via ATP acting on P2Y1 receptors and NO, both of which activates SK channels and CaCCs channels to cause relaxation of colonic smooth muscle.   59     2.2 Introduction Disturbances in colonic smooth muscle relaxation leads to constipation, bowel incontinence and other gastrointestinal (GI) motility disturbances (Engel, 2004). Colonic smooth muscle relaxation occurs via the release of inhibitory neurotransmitters from the non-adrenergic, non-cholinergic (NANC) inhibitory motor neurons. Action of inhibitory neurotransmitters on smooth muscle produces a hyperpolarizing response, known as the inhibitory junction potential (IJP). IJPs are electrophysiological correlate of smooth muscle relaxation. IJPs are mediated by purinergic, and nitrergic (nitric oxide mediated) pathways (Serio et al., 2003; Mañé et al., 2014). Purinergic signaling involves activation of post-junctional P2 receptors on the smooth muscles to mediate fast IJPs (fIJPs) (Zhang et al., 2010) whereas the nitrergic pathway requires nitric oxide (NO) action on smooth muscle cells to mediate slow IJPs (sIJPs) (Xue et al., 1999; Lies et al., 2014). Purinergic and nitrergic pathways activate different ion channels on smooth muscles to mediate fIJPs and sIJPs. Although small conductance Ca2+activated K+ channels (SK channel) contribute to IJP production, the contribution of other channels to IJPs is unclear (Zhang et al., 2010; Martínez-Cutillas et al., 2014). The Cl- channel blocker 4,4’ – diisothiocyanostilbene- 2,2’ disulfonic acid (DIDS), blocks IJPs in guinea pig ileal circular smooth muscle cells, indicating that IJPs were partly due to closure of Cl- channels (Crist et al., 1991). In a more recent study, a more selective Ca2+- activated Cl- channel (CaCCs) blocker, niflumic acid (NFA; 200 µM) blocked the sIJPs but not the fIJP in opossum esophageal smooth muscle (Zhang & Paterson, 2002). The contribution of CaCC to IJPs in other tissues is unclear. P2Y1 receptors mediate fIJPs, but the exact identity of the purinergic neurotransmitter is controversial. Functional studies suggests that ATP is the inhibitory purine neurotransmitter in   60     the gut (Xue et al., 1999). α,β-meATP, a stable analogue of ATP, closely mimics IJPs or muscle relaxation in the human and the rat colon (Martínez-Cutillas et al., 2014). However, other studies suggest that ATP does not mimic the effect of a neurotransmitter (Hwang et al., 2011). Due to these controversial reports, investigators continue to refer to the actual neurotransmitter as a purine substance. β–nicotinamide adenine dinucleotide (β–NAD), a purine molecule, may be the purine neurotransmitter as it is released in response to nerve stimulation in the murine vascular and visceral smooth muscle cells and exogenous application of β–NAD mimics IJPs (Smyth et al., 2004; Mutafova-Yambolieva et al., 2007; Goyal, 2011). In addition, electrical stimulation of human and non-human primate colon evoked co-release of ATP and β–NAD (Hwang et al., 2011). The release of β–NAD was neuronally mediated as opposed to ATP release (Hwang et al., 2011). I attempted to address this controversy and evaluate whether ATP or β–NAD mediate IJPs. One way to resolve this issue is by evaluating the effects of ectonucleotidase (NTPDase) inhibitors on GI smooth muscle responses caused by local and brief application of ATP and ADP to mimic IJPs. NTPDase specifically metabolizes ATP into ADP, which is a primary agonist for post-junctional P2Y-receptors (Müller et al., 2006; Wall et al., 2008). Inhibition of NTPDase can help to identify the neurotransmitter involved in inhibitory purinergic neuromuscular signaling.   61     2.3 Materials and methods 2.3.1 Ethics statement All studies were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Publication No. 85-23, revised 1996) and approved by Institutional Animal Care and Use Committee of Michigan State University. 2.3.2 Animals Male C57BL/6J mice, 10-12 weeks old were obtained from Jackson laboratories (Bar Harbor, ME, USA). Animals were kept in 12/12 hr light/dark cycle and had access to food and water ad libitum. Mice were euthanized by isoflurane anesthesia followed by cervical dislocation. 2.3.3 Electrophysiological recordings The distal colon segment was quickly removed and placed in carboxygenated Kreb’s solution and transferred to a petri dish. The mesenteric fat was removed and the segment was opened along the mesenteric border. The tissue segment was stretched and pinned flat on the Sylgard® base with fine pins (0.1 mm) (Fine Science Tools, Foster City, CA). The chamber was placed on the stage of an inverted microscope and was immobilized using modeling clay. Carboxygenated Krebs solution heated to 37°C was continuously flowed through the chamber at the rate of 3-4 ml/min. Drugs were added at a known concentration to the flowing Krebs solution using a system of 3-way stopcocks. The circular smooth muscle cells were impaled with glass microelectrode filled with 2 M KCl (40 - 80 MΩ tip resistance). Membrane potential was recorded using an Axoclamp 2A amplifier, a digidata 1332A analog-digital converter, and axoscope 9.2 software (all from Molecular Devices, Sunnyvale, CA). Electrical field stimulation was applied using two parallel silver wires placed 3.5 cm apart (Ø=0.03", A-M systems,   62     Carlsborg, WA). The wires were placed perpendicular to the longitudinal axis of the preparation. The electrical field stimulation (EFS) had following parameters: total train duration, 200 ms; Frequency, 10 Hz; pulse duration 0.8 ms; stimulation voltage 90-100 V. The amplitude and the area of EFS induced inhibitory junction potential (IJP) were measured before and after the infusion of each drug. The Ca2+ channel blocker nifedipine (1µM) was added to abolish spontaneous muscle contraction and obtain stable impalements for neurogenic recordings. 2.3.4 Local drug applications ATP, Adenosine 5′-[β-thio]diphosphate (ADP-β-S) and β–nicotinamide adenine dinucleotide (β-NAD), were loaded into the micropipette (~10 µm tip diameter) positioned close to the recording site where agonists were applied by pressure ejection (5-10 psi, 20 ms puff duration) (Pico-spritzer; General Valve, East Hanover, NJ). The amplitude and the area under the curve of drug-induced responses were measured. 2.3.5 Solution and Drugs Krebs solution composition was as follows (mM): Glucose, 11; NaCl, 117; NaHCO3, 25; KCl, 4.7; NaH2PO4.H2O, 1.2; CaCl2. 2H2O, 2.5 and MgCl2. 6H2O, 1.2 (pH 7.3–7.4). The Krebs solution was bubbled with 95% O2 and 5% CO2 mixture. All reagents were obtained from J.T. Baker (Phillipsburg, NJ). The following drugs were used: nifedipine (1 µM), Nω-nitro-Larginine (L-NNA, 100 µM), 2′-deoxy-N6-methyladenosine 3′,5′-bisphosphate tetrasodium salt (MRS 2179, 10 µM), apamin (0.1 µM), paxilline (0.5 µM), niflumic acid (50 µM), adenosine 5’triphosphate disodium salt solution (ATP, 5 mM), sodium polyoxotungstate 1 POM-1 (10 µM), adenosine 5′-[β-thio]diphosphate trilithium salt (ADP-β-S; 5 µM) and β-nicotinamide adenine dinucleotide hydrate (β-NAD, 5 mM). Stock solutions of nifedipine and niflumic acid were prepared by dissolving in dimethyl sulfoxide (DMSO). Stock solution of L-NNA was prepared   63     by dissolving in 1N HCl. Stock solutions for all of the other drugs were prepared by dissolving the drug in distilled water. POM-1 was obtained from Tocris Bioscience (Bristol, UK) while the rest of the drugs were obtained from Sigma-Aldrich (St. Louis, MO, USA). 2.3.6 Statistics Data were analyzed only from individual recordings that lasted ≥2 min. Normalized responses to the drug were measured by evaluating both the amplitude and the area of EFSinduced IJP responses. Effects of drug on the amplitude and area of IJP were compared using paired t-test. Two-group comparison was made using two-way ANOVA with Bonferroni posthoc test. Data are mean ± SEM. Differences were considered significant when p<0.05. Analysis was done in GraphPad Prism 4.0 software (San Diego, CA).   64     2.4 Results 2.4.1 Nifedipine application does not affect Resting membrane potential (RMP) in the mouse colon Nifedipine (1 µM) did not change the RMP of the colonic smooth muscle cells. This indicates that L-type Ca2+ channels do not play a vital role in mediating resting membrane in mouse colonic smooth muscles. Nifedipine can therefore be used to obtain stable impalements to record IJP in the mouse distal colon. Figure 2.1 Effect of nifedipine on colonic smooth muscle resting membrane potential. Nifedipine (1 µM) did not change the resting membrane potential (RMP) of colonic circular smooth muscle cells (n=6, p>0.05). 2.4.2 IJPs display both uniphasic and biphasic patterns When electrical field stimulation (10 Hz, 0.2 s) was used to elicit an IJP, two types of IJPs were identified. An initial fIJP characterized by larger amplitude and shorter time duration followed by a smaller amplitude and longer duration slow IJP (sIJP). In some instances fIJP and the sIJP merged to form a uniphasic IJP. The amplitude and the area of the IJP were used as a measure of fIJP and sIJP respectively.   65     Figure 2.2 IJPs have both a uniphasic and biphasic patterns. IJPs were obtained in (A) absence and in presence (B and C) of nifedipine (1µM). The shapes of IJP are either uniphasic (A and C) or biphasic. (B) Biphasic IJP consists of shorter duration fast IJP followed by a longer lasting slow IJP. NS: nerve stimulation.   66     2.4.3 IJPs are neurogenic In order to verify the neuronal origin of the IJP, tetrodotoxin (TTX), a voltage gated sodium (Na+) channel blocker was used. TTX blocks axonal action potential (Bane et al., 2014). This impairs neurotransmitter release in the neuromuscular junction. TTX completely blocked IJPs confirming a neuronal response. Figure 2.3 Schematic diagram illustrating neuromuscular transmission in the mouse colon. (A) Electrical stimulation opens voltage gated Na+ and K+ channels causing action potential generation in the inhibitory motor neurons. Action potential upon reaching the synaptic terminal activates Ca2+ entry via voltage gated Ca2+ channel causing neurotransmitter release in the neuromuscular junction. Post-junctional hyperpolarizing response is then measured from the smooth muscles via sharp microelectrode connected to a differential amplifier. (B) Tetrodotoxin (TTX; 0.3µM) blocks voltage gated Na+ channels and completely abolishes IJP in mouse distal colon.   67     2.4.4 Identification of receptors and neurotransmitters that mediate IJPs in the mouse colon In order to identify the receptors that mediate IJP in the mouse colon I applied MRS 2179, a P2Y1 receptor antagonist, on colonic smooth muscles (Figure 2.4). The RMP of the smooth muscle cells was unaffected by MRS 2179. MRS 2179 reduced IJP amplitude by 66 ± 4% but did not reduce the IJP area. This suggests that the P2Y1 receptor mediates the fIJP but not the sIJP. Next I studied the effect of L-NNA on evoked IJP (Figure 2.5). Nω-nitro-Larginine (L-NNA), a NOS inhibitor, did not affect the smooth muscle RMP. L-NNA did not reduce the amplitude of IJP but reduced the IJP area by 33 ± 8% (*p<0.05). This suggests that NO mediates the sIJP but not the fIJP. Finally, I co-applied MRS 2179 and LNNA together (Figure 2.6). When MRS 2179 and L-NNA were co-applied both the amplitude and the area of evoked IJP was inhibited by 89 ± 6% and 88 ± 8% respectively. This suggests that IJPs in the mouse distal colon are mediated via P2Y1 receptors and NO mediated pathways. Figure 2.4 The effect of P2Y1 receptor blocker on evoked IJPs. (A) MRS 2179 (10 µM) did not change the RMP. (B) Time course of inhibition of normalized IJP amplitude and area after MRS 2179 application. MRS 2179 shows maximum inhibition effect after 4 min of application. (C) Five minutes of MRS 2179 application significantly reduces the IJP amplitude but not the area (n=6, *p<0.05).   68     Figure 2.5 The effect of NOS inhibition on evoked IJPs. (A) L-NNA (100 µM), a NOS inhibitor, did not change the resting membrane potential. (B) Time course of L-NNA effect on normalized IJP amplitude and area after L-NNA application. (C) Five min of L-NNA application significantly reduced IJP area but not the amplitude (n=6, *p<0.05). Figure 2.6 Effects of MRS 2179 and L-NNA co-application. (A) MRS 2179 and L-NNA coapplication did not change the RMP. (B) Time course of MRS 2179 and L-NNA co-application on normalized IJP amplitude and area. MRS 2179 was applied for four min followed by coapplication of MRS 2179 and L-NNA. (C) MRS 2179 and L-NNA co-application blocked both the amplitude and the area of the IJP (n=8, *p<0.05).   69     2.4.5 Identification of smooth muscle ion channels involved in mediating IJPs. The SK channel blocker, apamin, was used to determine if SK channels are involved in mediating IJPs (Figure 2.7). Apamin application depolarizes the membrane potential by 4 ± 1.7 mV, and inhibits both the amplitude and the area of IJP by 40 ± 8% and by 35 ± 6% respectively. I used niflumic acid (NFA), to examine the role of CaCCs in inhibitory neuromuscular transmission (Figure 2.8). NFA hyperpolarized the membrane potential by 5 ± 2 mV and reduces both the area and the amplitude of the IJP by 44 ± 6% and 52 ± 6% respectively. I also co-applied apamin and NFA on the tissue (Figure 2.9). Co-application of NFA with apamin abolished IJPs mimicking the action of MRS 2179 and L-NNA co-application. These data suggests that IJPs are mediated via SK and CaCCs in the mouse distal colon. Figure 2.7 Effects of SK channel blocker on evoked IJPs. (A) Apamin (0.1 µM) application significantly depolarizes the colonic smooth muscle. (B) Time course of apamin effect on normalized IJP amplitude and area. Apamin shows maximum effect after 3 min of application. (C) Five minutes of apamin application significantly reduced both the area and the amplitude of the IJP (n=6, *p<0.05).   70     Figure 2.8 Effects of CaCC blocker on evoked IJP. (A) NFA (50 µM), a CaCC blocker, hyperpolarized the membrane potential by 5 ± 2 mV. (B) Time course of NFA effect on normalized IJP amplitude and area. The inhibitory effects of NFA on IJPs peaks after six minutes of application. (C) NFA application for six minutes significantly reduces both the area and the amplitude of the IJP (n=6, *p<0.05) Figure 2.9 Effects of apamin and niflumic acid (NFA) co-application. (A) Apamin and niflumic acid co-application did not affect the resting membrane potential. (B) Time course of NFA and apamin co-application on normalized IJP. NFA (50 µM) applied for four min followed by co-application of NFA and Apamin blocks IJPs. (C) Both the amplitude and the area of the IJP is disrupted (n=6, *p<0.05).   71     2.4.6 Identification of specific purinergic neurotransmitter involved in mediating the IJP. It is controversial whether the ATP or β-NAD mediates IJPs in the GI tract. Therefore, in order to identify the neurotransmitter mediating the IJP, ATP and β-NAD were applied from a micropipette near the recording site and electrophysiological responses were measured. ATP produced larger significantly larger hyperpolarizations compared to β-NAD (Figure 2.10). ADP, a metabolite of ATP, is more effective in causing smooth muscle hyperpolarization. Therefore, POM-1, an ectonucleotidase inhibitor (Wall et al., 2008) was used to study the role of ectonucleotidases in mediating inhibitory neuromuscular transmission (Figure 2.11). POM-1 decreased the amplitude and the area of IJP. POM-1 was then used to test if also inhibits local ATP response (Figure 2.12). I found that ATP responses were completely blocked by POM-1 verifying that an ATP metabolite mediates hyperpolarizing responses (n=3). Next I studied the effects of POM-1 on exogenous Adenosine 5′-[β-thio]diphosphate (ADP-β-S) application (Figure 2.13). ADP-β-S produced hyperpolarizing responses that were not blocked by POM-1. Finally, MRS 2179 was exogenously applied to verify that these effects were mediated via P2Y1 receptor activation (Figure 2.14). MRS 2179 completely abolished ADP-β-S induced hyperpolarizing responses in a reversible manner.   72     Figure 2.10 ATP and β-NAD induced hyperpolarizing response in the mouse distal colon. Representative hyperpolarizing response to 5 mM, 20 ms (A) ATP and (B) β-NAD puff. (C) ATP/ β-NAD response amplitude – puff duration calibration curve. Response amplitude/area reaches plateau at 20ms puff duration. ATP produced larger hyperpolarization amplitude (20 ± 2 mV) compared to β-NAD (10 ± 3 mV). (D) ATP/β-NAD response area – puff duration calibration curve. ATP produced larger hyperpolarization (67 ± 11 mV) compared to β-NAD (29 ± 10 mV) (n=8 ATP, n=5 β-NAD, *p<0.05)   73     Figure 2.11 Effects of POM-1 on the amplitude and area of IJP in the mouse distal colon. (A) POM-1 application depolarized the membrane potential by 3 ± 3 mV. (B) Time course of POM-1 application on normalized IJP. (C) POM-1 inhibits both the amplitude and the area of the IJP by 76 ± 7% and 66 ± 14% respectively. (D) Representative trace of effect of POM-1 on IJP. Data are mean ± S.E.M (n=6, *p<0.05)   74     Figure 2.12 Effect of POM-1 on ATP response. Figure 2.13 Effect of POM-1 on exogenous Adenosine 5′-[β-thio]diphosphate (ADP-β-S). ADP-β-S (5 mM), a stable ADP analogue produced prominent hyperpolarizing responses. Application of POM-1 did not inhibit ADP-β-S induced hyperpolarizing responses. Figure 2.14 Effect of MRS 2179 on exogenous ADP-β-S. ADP-β-S (5 mM) produced a prominent hyperpolarization, which was reversibly blocked by MRS 2179 (100 µM), a P2Y1 receptor antagonist.   75     2.5 Discussion These results provide evidence that IJPs are mediated by a purine and NO and involve P2Y1 receptors, SK and the ICl(Ca) channels based on following data: 1) blockade of the IJP by MRS-2179 and L-NNA, 2) blockade of the IJP by apamin and NFA, 3) blockade of the IJP and ATP responses by POM 1, and 4) blockade of ADP-β-S responses by MRS 2179. The fIJP is mediated via purinergic signaling mechanisms (Xue et al., 1999). Purinergic signaling is complex because there are several nucleotide candidate mediators including ATP, ADP, UTP, UDP and β-NAD, which activate multiple P2 receptors (Burnstock, 2006a). In this study MRS-2179, a P2Y1 receptor specific antagonist reduced peak IJP amplitude but not duration. This suggests that IJP is at least in part mediated by P2Y1 receptors. NO signaling produces a smaller amplitude but longer lasting slow hyperpolarization (sIJP) (Serio et al., 2003; El-Mahmoudy et al., 2006; Grasa et al., 2009; Mañé et al., 2014). NO is produced in the myenteric neurons via activation of Ca2+ sensitive nNOS. We found that LNNA, a nNOS inhibitor, significantly decreased IJP area but not amplitude. This suggests that NO acts on post junctional smooth muscle target to mediate MRS 2179 insensitive sIJP. Neither MRS 2179 nor L-NNA changed resting membrane potential. Co-application of MRS 2179 and L-NNA completely blocked the IJP without changing resting membrane potential. P2Y1 receptor activation leads to increase in intracellular Ca2+ concentration via inositol triphosphate (IP3) dependent Ca2+ release from the intracellular stores (Hu et al., 2003; Matsuda & Miller, 2010). NO production on the other hand, leads to activation of guanylate cyclase (GC) in the smooth muscle (El-Mahmoudy et al., 2006). GC catalyzes the conversion of GTP to cyclic GMP which activates protein kinase G (PKG) (Carvajal et al., 2000; El-Mahmoudy et al., 2006). The increase in intracellular Ca2+ concentration and PKG production activate different   76     ion channels to mediate IJP either directly via activation of Ca2+ dependent K+ channels or indirectly via PKG dependent phosphorylation of K+ channels (Matsuda & Miller, 2010). PKG also phosphorylates phospholamban (PLB) which causes disinhibition of smooth endoplasmic reticulum Ca2+ ATPase (SERCA) and reduces [Ca2+i]. Subsequent reduction in [Ca2+i] leads to closure of Ca2+-activated Cl- channels (Hartzell et al., 2005). Therefore in order to delineate the ion channels mediating IJPs, apamin was used to block smooth muscle SK channels. Apamin depolarized the membrane potential and reduced IJP area and amplitude. However, there was a significant proportion of IJP that were apamin and paxilline (data not shown) insensitive (Zhang et al., 2000; Zhang et al., 2010). Previous studies showed that closure of CaCCs causes IJPs in opossum esophageal smooth muscle and in guinea pig ileum (Zhang & Paterson, 2002). In the guinea pig ileum, apamin insensitive IJPs had a reversal potential of -25 mV, which was close to Cl- reversal potential (-20-30 mV) suggesting that Cl- ion mediates junction potential in these tissues (Crist et al., 1991). It is hypothesized that subplasmalemmal Cl- efflux from Cl- channels is essential for Ca2+ efflux from the sarcoplamsic reticulum (SR) (Janssen, 2002). NFA dependent inhibition of CaCCs can therefore hyperpolarize the membrane and disrupt SR Ca2+ release into the subplasmalemmal space, which is essential for IJP production (Cruickshank et al., 2003). To test if CaCC contributes to IJPs in the mouse colon CaCCs blocker, NFA was used. I used lower concentration of NFA than used previously in the literature because NFA in concentrations ≥ 100 µM blocks cation channels in the smooth muscle (Zhang & Paterson, 2002). NFA hyperpolarizes the membrane potential and blocks both the amplitude and the area of IJPs. The NFA sensitive IJP was apamin insensitive and co-application of apamin and NFA blocked IJPs similar to MRS 2179 and L-NNA co-application. Taken together these data suggests that IJP is   77     mediated via ATP and NO mediated activation of SK and ICl(Ca) channels. The opposing activity of SK and ICl(Ca) sets the resting membrane potential of smooth muscle in the mouse distal colon. Next, I set to identify the purine molecule involved in mediating fIJP. The literature suggests either ATP or β-NAD as the likely purine candidate involved in mediating fIJP (Xue et al., 1999; Burnstock, 2006b; Hwang et al., 2011). purinoceptors in smooth muscle. ATP and β-NAD both activate P2Y1 ATP and β-NAD were locally applied and their hyperpolarizing responses were measured. ATP had significantly greater response amplitude and area than β-NAD across varying puff durations suggesting greater ATP sensitivity than βNAD. ATP is metabolized in the extracellular space by ecto-nucleotidases (NTPDases 1,2 and 3) where it is broken down to ADP. ADP then act on post junctional P2 receptors (Müller et al., 2006). ARL 67156 and polyoxotungstate 1 (POM-1) are the two major NTPDase effective drugs available (Wall et al., 2008). Among these POM-1 is a more potent inhibitor of NTPDases than ARL67156 because unlike ARL67156, which only inhibits NTPDases1 and 3, POM-1 inhibits NTPDase1, 2 and 3 (Müller et al., 2006). POM-1 is also more efficacious and stable in aqueous solution and biological pH than ARL67156 (Wall et al., 2008). POM-1 was therefore chosen as an inhibitor of NTPDase to test if ATP plays role in regulating inhibitory neuromuscular transmission in the distal colon. POM-1 almost completely inhibits both the IJP and ATP response, suggesting that ATP puff mimics IJP production in the mouse distal colon. Small portion of POM-1 insensitive IJP are possibly NO dependent because they have much smaller amplitude but larger duration. ADP-β-S response on the other hand is insensitive to POM-1 but is blocked completely by MRS-2179 in a reversible manner. Taken together these results suggest that ATP release from the inhibitory motor neurons is the neurotransmitter involved in   78     mediating IJP signaling. In conclusion, these data suggests that ATP acting on P2Y1 receptors and NO affects SK and CaCCs to mediate IJP in the mouse distal colon.   79     BIBLIOGRAPHY   80     BIBLIOGRAPHY Bane V, Lehane M, Dikshit M, O'Riordan A & Furey A. (2014). 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Release of beta-nicotinamide adenine dinucleotide upon stimulation of postganglionic nerve terminals in blood vessels and urinary bladder. J Biol Chem 279, 48893-48903.   82     Wall MJ, Wigmore G, Lopatár J, Frenguelli BG & Dale N. (2008). The novel NTPDase inhibitor sodium polyoxotungstate (POM-1) inhibits ATP breakdown but also blocks central synaptic transmission, an action independent of NTPDase inhibition. Neuropharmacology 55, 1251-1258. Xue L, Farrugia G, Sarr MG & Szurszewski JH. (1999). ATP is a mediator of the fast inhibitory junction potential in human jejunal circular smooth muscle. Am J Physiol 276, G13731379. Zhang Y, Lomax AE & Paterson WG. (2010). P2Y1 receptors mediate apamin-sensitive and insensitive inhibitory junction potentials in murine colonic circular smooth muscle. J Pharmacol Exp Ther 333, 602-611. Zhang Y, Miller DV & Paterson WG. (2000). Opposing roles of K(+) and Cl(-) channels in maintenance of opossum lower esophageal sphincter tone. Am J Physiol Gastrointest Liver Physiol 279, G1226-1234. Zhang Y & Paterson WG. (2002). Role of Ca2+-activated Cl- channels and MLCK in slow IJP in opossum esophageal smooth muscle. Am J Physiol Gastrointest Liver Physiol 283, G104-114.   83     CHAPTER 3 β1BK CHANNEL REGULATES SMOOTH MUSCLE EXCITABILITY IN THE MOUSE DISTAL COLON.   84     3.1 Abstract Smooth muscle specific large conductance Ca2+ activating K+ (β1BK) channels consist of an α and β1 subunit. The α subunit is a pore forming subunit while the β1 subunit modulates channel gating and activation properties. β1BK channel plays an important role in regulating cellular excitability and neurotransmitter release in the neuromuscular junction. Altered β1 subunit expression disrupts coupling between Ca2+ spark and β1BK channel mediated outward K+ currents resulting in increased muscle tone and excitability in urinary bladder, vascular and tracheal smooth muscles. However, its effect on GI smooth muscle is yet unknown. I therefore studied the effects of β1BK channel function on neuromuscular transmission and colonic smooth muscle excitability in-vitro using WT and BK β1-subunit KO mice. I found that while β1subunit KO mice do not show any impairment in inhibitory junction potential (IJPs), the smooth muscles in β1-subunit KO have a depolarized membrane potential and higher action potential firing frequency. This suggests that although β1BK channels are not involved in mediating neuromuscular transmission, they are crucial in regulating smooth muscle excitability in the mouse distal colon.   85     3.2 Introduction Large conductance Ca2+ activating K+ (BK) channels are voltage gated and Ca2+ sensitive channels that lead to net K+ efflux causing membrane hyperpolarization. BK channels are expressed in various cells types including skeletal muscle cell (Tricarico et al., 2005), smooth muscle cell (Brenner et al., 2000b; Chang et al., 2006; Rueda et al., 2013; Shi et al., 2013a; Zheng et al., 2013), neurons (Meera et al., 2000) and endocrine cells (Braun et al., 2008). BK channel consists of an α and β subunit. The α subunit is a pore forming subunit while the βsubunit is an accessory subunit that enhances Ca2+ sensitivity of the pore forming α subunit (Nelson et al., 1995; Brenner et al., 2000b; Cox & Aldrich, 2000; Ohi et al., 2001; Qian & Magleby, 2003; Sweet & Cox, 2009). Based on molecular and electrophysiological studies, 4 different β-subunits (β1-β4) of BK channel have been identified (Brenner et al., 2000a). Among these subunits, the β1 subunit in particular is smooth muscle specific and plays a vital role to mediate smooth muscle tone in vascular (Leo et al., 2014; Pabbidi et al., 2014) and urinary bladder (Petkov et al., 2001) smooth muscle cells. In smooth muscle β1BK channels are located very close to intracellular calcium release sites. When Ca2+ enters through L-type calcium channels, it activates further Ca2+ release through ryanodine receptors (via Ca2+ induced Ca2+ release mechanisms) of the smooth endoplasmic reticulum resulting in high intracellular Ca2+ (>10 µmol L−1) concentration. Increase in intracellular Ca2+ above the threshold activates β1BK channel and initiates spontaneous transient outward current to cause closure of L type Ca2+ channels and smooth muscle relaxation (Nelson et al., 1995; Bolton & Imaizumi, 1996). Although altered expression of β1subunit of the BK channel impairs vascular smooth muscle and urinary bladder smooth muscle relaxation to cause hypertension (Plüger et al., 2000; Yang et al.,   86     2013), heart failure (Wan et al., 2013), and urinary bladder over-activity (Petkov et al., 2001) the functional effect of altered β1BK channel expression in GI smooth muscles is yet unknown. The β1-subunit KO mouse model has been studied in myocytes from urinary bladder (Petkov et al., 2001), trachea (Semenov et al., 2006; Evseev et al., 2013) and colon (Hagen et al., 2003) where they exhibit lower β1BK channel open probabilities and reduced coupling of Ca2+ sparks and outward K+ currents. In addition, the BK channel also controls quantal release of neurotransmitter in the neuromuscular junction (Wang et al., 2001). I therefore hypothesized that reduced coupling of Ca2+ sparks and β1BK channel mediated spontaneous outward K+ currents in β1-subunit KO mice leads to impaired neuromuscular communication and/or altered smooth muscle excitability in the mouse distal colon. In order to test this hypothesis I used intracellular recordings and measured inhibitory junction potentials (IJPs) and action potential firing activity in the GI smooth muscle cells.   87     3.3 Materials and Methods 3.3.1 Ethics Statement All studies were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Publication No. 85-23, revised 1996) and approved by Institutional Animal Care and Use Committee of Michigan State University. 3.3.2 Animals All animal use protocols were approved by the Institutional Animal Care and Use Committee at Michigan State University. Homozygous β1-subunit KO mice were obtained from Dr. Robert Brenner, University of Texas Health Science Center at San Antonio and bred inhouse at Michigan State University. C57BL/6 (WT) mice were either fed standard chow diet, control diet (10 kcal% fat; D12450B) or HF diet (60 kcal% fat; D12492), all from Jackson Laboratories (Bar Harbor, ME, USA). β1-subunit KO mice were weaned at 3 weeks of age. Mice consuming normal diet were studied at 10–12 weeks of age (male, 25–30 g) and mice on control or HF diets were studied at 18-22 weeks (male, 30-52 g). Mice were euthanized using isoflurane anesthesia followed by cervical dislocation. 3.3.3 Electrophysiological recordings Conventional intracellular electrophysiological techniques were used to record IJPs and smooth muscle electrical activity in the distal colon of WT and β1-subunit KO mice. Small segments of distal colon were placed in a petri dish containing Krebs buffer solution. The segment was cut open along the mesenteric border and pinned flat. The mucosa and submucosa were removed using fine forceps. A ~5 mm2 section was cut out and transferred to a silastic elastomer-lined recording chamber. The section was stretched lightly and pinned to the chamber floor using small stainless steel pins. The chamber was placed on the stage of an inverted   88     microscope and the chamber was then perfused with oxygenated (95% O2, 5% CO2) Krebs buffer solution (37 °C) at a flow rate of 3 mL min−1. Drugs were added in known concentrations to the flowing Krebs solution using a system of three-way stopcocks. Drug concentrations reached steady state in the recording chamber within 4 min. Smooth muscle cells were impaled with glass microelectrodes (filled with 2 M KCl, tip resistance = 60–90 MΩ). IJPs were recorder in presence and in absence of nifedipine (1µM). Although the tissue was anchored to the floor of the recording chamber, there was still substantial muscle movement throughout the recording periods when nifedipine was not applied. Despite this complication, it was possible to maintain impalements for periods of 2–60 min. Membrane potential was recorded using an Axoclamp 2A amplifier, a Digidata 1322A analogdigital converter, and Axoscope 9.2 software (all from Molecular Devices, Sunnyvale, CA, USA). Data were only collected and analyzed from individual recordings that lasted ≥ 2 min. 3.3.4 Drugs Drugs and reagents were obtained from Sigma Aldrich Chemical Company (St. Louis, MO, USA). 3.3.5 Statistics Two group comparisons were made using two-way ANOVA with Bonferroni post-hoc test, paired or unpaired Student’s t-test, and Fisher’s exact test. statistically significant. Data are reported as mean ± SEM.   89   p< 0.05 was considered   3.4 Results 3.4.1 β1Bk channel does not mediate inhibitory neuromuscular transmission in the mouse distal colon To assess if β1BK channels play a role in mediating neuromuscular transmission in mouse distal colon, I stimulated colonic tissues at varying voltage (20 -140 V) and measured the corresponding amplitude and the area of the IJPs in presence and in absence of nifedipine. I found that both the amplitude and the area of IJPs were similar between WT and β1-subunit KO either in presence or in absence of nifedipine (p>0.05, two-way Anova with Bonferroni posthoc). Figure 3.1 IJP measurements from WT and β1-subunit KO. Voltage – Amplitude/Area calibration curve in presence (A and B) and in absence (C and D) of nifedipine. Amplitude (A and C) and the area (B and D) of the inhibitory junction potentials (IJPs) are similar between WT and KO.   90     Since IJPs were not disrupted in β1-subunit KO mice, we tested if paxilline, a α subunit blocker of the BK channel plays a role in mediating IJPs. I applied paxilline (0.5 µmol L−1) on colonic smooth muscles preparation and found that paxilline had no effect on either the amplitude or the area of the IJP. However, apamin (0.1µmol L−1), a SK channel blocker significantly decrease both the amplitude and the area of the IJP in both wild type and β1-subunit KO. Figure 3.2 Effects of paxilline on IJPs. Paxilline did not impair the either the (A) Amplitude or the (B) Area of IJP in β1-subunit KO mice. Data are represented as normalized values. Figure 3.3 Effect of co-application of apamin and paxilline on IJPs. Apamin and paxilline coapplication significantly decreased both the IJP amplitude and area by 50 ± 5%, and 62 ± 6% respectively in both wild type and β1-subunit KO. Data are represented as normalized values.   91     3.4.2 Tetrodotoxin (TTX) application does not block smooth muscle action potential in colonic smooth muscles Since IJPs were not impaired in either in β1-subunit KO mice or in paxilline applied tissues, we conclude that β1BK channel do not play a role in mediating IJPs. I tested if β1BK channels mediate smooth muscle excitability and action potential firing. In order to study how β1BK channel affect smooth muscle excitability and action potential firing, intracellular recordings were obtained from circular smooth muscle cells in-vitro in the distal colon of WT and β1-subunit KO mice in absence of nifedipine. I verified that the action potentials were non neurogenic by applying TTX (0. 3 µmol L−1), a Na+ channel blocker. TTX did not block action potential generation in muscles from either WT or β1-subunit KO mice. Figure 3.4 Effect of tetrodotoxin (TTX) on smooth muscle action potential. TTX does not block action potential generation from colonic smooth muscles verifying non-neuronal action potential firing.   92     3.4.3 Increased action potential activity in distal colonic smooth muscle from β1-subunit KO mice I used intracellular recording to measure smooth muscle action potentials in WT and β1subunit KO mice. Recordings were obtained from 42 cells in distal colonic tissues from 11 WT mice. I found that action potential firing occurred in short bursts with a 10.4 ± 0.6 s interburst interval in 30 cells whereas action potentials occurred continuously in the remaining 12 cells in WT (Figure 3.5A). Recordings were obtained from 29 cells in distal colonic tissues from seven β1-subunit KO mice. In KOs, action potential firing occurred continuously in 23 cells whereas bursting occurred in the remaining six cells (Figure 3.5B). The proportion of bursting cells was significantly higher in WT compared to β1-subunit KO mice (p<0.05, Fisher’s exact test). The average action potential firing frequency was also greater in tissues from β1-subunit KO compared to WT mice (Figure 3.5C). The resting membrane potential in circular smooth muscle cells from the distal colon of β1-subunit KO mice was significantly more depolarized compared to that in WT preparations (Figure 3.5D).   93     Figure 3.5 Action potential firing pattern in WT and β1-subunit KO. (A) Representative recording of action potential firing pattern in the distal colon of a WT mice. Action potentials occur in short bursts with a ~10 s inter-burst interval. (B) Recording of continuous action potential firing in the distal colon of a β1-subunit KO mouse. (C) Mean action potential firing rate is greater in distal colon of β1-subunit KO (n=31 cells from 7 mice) compared to WT (n=42 cells from 11 mice) mice (*p<0.05). (D) Resting membrane potential in distal colon circular smooth muscle cells were more depolarized from β1-subunit KO compared to WT mice in absence of nifedipine (*p<0.05). Reprinted from Neurogastroenterology & Motility, volume 24, France et al., Impaired propulsive motility in the distal but not proximal colon of BK channel β1subunit knockout mice, p e450-9, copyright (2012).   94     3.4.4 Effects of nifedipine application in action potential generation and smooth muscle membrane potential in WT and β1-subunit KO The L-type Ca2+ channel antagonist nifedipine (1 µmol L−1) completely blocked action potential firing in distal colon circular smooth muscle cells from WT and β1-subunit KO mice (Figure 3.6). Nifedipine depolarized the membrane potential in distal colon circular smooth muscle cells from WT mice (Figure 3.6A, mean depolarization after 5 min nifedipine application =4.4 ± 2.2 mV). Conversely, nifedipine did not change the membrane potential of distal colon smooth muscle cells from β1-subunit KO mice (Figure 3.6B, mean depolarization after 5 min nifedipine application = 2.6 ± 2.3 mV). No difference in resting membrane potential was seen after nifedipine application.   95     Figure 3.6 Nifedipine blocks action potential firing in WT and BK β1-subunit KOs. Nifedipine caused a small depolarization in tissues from WT but not BK β1-subunit KO mice. (C) RMP was similar in both WT and β1-subunit KO in presence of nifedipine (1 µM). Reprinted from Neurogastroenterology & Motility, volume 24, France et al., Impaired propulsive motility in the distal but not proximal colon of BK channel β1-subunit knockout mice, p e450-9, copyright (2012).   96     3.4.5 Effects of Paxilline on membrane potential and action potential firing in distal colonic smooth muscle of WT mice Since paxilline is a BK channel blocker, it would be expected that paxilline treatment should mimic the effects of β1-subunit KO. Paxilline treatment for a minimum of 4 min caused a membrane potential depolarization Figure 3.7, top trace). Paxilline did not change the bursting action potential firing pattern, but it did cause an increase in the number of action potentials during each burst (Figure 3.7, bottom traces) suggesting that blocking the BK channel has different effects on smooth muscle activity compared with deletion of a molecular component of the channel. Figure 3.7 Effect of paxilline on smooth muscle action potential firing. Upper trace shows a slow time base recording of action potential bursting in WT distal colonic smooth muscles. Paxilline causes a membrane depolarization but action potential bursting persists. Lower traces show expanded views of action potential bursting in the absence (left) and presence (right) of paxilline. Paxilline caused a prolongation of the bursts of action potentials. Reprinted from Neurogastroenterology & Motility, volume 24, France et al., Impaired propulsive motility in the distal but not proximal colon of BK channel β1-subunit knockout mice, p e450-9, copyright (2012).   97     Figure 3.8 Effect of paxilline on smooth muscle membrane potential. Paxilline caused a significant depolarization in distal colon smooth muscle cells from WT mice (recordings from 9 cells from 3 mice) (*p<0.05). Paxilline also caused an increase in the number of action potentials per burst (*p<0.05). Reprinted from Neurogastroenterology & Motility, volume 24, France et al., Impaired propulsive motility in the distal but not proximal colon of BK channel β1-subunit knockout mice, p e450-9, copyright (2012).   98     3.5 Discussion My in vitro data indicate that although β1BK channel function is not important to maintain inhibitory neuromuscular communication, it is crucial to maintain the resting membrane potential and rhythmic action potential firing in the mouse distal colonic smooth muscles. I tested if β1Bk channel plays a role in mediating inhibitory neuromuscular communication and smooth muscle excitability in the distal colon by measuring IJPs and action potential activity in WT and β1-subunit KO. Mouse distal colon was the tissue of interest because unlike small intestine and proximal colon whose function is to absorb nutrients and water from the gut lumen, distal colon is primarily a storage organ, hence, the greater need for smooth muscle relaxation (which would be facilitated by BK channel activity). Nifedipine was applied to prevent spontaneous smooth muscle contraction and obtain stable impalements. I measured IJPs from WT and β1-subunit KOs but could detect any difference in either the amplitude or the area of IJPs. Since nifedipine blocks L-type voltage gated calcium channels and prevents Ca2+ entry to the cells, it is possible that application of nifedipine drops intracellular calcium level below the threshold needed for β1BK channel function (1-10µM). In order to address this issue, the amplitude and the area of IJPs were measured in absence of nifedipine. However, no significant difference between WT and β1subunit KOs were found. This suggests inhibitory neuromuscular transmission is not impaired in β1-subunit KOs. Next I applied paxilline in order to study if the α subunit of BK channel plays a role in mediating inhibitory neuromuscular transmission. Paxilline did not disrupt IJPs in both WT and β1-subunit KOs, however, apamin a SK channel blocker significantly disrupted IJPs in both tissues. This suggests that unlike SK channels, β1BK channel does not participate in mediating inhibitory neuromuscular communication in the mouse distal colon.   99     Next I tested if the β1BK channel regulates smooth muscle excitability. Since β1BK channels are activated by Ca2+ induced Ca2+ release from intracellular stores and the β1-subunit modulates channel function by increasing Ca2+ sensitivity (Ohi et al., 2001; Qian & Magleby, 2003; Sweet & Cox, 2009), absence of the β1-subunit would reduce Ca2+ sensitivity and therefore reduce channel opening. Reductions in outward K+ currents would increase smooth muscle excitability (Petkov et al., 2001; Semenov et al., 2011). This theory is supported by my electrophysiological data where I found that action potential firing is increased in β1-subunit KO mice as seen by continuous action potential firing in distal colon smooth muscle cells from these mice. Conversely, action potentials occurred in short bursts with an interburst interval of ~10 s in WT mice. Action potentials in all smooth muscle cells were Na+ independent and Ca2+-dependent because the L-type Ca2+ channel antagonist, nifedipine, blocked action potentials while TTX could not. Bursting behavior in WT cells would result from Ca2+ entry during action potential generation subsequently activating BK channels and bringing the membrane potential below action potential threshold. Firing would cease until intracellular Ca2+ levels fell below those needed to activate BK channels allowing the membrane to depolarize again leading to another burst of action potentials. In β1-subunit KO cells, the BK channel would be less sensitive to Ca2+ and would not open during intracellular Ca2+ increases that would occur during action potential bursts. Post-burst hyperpolarization would be absent and therefore smooth muscle cells could fire action potentials continuously. Continuous firing would also be driven by the depolarized membrane potential as measured in β1-subunit KO smooth muscle cells. The BK channel blocker, paxilline partially mimicked the effects of β1-subunit KO. Paxilline caused a depolarization in smooth muscle membrane potential supporting the role of the BK channel in   100     regulating membrane potential of the smooth muscle cells. However, paxilline did not change the action potential firing pattern from bursting to continuous in WT tissues. In the presence of paxilline, bursting behavior was sustained but the duration of each burst was prolonged. These data suggest that blocking the BK channel has different effects on smooth muscle activity compared with deletion of a molecular component of the channel. Alternatively, there may be other molecular changes in the mechanisms controlling smooth muscle excitability in response to β1-subunit KO that are not mimicked by acute pharmacological blockade of the BK channel. Nifedipine caused a depolarization in distal colon smooth muscle cells from WT, but not β1-subunit KO mice. This is consistent with Ca2+ entry through L-type Ca2+ channels driving BK channel activation and hyperpolarization of the membrane potential. In the absence of the β1-subunit, the α-subunit is less sensitive to Ca2+ activation and therefore the membrane potential of smooth muscle cells is less polarized. As stated above, nifedipine was able to block action potentials in both WT and β1-subunit KO cells. In β1-subunit KO cells, action potentials occur continuously until the addition of nifedipine. Nifedipine acts to reduce gradually the action potential firing frequency as nifedipine concentrations slowly reached a steady state level. Firing frequency decreased but I did not detect bursting in β1-subunit KO cells as action potentials fired continuously, but at a progressively lower frequency. In conclusion these data suggests that while β1BK channel do not play a major role in mediating inhibitory neuromuscular transmission, it is imperative to maintain smooth muscle excitability in the mouse distal colon. Genetic KO of β1-subunit could therefore lead to disrupted colonic propulsive motility patterns (France et al., 2012).   101     BIBLIOGRAPHY   102     BIBLIOGRAPHY Bolton TB & Imaizumi Y. (1996). Spontaneous transient outward currents in smooth muscle cells. Cell Calcium 20, 141-152. Braun M, Ramracheya R, Bengtsson M, Zhang Q, Karanauskaite J, Partridge C, Johnson PR & Rorsman P. 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BK channel β1 subunits regulate airway contraction secondary to M2 muscarinic acetylcholine receptor mediated depolarization. J Physiol 589, 1803-1817. Shi L, Liu B, Li N, Xue Z & Liu X. (2013). Aerobic exercise increases BK(Ca) channel contribution to regulation of mesenteric arterial tone by upregulating β1-subunit. Exp Physiol 98, 326-336. Sweet TB & Cox DH. (2009). Measuring the influence of the BKCa {beta}1 subunit on Ca2+ binding to the BKCa channel. J Gen Physiol 133, 139-150. Tricarico D, Mele A & Conte Camerino D. (2005). Phenotype-dependent functional and pharmacological properties of BK channels in skeletal muscle: effects of microgravity. Neurobiol Dis 20, 296-302. Wan E, Kushner JS, Zakharov S, Nui XW, Chudasama N, Kelly C, Waase M, Doshi D, Liu G, Iwata S, Shiomi T, Katchman A, D'Armiento J, Homma S & Marx SO. (2013). Reduced vascular smooth muscle BK channel current underlies heart failure-induced vasoconstriction in mice. FASEB J 27, 1859-1867. Wang ZW, Saifee O, Nonet ML & Salkoff L. (2001). SLO-1 potassium channels control quantal content of neurotransmitter release at the C. elegans neuromuscular junction. Neuron 32, 867-881. Yang Y, Li PY, Cheng J, Mao L, Wen J, Tan XQ, Liu ZF & Zeng XR. (2013). Function of BKCa channels is reduced in human vascular smooth muscle cells from Han Chinese patients with hypertension. Hypertension 61, 519-525. Zheng YM, Park SW, Stokes L, Tang Q, Xiao JH & Wang YX. (2013). Distinct activity of BK channel β1-subunit in cerebral and pulmonary artery smooth muscle cells. Am J Physiol Cell Physiol 304, C780-789.   105     CHAPTER 4 HIGH FAT DIET DISRUPTS INHIBITORY NEUROMUSCULAR TRANSMISSION AND SMOOTH MUSCLE EXCITABILITY IN THE MOUSE DISTAL COLON   106     4.1 Abstract High fat (HF) diet disrupts gastrointestinal (GI) motility by unclear mechanisms. We studied the effect of HF diet on nitrergic neurons, neuromuscular transmission, smooth muscle cell (SMC) excitability and colonic motility in wild type (WT; C57BL/6J) and β1-subunit knockout (KO) mice. HF diet caused increased myenteric plexus oxidative stress, a 28% decrease in nitrergic myenteric neurons and a 20% decrease in basal NO levels in the myenteric plexus. Circular muscle inhibitory junction potentials (IJPs) were smaller in the colon of WT HF fed mice. NLA (NOS inhibitor) was less effective at inhibiting relaxations in HF fed compared to control fed WT mice (11% vs 37%). SMCs in HF fed WT mice exhibited depolarized membrane potential (-47 ± 2 mV, n=6) and continuous action potential firing compared to control fed WT mice (-53 ± 2 mV n=6, p<0.05), which showed rhythmic firing. SMCs from control and HF fed β1BK KO mice exhibited continuous action potential firing. NLA depolarized colonic SMC membrane potential and caused continuous firing in SMCs from control fed WT mice. Sodium nitroprusside (NO donor) hyperpolarized membrane potential and changed continuous to rhythmic action potential firing in colonic tissues from WT HF fed and β1BK KO mice. Colonic migrating motor complexes were disrupted in β1BK KO mice and HF fed WT mice. Diet did not change BK α-subunit protein or mRNA expression or β1BK-subunit mRNA expression. We conclude that HF diet disrupts IJPs, SMC excitability and GI motility by promoting oxidative stress, loss of nitrergic neurons and disruption of SMC β1BK channel function.   107     4.2 Introduction Obesity in humans is mostly caused by the consumption of high fat (HF) diet (Y Lee, 2013) and leads to Type 2 diabetes (T2D) (Colditz et al., 1995), heart disease (Huang et al., 1998), arthritis (Heliövaara et al., 1993) and cancer (Kopelman, 2000; Farag & Gaballa, 2011). Obesity and HF diet are also associated with increased risk of gastrointestinal (GI) motility disorders (Delgado-Aros et al., 2004; Xing & Chen, 2004; Mushref & Srinivasan, 2013; Y Lee, 2013). GI motility is controlled largely by interactions between enteric neurons, interstitial cells of Cajal (ICC), and smooth muscle cells (SMCs) but how obesity-induced changes in these cell types contribute to GI dysmotility is poorly understood. Animal models of obesity show an increase in oxidative stress followed by altered function of some myenteric neurons (Chandrasekharan et al., 2011a; Voukali et al., 2011; Voss et al., 2013). The neuropathological changes that cause GI dysmotility vary among animal models and location in the GI tract (Chandrasekharan & Srinivasan, 2007). Inhibitory nitric oxide (NO) motor neurons are most susceptible to damage caused by obesity (Chandrasekharan et al., 2011a; Beraldi et al., 2014). Since activation of NO motor neurons cause smooth muscle (SM) relaxation (Murr et al., 1999; Grasa et al., 2009), it is crucial to determine if myenteric NO neurons are lost in obesity and how this contributes to disrupted GI motility. GI SMC function also plays a crucial role producing propulsive motility. GI SMCs have rhythmic electrical activity (slow waves), initiated by ICCs and slow waves control contraction frequency. The large conductance Ca2+ activated K+ channel (BK channel) helps to regulate membrane potential and excitability of GI SMCs (Vergara et al., 1998; France et al., 2012). The SMC BK channel is composed of an α subunit and the SM specific β1 subunit. The BK β1 subunit enhances Ca2+ sensitivity of the pore forming α subunit (Cox & Aldrich, 2000; Qian &   108     Magleby, 2003). Down-regulation of BK β1-subunit expression reduces the calcium sensitivity of the α subunit and causes increased SM contractility (Brenner et al., 2000b; Plüger et al., 2000). This leads to various disorders that include hypertension (Yang et al., 2013), heart failure (Wan et al., 2013) and urinary bladder over activity (Petkov et al., 2001). Although obesity (Howitt et al., 2011), oxidative stress (Santarelli et al., 2004; Tang et al., 2004) and NO availability (Leo et al., 2014) all modulate the function/expression of β1 subunit in vascular SMCs, it is unclear is similar modulation occurs in the gut. The etiology of obesity induced GI motility disorders is multifactorial and includes alterations to multiple cell types. My aim in the present study was to address how inhibitory neuromuscular transmission is affected by obesity. I hypothesized that in obesity, oxidative stress causes loss of inhibitory nitrergic motor neurons and disrupts β1BK channel function/expression leading to impaired inhibitory neuromuscular transmission and increased SMC excitability. I tested my hypothesis using colonic tissues from wild type (WT) and β1BK KO mouse models placed either on control or HF diet. My findings show that HF diet causes oxidative stress induced loss of NO motor neurons, which disrupts inhibitory neuromuscular transmission and SM excitability causing impaired colonic motility.   109     4.3 Materials and methods 4.3.1 Ethics Statement All studies were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Publication No. 85-23, revised 1996) and approved by Institutional Animal Care and Use Committee of Michigan State University. 4.3.2 Animals Wild type (WT) 18-24 weeks old male C57BL/6J mice maintained either on control (10 kcal% fat; D12450B) or HF diet (60 kcal% fat; D12492), were obtained from Jackson laboratories (Bar Harbor, Maine, USA) (http://jaxmice.jax.org/diomice/diets.html), while β1BK KO mice were obtained as a gift from Dr. Robert Brenner (University of Texas Health Science Center, San Antonio). β1BK KO mice were maintained as homozygous lines at Michigan State University (Xu et al., 2011; France et al., 2012; Xu et al., 2012; Xu et al., 2014). Pups of β1BK KO mice were weaned at 3 weeks. After 3 weeks, mice were placed either on control or a HF diet. Animals were kept in 12/12 hr light/dark cycle, and had access to food and water ad libitum. WT and β1BK KO mice were age matched for the study. WT mice were maintained in our animal facility for 1-week prior to use in any experiments. Euthanasia was carried out by isoflurane anesthesia followed by cervical dislocation. 4.3.3 Oxidative stress measurements Whole colon segments from control or HF WT mice were cut open along the mesenteric border, washed, and stretched using fine pins (0.1mm) (Fine Science Tools, Foster City, CA) in a silastic elastomer-lined (VWR, Buffalo Grove, IL) petri dish. Tissues were dissected to expose the longitudinal muscle myenteric plexus (LMMP). Reduced thiols were labeled with Alexa fluor 680-conjugated C2 maleimide (1 µM; Life Technologies, Grand Island, NY) in 4% paraformaldehyde (PFA), 0.02% Triton X-100 and 1 mM N-ethylmaleimide (NEM, E3876;   110     Sigma-Aldrich, St. Louis, MO) in 1 M PBS for 20 min at room temperature. Tissues were rinsed with PBS. In order to visualize oxidized thiols, LMMP preparations were then treated with 5 mM tris (2-carboxyethyl) phosphine hydrochloride (TCEP) (Catalog #C4706; Sigma-Aldrich, St. Louis, MO) for 20 min. TCEP chemically reacts with oxidized thiol to form reduced free thiols. The newly formed reduced thiols were labeled with 1 µM Alexa fluor 546 C5 maleimide in 1 mM NEM in PBS for another 20 min at room temperature. Finally, the ratio of 546-maleimide (oxidized thiol) /680-maleimide (reduced thiol) (SS/SH ratio) fluorescence intensity was calculated using Image J (Version 1.44 g, National Institute of Health, USA) software. Higher ratios indicate increased oxidative stress. 4.3.4 Immunohistochemistry for nitric oxide synthase (NOS) containing neurons Mucosa, submucosa and the circular muscle were dissected away to expose myenteric ganglia from control and HF mice. Whole-mount LMMP preparations were fixed in 4% PFA for 1 hour at room temperature. The tissues were then washed (3 x 10 min) with 0.1M PBS prior to incubation in 0.5% Triton X-100 and 4% normal donkey serum. Tissues were incubated overnight in sheep anti nNOS primaries (Cat #AB1529, Millipore, USA; 1:200 dilution). After washing the tissues (3 x 15 min) in PBS the following day, the antibody staining was visualized using fluorescein isothiocyanate (FITC)-conjugated donkey anti sheep antibody (Cat # 713-486147, Jackson Lab, West Grove, PA; 1:100 dilution). Fluorescent images were obtained through 40X (PlanFluor, 0.75 numerical aperture) objective of an upright epiflourescence microscope (Nikon Eclipse, Melville, NY) equipped with a Retiga 2000R camera (QImaging, Surrey, BC, Canada) controlled by QCapture Pro 7.0 (QImaging) software. Primary and secondary only controls were run and showed no labeling. NO positive neurons number and ganglionic area   111     were obtained from five random locations on the tissues. Images were analyzed using Image J (Version 1.44g, National Institute of Health, USA) software. 4.3.5 Nitric oxide (NO) measurements Freshly dissected LMMP preparations from either control or HF mice were treated with collagenase type II (156µg/ml) (Cat #17101-015; Gibco, Grand Island, NY) and dispase (1µg/ml) (Cat #17105-041; Gibco, Grand Island, NY) dissolved in DMEM/F-12 media (Life technologies, USA) for 15min. The tissues were then loaded with diaminofluorophore 4-amino5-methylamino-2′-7′-difluorofluorescein diacetate (DAF-FM-DA; 4µM; Catalog # D-2321, Life Technologies, Grand Island, NY; 1:1000) dissolved in DMEM/F-12 supplemented with 0.08% probenecid for 30 min. Tissues were washed (3 x 1 min) with DMEM/F12 and allowed to deesterify for 20 min in presence of 200 µM probenecid. DAF-FM fluorescence in myenteric ganglia was captured through the 40X water-immersion objective of an upright Olympus BX51WI fixed-stage microscope (Olympus, Center Valley, PA) equipped with a Neo sCMOS digital camera (Andor, South Windsor, CT) controlled by Andor IQ3 software. Images were analyzed offline using Image J software. 4.3.6 Electrophysiological recordings from colonic circular muscle SMCs Distal colon segments were removed from euthanized mice and placed in carboxygenated (95% O2 and 5% CO2 mixture) Krebs solution containing (mM) 11 glucose, 117 NaCl, 25 NaHCO3, 4.7 KCl, 1.2 NaH2PO4.H2O, 2.5 CaCl2.2H2O, and 1.2 MgCl2 6H2O, (pH 7.3–7.4). The mesenteric fat was removed and the segment was opened along the mesenteric border. The tissue segment was stretched and pinned flat on a silastic elastomer lined petri dish with stainless steel pins. Mucosa and submucosa were removed using fine forceps. A small tissue section (~3 x 1 cm) with intact myenteric plexus and circular muscle was cut out and transferred to a silastic   112     elastomer lined recording chamber. The chamber was then placed on the stage of an inverted microscope (Nikon Eclipse TS100). Warm carboxygenated Krebs solution (37°C) was continuously perfused through the chamber at 3-4 ml/min. Circular SMCs were impaled with a glass microelectrode filled with 2M KCl (50 - 80MΩ tip resistance). Membrane potential was recorded using an Axoclamp 2A amplifier, a Digidata 1332A analog-digital converter and axoscope 9.2 software (all from Molecular Devices, Sunnyvale, CA). Resting membrane potential of the SMC and the spontaneous action potential activity was recorded in absence of nifedipine. Individual recordings that lasted ≥ 2 min were later used for data analysis. Inhibitory junction potentials (IJPs) were evoked using electrical field stimulation (train duration, 200-1000 ms; frequency, 10 Hz; pulse duration 0.8 ms; voltages 60-120V) was applied using two parallel silver chloride wires placed 3.5 cm apart (Ø=0.03", A-M systems, Carlsberg, WA). The wires were placed perpendicular to the longitudinal axis of the preparation. The Ltype Ca2+ channel blocker, nifedipine (1 µM) was added to abolish spontaneous muscle contractions. 4.3.7 Isometric tension measurement of muscle relaxation in-vitro Experiments were designed to assess the contribution of β1BK channels in mediating SM relaxation. One-centimeter segments of distal colon from WT and β1BK KO mice fed a control and HF diets were used. Tissues were attached horizontally (in the axis of the circular muscle) to the tissue holder and connected to a force-displacement transducer (FT03C; Grass Instruments, Quincy, MA) using silk thread. Preparations were suspended horizontally and placed under 1 g of passive tension in 20 ml of glass organ baths containing warm carboxygenated Krebs solution (37°C). Tissues were then allowed to equilibrate for 40 min prior to the start of experiments. During this time Krebs solution was changed every 10 min. Electrical field stimulation (EFS; 20   113     mA, 10 – 40 Hz) was applied using concentric platinum wire electrodes to elicit neurogenic relaxation in the presence of PGF2α (5 µM, to increase muscle tone) and scopolamine (1 µM to block muscarinic receptors). The amplitude and area of the relaxation were measured. Data were analyzed using Labscribe software (Iworx, Dover, NH). The Na+ channel blocker tetrodotoxin (TTX, 1 µM) was used to confirm that stimulation evoked relaxations were neurogenic. 4.3.8 In-vitro assessment of colonic motility Whole colons were harvested from WT and β1BK KO mice, and luminal contents were flushed with Krebs buffer. A stainless-steel rod was then inserted into the lumen. The tissue was secured at each end (~1.5 cm apart) with small needles that were attached to surgical silk thread. The silk threads were then attached to two force transducers. The rod holding the tissue was placed in a 60 ml reservoir filled with warm carboxygenated krebs solution and stretched to an initial tension of 0.5 g. The tissues were then allowed to equilibrate for 30 min. After regular colonic migrating contractions (CMMC) were established, CMMC activity was recorded from control fed and HF tissue for an hour. Recordings were obtained using two strain gauge amplifiers (Grass Instruments CP122A) attached to an analog/digital converter (Minidigi 1A, Molecular Devices) and finally analyzed with axoscope 9.2 software (Molecular Devices). Retrograde (anal to oral), anterograde (oral to anal) and simultaneous (oral and anal occurring at the same time) contractions were identified and recorded. Velocity of anterograde CMMC propagation was obtained by dividing the distance between the clips (1.5 cm) with the time delay for oral contraction to reach the anal end. 4.3.9 In-vivo assessment of colonic motility WT and β1BK KO mice placed on control and HF diet were individually housed in clean cages where food and water were removed for 1 h between 11 AM and 12 PM. Fecal pellets   114     collected from each mouse during this time and were counted and the wet weight measured. Fecal pellets were later stored in a warm (40°C) oven for 24 hours and the dry weight was measured the next day. This protocol was repeated for four consecutive days. Fecal pellet number, wet weight and dry weight were all normalized to the body weight of the animal over the four-day period. 4.3.10 Western blot for α subunit protein expression Colon samples from WT control and HF mice were collected. Mucosa and submucosal layers were quickly removed in cold Krebs solution containing protease inhibitor cocktail (P8340; Sigma-Aldrich, St. Louis, MO) 10 mmol/L sodium fluoride (S7920; Sigma‐Aldrich), and 1 mmol/L sodium orthovanadate (S6508; Sigma-Aldrich) to prevent protein degradation. Tissues were stored at -80° C until ready to be processed and later homogenized with a mortar and pestle that had been pre-cooled with liquid nitrogen. Homogenized tissues were lysed with 1X lysis buffer (12.5 ml 0.5M Tris HCl, 20 ml 10% SDS, 10 ml Glycerol, 57.5 ml dH2O) that contained protease inhibitor cocktail. Protein (5 mg) was loaded for SDS-PAGE along with the full-range rainbow molecular weight marker (GE Healthcare, RPN800E). Proteins were separated at 100 V on a 10-12% gel, and electro-transferred overnight onto a 0.45 mm PVDF at 30 V (Milipore, IPVH00010). Ponceau S was used to verify efficient protein transfer. The following day membranes were blocked for 1 hour at room temperature in 5% milk-TBS-T to prevent non-specific binding. Membranes were blocked for 2hrs, with BK α-subunit primary antibodies (Alomone labs, APC-107; 1:20,000). Membranes were then washed (3 X 10 minutes) with TBS-T, and blotted with the appropriate HRP-conjugated secondary antibodies (1:4000) for 1 hour at room temperature in blocking solution. Blots were imaged with ECL Plus Western Blotting Detection System (GE Healthcare).   115     4.3.11 Real time rt-PCR for α and β1 subunits mRNA expression I used rt-PCR to detect if there were HF diet-induced changes in α and β1 mRNA expression in WT mice. Mucosa and submucosa layers were dissected and colonic SMs were collected. SMs were stored in All Protect Reagent (Qiagen, 76405) until ready to be processed. RNA was isolated using the MELT Total Nucleic Acid Isolation System kit (Ambion, no. AM1983). RNA isolates were reverse-transcribed to cDNA using High Capacity RNA to cDNA kits (Applied Biosystems, 4387406). Resultant cDNA was used for real-time PCR assays. Applied Biosystems TaqMan Fast Advanced master mix (Applied Biosystems, 4444556) was used to prepare reactions (20 µl). Reactions were assayed with TaqMan Gene Expression assays (KCNMA1-no Mm00516078_m1, Gapdh – no. Rn01775763_g1). Samples were analyzed in duplicate. 4.3.12 Statistics Potential differences between treatment groups were tested using either two-way ANOVA with Bonferroni post-hoc test or Student’s t-test. Data are mean ± SEM. Differences were considered significant when p< 0.05. Analysis was done in Graph Pad Prism 4.0 software (San Diego, CA).   116     4.4 Results 4.4.1 HF diet increases body weight and abdominal fat pad weight I tested if mice develop obesity after consuming a HF diet for 18-22 weeks and found that HF fed mice had a significant increase in body weight (41.7 ± 1.4 g, n=21) compared to control fed mice (28.3 ± 0.47 g, n=21; p<0.05). HF fed mice also had a significant increase in abdominal fat pad weight compared to control fed groups (0.4 ± 0.04g, n=9 Vs 1.7 ± 0.2g, n=9; p<0.05). 4.4.2 WT HF mice exhibit increased neuronal oxidative stress, decreased NOS neuron density and lower NO availability. I measured neuronal thiol oxidation to determine if consumption of a HF diet drives oxidative stress in myenteric NO neurons. I observed an increase in the GSSG/GSH fluorescence ratio in NOS and non NOS neurons of HF fed mice compared to control fed mice (Figure 4.1).   117     Figure 4.1 Glutathione fluorescence in the myenteric plexus of the mouse distal colon. Representative images of (A) oxidized and (B) reduced glutathione fluorescence intensity. (C) nNOS positive neurons were co-localized with glutathione fluorescence measurements to measure oxidative stress (increased oxidized glutathione) in these neurons. (D) Fluorescence intensity measurements show 17 ± 2% increase in oxidized/reduced glutathione ratio in myenteric nitric oxide neurons of HF (n=6) compared to control fed mice (n=6). (E) Increase in oxidized/ reduced GSH ratio was also seen in non-nNOS positive neurons from HF fed (n=13) compared to control fed mice (n=14). *p<0.05. I tested to see if an increase in oxidative stress is accompanied by loss of NOS neurons in HF fed mice. nNOS positive neurons were detected in the myenteric plexus of control fed and HF mice but nNOS neuron density was significantly lower in the myenteric plexus of HF fed compared to control fed mice (Figure 4.2C). Given the lower frequency of neurons containing nNOS in mice fed a HF diet, I reasoned that myenteric ganglia in these animals would contain lower levels of NO availability. Therefore, I assessed NO levels within myenteric ganglia using the fluorescent NO indicator dye, DAF-FM-DA. DAF-FM-DA reliably detects NO levels in vertebrates (Hong et al., 2009) and does not fluoresce in the absence of NO. In the presence of   118     NO, DAF-FM-DA is converted into fluorescent benzotriazole molecules whose intensity is directly correlated to bioavailability of NO in the tissue (Hong et al., 2009). I measured the fluorescence intensity of benzotriazole derivative as an indirect measure of basal NO availability. The fluorescence intensity of the benzotriazole derivative was greater in control mice suggesting lower NO availability in the myenteric ganglia of HF mice (Figure 4.2D) Figure 4.2 Immunohistochemical labeling for neuronal nitric oxide synthase (nNOS) containing neurons from control and high fat (HF) mice. (C) Number of nNOS positive neurons per ganglionic area was higher in control fed (n=6) compared to HF fed mice (n=6). (D) Fluorescence intensity of DAF-FM-DA derived benzotriazole derivative is greater in HF fed (n=6) compared to control fed mice (n=6). *p<0.05. 4.4.3 HF diet impairs inhibitory nitrergic neuromuscular transmission Inhibitory neuromuscular transmission in the gut involves purinergic and nitrergic signaling (Murr et al., 1999; Gallego et al., 2008; Grasa et al., 2009). A purine mediates fast inhibitory junction potentials (IJPs) and fast relaxation while NO mediates slowly developing but   119     longer lasting IJPs and relaxations. I used intracellular microelectrode recording to measure IJPs at varying train duration (200-1000 ms) and measured amplitude (indicative of fast purinergic IJP) and the area (indicative of nitrergic slow IJP) in control and HF fed mice. At higher train duration (800 ms), the area of IJP but not the amplitude was significantly smaller (52% 100 V, p<0.05) in HF mice compared to controls (Figure 4.3). Figure 4.3 IJP recordings from control and HF fed mice. (A) Representative recording of IJPs in mouse distal colon (EFS: 100 V, 10 Hz and 0.2-1 s train duration) from control and HF tissues. (B) IJP area was reduced only at the 0.8 s train duration in tissues from HF fed mice. (C) IJP amplitude was similar in tissues from control and HF fed mice. *p<0.05. 4.4.4 HF diet impairs neurogenic smooth muscle relaxation To test if disrupted inhibitory neuromuscular transmission can lead to impairment in neurogenic SM relaxation, I measured the force of relaxation generated by the colonic circular   120     smooth muscle (Figure 4.4A). We found that neurogenic relaxations in tissues from WT HF fed were smaller than in tissues from control fed mice (p<0.05, two-way ANOVA), however no such differences were observed in β1BK KOs (Figure 4.4B). NLA (100 µM) also caused a smaller inhibition of neurogenic relaxation in tissues from HF fed mice (11 ± 3%) compared to control fed mice (37 ± 6%, p<0.05) (Figure 4.4C). Figure 4.4 Measurement of neurogenic smooth muscle relaxation in the mouse distal colon. (A) Neurogenic smooth muscle relaxation in control and HF groups in presence of prostaglandin (5 µM) and scopolamine (10 µM). (B) HF mice showed significantly impaired neurogenic smooth muscle relaxation across varying frequencies compared to control groups. (C) Application of NLA (100 µM; Nitric oxide synthase inhibitor) significantly decreased neurogenic relaxation in both groups. However, the percentage inhibition of relaxation in presence of NLA was significantly greater in controls than in HF. *p<0.05. All tissues were precontracted with PGF2a (5 mM) and muscarinic receptors were blocked with scopolamine (1 mM).   121     4.4.5 HF diet mimics the effect of β1BK KO on SM electrical activity I tested the hypothesis that HF diet dependent increase in oxidative stress alters smooth muscle electrical activity by impairing β1BK channel function. Intracellular recording were obtained from colonic circular SMCs from control and HF fed mice. Colon SMCs from HF fed mice had a depolarized membrane potential (-47 ± 1.9 mV n=16) compared to control fed mice (-53 ± 1.7 mV n=13; p<0.05). SMCs in WT control fed mice showed rhythmic action potential firing, while SMCs in tissues from HF fed mice showed continuous action potential firing activity. 13 out of 17 cells from 7 WT HF mice exhibited continuous action potential firing behavior while 11 out of 13 cells impaled in 6 WT controls exhibited bursting behavior. (p<0.05, Fischer exact test, n=6) (Figure 4.5). SMCs from β1BK KO mice showed continuous action potential firing independent of diet. Only 4 out of 11 cells recorded from 6 control fed β1BK KO mice showed bursting action potential firing while remaining 7 cells showed continuous firing. Only 3 out of 18 cells from 7 β1BK KO HF mice showed bursting action potential firing while remaining 15 cells showed continuous firing (p<0.05, Fischer exact test). 4.4.6 NO availability affects SM excitability in WT and β1BK KO mice NO modulates BK channel activity in vascular SMCs (Mistry & Garland, 1998; L'Heureux et al., 2010). To test if NO availability alters GI SM excitability, I applied NLA (100 µM) and SNP (10 mM) to WT and β1BK KO tissues. NLA depolarized (-50 ± 2.4 mV Vs -46 ± 2.5 mV n=4; p<0.05, paired t-test) SMCs and changed the action potential firing from a rhythmic pattern to continuous firing in tissues from control fed WT mice. NLA did not change membrane potential or action potential firing in SMCs from HF WT mice. SNP hyperpolarized (-51 ± 0.7 mV Vs -67 ± 1.8 mV n=3; p<0.05, paired t-test) the membrane potential and stopped action potential firing in colonic SMCs from WT HF. In β1BKKOs, SNP did not significantly   122     (p>0.05) hyperpolarize the membrane potential but changed action potential firing from continuous to bursting pattern in both β1BKKO control and β1BKKO HF tissues (Figure 4.5). Figure 4.5 NO modulates smooth muscle excitability. Effects of L-NNA (A and B) and SNP (C, D and E) on smooth muscle excitability. (A) WT HF mice show continuous action potential firing compared to WT on control diet which shows bursting like action potential firing behavior *p<0.05. (A) In WT controls, L-NNA depolarizes the membrane and increases the firing frequency (*p<0.05, paired t-test, n=3) (B) Application of L-NNA in WT HF does not depolarize the cell membrane. (C) Application of SNP in WT HF mice hyperpolarized the membrane and stopped action potential firing in a reversible manner. (D and E) BK KO mice irrespective of the diet had continuous action potential firing. Application of SNP on BK KO mice changed action potential firing to bursting like firing with no apparent change in RMP. (E) Application of SNP on BK KO mice fed a control diet changed action potential firing to a greater extent than HF tissues.   123     Figure 4.5 (cont’d) 4.4.7 WT HF mice exhibit disruptions in colonic migrating motor complex (CMMC) activity and CMMCs are disrupted in β1BK KO mice regardless of diet In order to test the hypothesis that HF diet disrupts in-vitro colonic motility, CMMCs were examined (Figure 4.6). I found that colonic contractions from control mice propagated aborally (anterograde) in all cases (100% anterograde). However, in HF mice only 64 ± 10% of contractions recorded at the oral end propagated anterogradely while 17 ± 8% of the contractions propagated retrogradely (anal to oral) and 6 ± 4% of contraction occurred simultaneously. The remaining (15 ± 10%) contractions did not propagate at all (i.e. they were seen either only in the oral segment or in the anal segment) (Figure 4.6B). The CMMC pattern was also disrupted in the colon from control and HF fed β1BK KO mice (Figure 4.6C). Oral and aboral contraction area was greater in WT HF than WT controls while no such difference were observed among KOs (Figure 4.6D and 4.6E). I found that the velocity of CMMC propagation was significantly faster in WT HF diet fed mice compared to WT controls (Figure 4.6F) (p<0.05).   124     Figure 4.6 In vitro recordings of colonic migrating motor complex (CMMC) in colonic segments from WT and β1-subunit KO mice placed on control and HF diet. (A) Representative recording of CMMC in the colon from a control fed mouse. Recording shows contractions propagate regularly in an oral to anal direction. (B) Summary of propagation patterns in colon segments from control (n=6) and HF fed (n=9) mice. Almost all of CMMCs propagate in anterograde direction in tissues from WT control fed mice while only 64% of CMMC propagate anterogradely in colon segments from WT HF fed mice. There was a significant increase in the percentage of retrogradely propagated contractions, and contractions that either occurred simultaneously or only at one recording site in the colon from HF mice (*p < 0.05). (C) CMMC propagation disrupted in β1-subunit KOs irrespective of the diet. Contraction area was greater in segments from WT HF compared to WT control fed mice at oral (D) aboral (E) ends of the segments, however no differences were detected among β1-subunit KOs. (F) CMMC velocity was faster (*p < 0.05) in colon segments from WT HF mice (AUC: Area under the curve) *p<0.05.   125     4.4.8 Consumption of a HF diet increases fecal pellet output in WT mice WT mice show significantly higher fecal pellet output per gram of body weight compared to β1BK KOs (Figure 4.7). Among WTs, mice fed a HF diet exhibit greater fecal pellet output compared to control fed. However, there was no difference in either the dry weight or the wet weight of the pellet among WTs and β1BK KOs. I also did not detect any diet dependent difference in either fecal pellet output, wet weight or the dry weight between β1BK KO mice fed a control and β1BK KO fed a HF diet. Figure 4.7 In vivo colonic motility measurements from WT and β1-subunit KO mice. WT mice had increased fecal output than β1BK KOs. Among WTs, mice fed a HF diet have increased fecal pellet output per gram of body weight than controls (n=5; *p < 0.05).   126     4.4.9 α-subunit protein expression of a β1BK channel is not altered in HF mice I did not find any difference in BKα subunit expression between WT control and HF mice. Although I had hypothesized that expression of β1subunit is altered in HF mice, I was unable to study the expression of β1subunit using western blot due to the lack of commercially available specific antibody for β1subunit (Bhattarai et al., 2014). Figure 4.8 BK channel α-subunit western blot. (A) The BK α-subunit antibody detected a protein band at ~110 kDa from the mouse distal colonic smooth muscle cell preparations. Antibody binding was blocked pre-incubating the primary antibody with its competing peptide and using secondary only controls. (B) Representative blot of BKCa α subunit and β-actin expression in colonic SMs of mice placed on control and HF diet. (C) No significant difference in BKCa α subunit / β-actin subunit protein expression was observed between WT controls and WT HFs.   127     4.4.10 β1-subunit mRNA expression is not changed in HF mice I used rt-PCR to detect changes in β1subunit gene expression between control and HF mice. However, I did not detect any significant change in the expression of either the α or the β1-subunit mRNA Figure 4.9 rt-PCR measurements for the α and the β1subunit mRNA expression. rt-PCR showed no difference in either (A) the α or (B) the β1 subunit expression between HF and controls   128     4.5 Discussion The aim of this study was to identify the pathogenesis of HF associated enteric neuron loss and altered SM electrical activity that contributes to GI dysmotility in obesity. My results show that HF diet disrupts inhibitory neuromuscular transmission, SM relaxation and excitability in WT but not β1BK KO mice. This effect is due to oxidative stress induced loss of NO neurons, which ultimately leads to impaired in-vitro and in-vivo colonic propulsive motility. Electrophysiological studies suggest that compromised BK channel β1-subunit function might play a role in mediating these disruptions. Consumption of a free fatty acid rich diet increases reactive oxygen species (ROS) generation via increased mitochondrial uncoupling and β-oxidation (Evans et al., 2002). NO neurons are susceptible to oxidative damage because NO reacts with superoxide anion to produce peroxynitrite (ONOO-), a highly reactive oxidizing agent (Murphy et al., 1998; Rivera et al., 2011). Since peroxynitrite reacts with reduced glutathione (GSH) to produce oxidized glutathione disulphide (GSSG), I quantified the ratio of oxidized/reduced (SS/SH) GSH as an indicator of oxidative stress (Murphy et al., 1998). I found that myenteric neurons in the colon of HF fed mice have an increased GSSG/GSH ratio suggesting increased oxidative stress. Increased in oxidative stress was associated with a decrease in the number of myenteric NOS neurons. Since, NOS neurons constitute ~25-40% of total myenteric neuron population (Bagyánszki et al., 2000; Bódi et al., 2009; Bagyánszki & Bódi, 2012), I tested to see if loss of NOS neurons reduced NO availability in the myenteric ganglia of HF mice. I found that mice placed on HF diet showed significant decreased benzotriazole product fluorescence intensity compared to control fed WT mice, suggesting lower NO bioavailability myenteric ganglia of HF fed mice.   129     I tested to see if oxidative stress, reduced NOS neurons and NO bioavailability disrupted inhibitory neuromuscular transmission and smooth muscle relaxation. Stimulation of enteric neurons at lower frequencies produced purinergic IJPs while higher stimulation frequencies causes mixed purinergic/nitrergic IJPs (Gallego et al., 2008). SMCs in the colon of HF fed mice had reduced IJP area but not peak amplitude indicating impairment of nitrergic but not purinergic signaling. I also found that impairment in nitrergic signaling is sufficient to cause disruptions in neurogenic SM relaxation in WT HF mice. Inhibition of neurogenic relaxation in the presence of NLA was also lower in HF fed WT mice compared to control fed WT mice suggesting lower NO availability. NO mediated relaxation occurs through activation of β1BK channel either via cyclic GMP-dependent pathway or via S-nitrosylation of cysteine residues (George & Shibata, 1995; Mistry & Garland, 1998; Jaffrey et al., 2001; Hess et al., 2005). I therefore tested to see if loss of β1 subunit in itself is sufficient to disrupt SM relaxation in GI tissues. I measured neurogenic relaxation from β1BK KO mice fed either a control or a HF diet. I found that in contrast to wild type mice, β1BK KO mice showed no diet dependent difference in the colonic SM relaxation and NLA dependent inhibition of relaxation was similar in both mice irrespective of the diet. These data suggest that NO possibly mediates SM relaxation by activating β1BK channel function in colonic smooth muscle. In vascular SMCs, oxidative stress decreases BK channel open probability in a dose dependent fashion by preferentially attacking calcium sensing cysteine residues (Brzezinska et al., 2000), which disrupts calcium sensing mechanism and causes channel to behave as if the β1 subunit had been deleted (Tang et al., 2004). However, the effect is reversible with addition of antioxidant, GSH (Brzezinska et al., 2000). I therefore hypothesized that HF diet dependent   130     oxidative stress contributes to changes in SM excitability via altered β1BK channel function. Using intracellular recordings, I found out that while control diet fed mice had significantly hyperpolarized SM membrane potential and fired action potentials in bursts, HF mice showed depolarized membrane potential and fired action potential continuously. Previous study done in our lab has shown that β1BK KO mice are constipated (France et al., 2012) and exhibit depolarized RMP and continuous action potential firing similar to WT HF mice. This suggests that HF diet might alter SM excitability in such a way that the HF tissues behaves as β1BK KO. Since HF diet has also been associated with lower NO availability I tested to see if NO availability contributes to colonic SM excitability. I found out that application of NLA in control diet fed mice depolarized the resting membrane potential of colonic SM (by ~3-5 mV) and increased action potential firing. This caused control mice to behave as HF. Application of SNP to tissues from HF fed mice hyperpolarized SMCs in a reversible manner. This results suggests that NO modulates SM excitability. Continuous action potential firing in colonic SMCs from HF fed mice are possibly due to loss of NO availability leading to subsequent SM depolarization and increased SM excitability. I repeated these experiment in β1BK KO mice to test if loss of the β1 subunit is sufficient to increase SMC excitability. Although I did not see any diet dependent difference (action potential firing were continuous in both cases), application of SNP restored bursting like action potential firing in β1BK KO. I also observed that in β1BK KO HF tissues, the number of action potentials that remained after SNP application were greater compared to control tissues suggesting greater excitability in these tissues compared to WT. These results are similar to studies carried in rat hippocampus where application of SNP restored the activity of BK channels in the NO deficient neurons caused by intermittent hypoxia (Tjong et al., 2008) suggesting that although that NO and β1BK channel share a common pathway to modulate the   131     SM excitability, application of NO is sufficient to reverse the changes caused by abnormal BK channel function. To assess if alterations in NO dependent neurogenic relaxation and impairment in SM excitability leads to disrupted colonic motility in-vitro, I measured CMMC activities in WT and β1BK KO mice fed a control and a HF diet. CMMCs are waves of contractile activity that occurs at regular interval in fasting state. I found that the velocity, directionality and the force of contraction were all disrupted in WT HF fed mice and β1BK KO (irrespective of diet) compared to WT control. Application of NLA disrupted the frequency, the tone, and the directionality of the contraction in both controls and HF WT mice suggesting that loss of endogenous NO in WT HF mice disrupts in-vitro motility (Powell & Bywater, 2001). The changes that I saw in CMMC activity in WT and β1BK KO mice correspond to changes in in-vivo colonic motility. This was evident by the fact that the fecal pellet output in WT mice was significantly greater than β1BK KOs. Among WTs, mice placed on HF diet had greater fecal output than controls while no such difference was observed among β1BK KO. NO is involved in rapid anterograde trafficking of β1 subunit from the endosomes to the cell membrane and helps β1 subunit to associate with BKα subunit elevating channel Ca2+ sensitivity in myocytes and inducing vasodilation in vascular SMCs (Leo et al., 2014). Since I have shown that NO levels are lower in HF mice, it is possible that mechanisms that lead to the trafficking of β1 subunit to cell membrane are disrupted leading to disruptions in SMC excitability. Besides this, decreased BK channel β1/α subunit ratio, and altered Ca2+ spark generation has been observed in cerebral arteries of db/db mice (Rueda et al., 2013). I therefore hypothesized that HF diet dependent oxidative stress and lower NO availability leads to changes in β1 subunit protein expression of BK channel ultimately causing altered colonic SM   132     excitability. Although I showed no change in expression of the BK α-subunit, I was unable to study the expression of β1subunit using western blot analysis due to the lack of specificity in commercially available antibodies for β1subunit (Bhattarai et al., 2014). I did not find any dietinduced changes in β1subunit gene expression using rt-PCR. This suggests that if there is any diet dependent changes that occurs in β1subunit, they are downstream of mRNA transcription (i.e. disruptions in β1 trafficking, changes in post translational modification). These studies indicate that HF diet alters colonic motility via oxidative stress induced loss of NO motor neurons, and by causing changes in SM excitability. Although the results of this study insinuate that β1BK channel might play a crucial role in modulating these changes, measurement of β1BK protein expression is necessary to conclusively validate this point.   133     BIBLIOGRAPHY   134     BIBLIOGRAPHY Bagyánszki M & Bódi N. (2012). 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Selective responses of myenteric neurons to oxidative stress and diabetic stimuli. Neurogastroenterol Motil 23, 964-e411.   138     Wan E, Kushner JS, Zakharov S, Nui XW, Chudasama N, Kelly C, Waase M, Doshi D, Liu G, Iwata S, Shiomi T, Katchman A, D'Armiento J, Homma S & Marx SO. (2013). Reduced vascular smooth muscle BK channel current underlies heart failure-induced vasoconstriction in mice. FASEB J 27, 1859-1867. Xing J & Chen JD. (2004). Alterations of gastrointestinal motility in obesity. Obes Res 12, 17231732. Xu H, Garver H, Fernandes R, Galligan JJ & Fink GD. (2014). Altered L-type Ca2+ channel activity contributes to exacerbated hypoperfusion and mortality in smooth muscle cell BK channel deficient septic mice. Am J Physiol Regul Integr Comp Physiol. Xu H, Garver H, Galligan JJ & Fink GD. (2011). Large-conductance Ca2+-activated K+ channel beta1-subunit knockout mice are not hypertensive. Am J Physiol Heart Circ Physiol 300, H476-485. Xu H, Wang Y, Garver H, Galligan JJ & Fink GD. (2012). Vascular BK channel deficiency exacerbates organ damage and mortality in endotoxemic mice. J Cardiovasc Pharmacol 59, 207-214. Y Lee C. (2013). The Effect of High-Fat Diet-Induced Pathophysiological Changes in the Gut on Obesity: What Should be the Ideal Treatment? Clin Transl Gastroenterol 4, e39. Yang Y, Li PY, Cheng J, Mao L, Wen J, Tan XQ, Liu ZF & Zeng XR. (2013). Function of BKCa channels is reduced in human vascular smooth muscle cells from Han Chinese patients with hypertension. Hypertension 61, 519-525.   139     CHAPTER 5 WESTERN BLOT ANALYSIS OF BK CHANNEL β1-SUBUNIT EXPRESSION SHOULD BE INTERPRETED CAUTIOUSLY WHEN USING COMMERCIALLY AVAILABLE ANTIBODIES   140     5.1 Abstract Large conductance Ca2+ activated K+ (BK) channels consist of pore forming α- and accessory β-subunits. There are 4 β-subunit subtypes (β1-β4), and the BK β1-subunit is specific for smooth muscle cells (SMC). Reduced BK β1-subunit expression is associated with SMC dysfunction in animal models of human disease, since down-regulation of BK β1-subunit reduces channel activity and increases SMC contractility. Several anti-BK β1-subunit antibodies are commercially available. However, the specificity and the sensitivity of most antibodies have not been tested or confirmed in the tissues from BK β1-subunit knockout (KO) mice. In this study, I tested the specificity and sensitivity of six commercially available antibodies from five manufacturers. I performed western blot analysis on BK β1-subunit enriched tissues (mesenteric arteries and colons) and non-SM tissue (cortex of kidney) from wild-type (WT) and BK β1-KO mice. I found that all tested antibodies lack specificity for BK β1-subunits in murine tissues. Antibodies either detected protein bands of the appropriate molecular weight in tissues from both WT and BK β1-KO mice, or failed to detect protein bands at the appropriate molecular weight in tissues from WT mice. The absence of BK β1-subunit mRNA expression in arteries, and colons from BK β1-KO mice was confirmed by RT-PCR analysis. I conclude that these commercially available antibodies are not reliable tools for studying BK β1-subunit expression in murine tissues. Data obtained using these antibodies should be interpreted cautiously. Developing a highly specific BK β1-subunit antibody is needed for future studies of BK channel murine models of human disease. 141       5.2 Introduction BK (large conductance Ca2+-activated K+) channels are expressed in many mammalian cell types and BK channel activity is an important regulator of cell excitability. BK channels are composed of pore forming α-subunits and accessory β-subunits that modulate α-subunit Ca2+ sensitivity and channel activity (Nelson et al., 1995; Brenner et al., 2000b). There are 4 subtypes (β1-β4) of BK channel β-subunits. The BK β1-subunit is largely smooth muscle cell (SMC) specific, although it has been found in the distal nephron of murine kidney (Grimm et al., 2007), BK β2- and β3-subunits are expressed in endocrine cells (Xia et al., 1999; Braun et al., 2008), while BK β4-subunits are expressed in neurons (Meera et al., 2000) and in the distal nephron of the kidney (Grimm et al., 2007). In SMCs, BK channels stabilize membrane potential and excitability through negative-feedback modulation. Activation of BK channels causes SMC hyperpolarization, L-type Ca2+ channel closure and SMC relaxation (Nelson et al., 1995). Down-regulation of BK β1-subunit expression may contribute to human diseases including metabolic disorders (Zhang et al., 2010a), hypertension (Yang et al., 2013), heart failure (Wan et al., 2013), asthma (Semenov et al., 2011), maladaption of uteroplacental circulation (Hu et al., 2012), urinary bladder over activity (Petkov et al., 2001) and gastrointestinal motility disorders (France et al., 2012). Down-regulation of BK β1-subunit expression reduces α-subunit Ca2+ sensitivity, therefore, causing SMC membrane depolarization and increased muscle contractility (Brenner et al., 2000b; Plüger et al., 2000; Xu et al., 2011). BK β1-subunit expression in pathological situations is mainly affected by signaling at the post-transcriptional level where protein synthesis is reduced (Nelson et al., 1995; Grimm et al., 2009; Xu et al., 2011; Shi et al., 2013a; Shi et al., 2013b) or protein degradation is accelerated (Zhang et al., 2010a; Lu et al., 142       2012). Therefore, accurate measurement of BK β1-subunit protein expression is crucial in studies of BK channel dysfunction in diseases. The anti-BK β1-subunit antibodies used in published studies are either custom made (Lu et al., 2012; Yi et al., 2014) or purchased commercially. BK β1-subunit protein expression is typically measured using western blot analysis. Importantly, the specificity and sensitivity of commercially available BK β1-subunit antibodies has not been tested or confirmed in tissues from BK β1-subunit knockout (KO) mice, although BK β1-KO mice have been generated by two research groups (Brenner et al., 2000b; Plüger et al., 2000). BK β1-KO mice have been widely used as an animal model to study the contribution of BK channel dysfunction in hypertension (Grimm et al., 2007; Xu et al., 2011), septic shock (Xu et al., 2012; Xu et al., 2014), heart failure (Wan et al., 2013), asthma (Evseev et al., 2013), metabolic disorders (Lynch et al., 2013), irritable bowel syndrome (France et al., 2012) and urinary bladder over activity (Petkov et al., 2001). Conclusions about the physiological significance of BK β1-subunits in health and disease have been based in part on studies in mice. I used western blot analysis to test the specificity and sensitivity of six commercially available BK β1-subunit antibodies from five manufacturers in BK β1-subnuit enriched SM tissues (mesenteric arteries, MA, and colons) from wild-type (WT) and BK β1-KO mice (Brenner et al., 2000b; Grimm et al., 2007). I confirmed that BK β1-KO mice lack β1-subunit mRNA using RT-PCR analysis. 143       5.3 Materials and Methods 5.3.1 Ethics Statement All studies were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Publication No. 85-23, revised 1996) and approved by the Michigan State University Institutional Animal Care and Use Committee. 5.3.2 Animals Homozygous breeder male and female BK β1-KO mice were a gift from Dr. Robert Brenner (University of Texas Health Science Center, San Antonio). BK β1-KO mice are congenic as a result of seven generations of inbreeding to the C57BL/6 line and maintained originally as homozygous lines at Michigan State University (Xu et al., 2011; France et al., 2012; Xu et al., 2012; Xu et al., 2014). Pups of BK β1-KO mice were weaned at 3 weeks. Age matched WT (C57BL/6) mice were purchased from Jackson Laboratories (Bar Harbor, Maine, USA). All mice were fed a normal diet. Mice used in our studies were at 20-24 weeks of age (male). 5.3.3 Western blot Colons and MA from WT and BK β1-KO mice were homogenized and lysed with 1X lysis buffer (12.5 ml 0.5M Tris HCl, 20 ml 10% SDS, 10 ml Glycerol, 57.5 ml dH2O) along with a protease inhibitor cocktail (P8340, Sigma), sodium fluoride (S7920, Sigma), and sodium orthovanadate (S6508, Sigma). Protein levels were quantified with Sigma’s Bicinchoninic Assay (BCA1 and B9643). Samples were reduced with Laemmli Buffer (Bio-Rad 161-0737) and βmercaptoethanol and boiled for 5 minutes at 95 - 100 °C. Protein (30-100 mg for MA and colons, 500 mg for kidney) were loaded on SDS-PAGE (10-12%) gel along with the full-range rainbow molecular weight marker (RPN800E, GE Healthcare); separated at 100-120 V with 144       Laemmli running buffer (6 g Tris base, 28.8 g glycine, 1 g SDS, 1L dH20), and electrotransferred overnight at 30 V onto Nitrocellulose membranes (RPN68D, Amersham Hybond-ECL, USA) or PVDF (IPVH00010, Milipore, USA) overnight, or at 100 V for 1 hour, with transfer buffer (3 g Tris base, 14.4 g glycine, 200 ml methanol, 800 ml dH2O). Ponceau S was used to verify efficient protein transfer. Samples were blocked for 1 hour at room temperature in 5% milkTBS-T, 5% BSA-TBS-T, or 4% chicken ovalbumin-TBS-T. I tested six commercially available BK β1-subunit antibodies from five manufacturers (Table 5.1). 145       Table 5.1 List of commercially available anti-BK β1-subunit antibodies. Reprinted from Physiological reports, Volume 2, Bhattarai et.al., Western blot analysis of BK channel β1subunit expression should be interpreted cautiously when using commercially available antibodies, copyright (2014). Manufacturer Catalog # Immunogen Reactivity 2-17 MW (kDa) 28 Alomone APC-036 Abcam Ab3587 90-103 35 H, R, M Pierce PA1-924 90-103 28 H, R, M Pierce PA528284 1-191 28 H, M Santa Cruz sc-14749 1-191 28 H, R, M Biorbyt orb101774 10-80 21 H, R, M H, R, M Concentratio Secondary Ab n 1:100-1000 Anti-Rabbit 1:2000 (sc-2313; 7074S) 1:500 Anti-Rabbit 1:2000 (#7074S) 1:100-1000 Anti-Rabbit 1:2000 (sc-2313; 7074S) 1:100-1000 Anti-Rabbit 1:2000 (sc-2313) 1:200 Anti-Goat 1:500 (sc-2056) 1:100-500 Anti-Rabbit 1:2000 (sc-2313; 7074S) MW, manufacturer recommended molecular weight; H, human; R, rat; M, mouse; sc-1213, donkey anti-rabbit IgG-conjugated to horseradish peroxidase, Santa Cruz Biotechnology; sc2056, donkey anti-goat IgG-conjugated to horseradish peroxidase, Santa Cruz Biotechnology; 7074S, goat anti-rabbit IgG-conjugated to horseradish peroxidase, Cell Signaling. 146       I also tested an anti-BK α-subunit antibody (APC-107, anti-KCa1.1, 1:500, Alomone Labs) in protein extracts from colons and MA to confirm BK channel expression in these tissues. Protein loading and transfer were confirmed by re-blotting for β-actin (A228, Anti-β-Actin, 1:1000, Sigma) on each membrane after the BK β1-subunit antibody was stripped. Membranes were blotted overnight at 4°C or 2 hours at room temperature, with anti BK α- or β1-subunit primary antibodies used at the manufacturers’ suggested dilutions and after primary antibody incubation with antigen peptide (1:100) overnight (Table 5.1). The following day membranes were washed 3 X 10 minute intervals with TBS-T, and blotted with the appropriate HRPconjugated secondary antibodies (Table 5.1) for 1 hour at room temperature in blocking solution. Membranes were then washed with TBS-T for 3 X 10 minutes. Blots were imaged with Super Signal West Dura Extended Duration Substrate (34076, Pierce) using a LICOR-FC imager, or using the Enhanced Chemiluminescence (ECL) Plus Western Blotting Detection System (RPN2133, GE Healthcare). 5.3.4 Real time RT-PCR Samples were collected and stored in All Protect Reagent (76405, Qiagen, USA) until ready to be processed. Total RNA was isolated with the MELT Total Nucleic Acid Isolation System kit (AM1983, Ambion, USA) following the manufacturer’s instructions. RNA isolates were reverse-transcribed using High Capacity RNA to cDNA kits (4387406, Applied Biosystems, USA), following the manufacturer’s protocols. As a control for genomic DNA contamination, all cDNA synthesis reactions were set up with additional samples lacking reverse transcriptase. Resultant cDNA was used for real-time PCR assays. Reactions (20 ml) were prepared with TaqMan Fast Advanced master mix (4444556, Applied Biosystems, USA) and inventoried with TaqMan Gene Expression assays (Mm00466621_m1 for KCNMB1, 147       Rn01775763_g1 for Gapdh, Applied Biosystems, USA). Samples were analyzed in duplicate. Real time RT-PCR products at 40 cycles were also determined by agarose gel analysis (1.5% TBE agarose gel at 100V for 1 hour). 148       5.4 Results 5.4.1 Western blots in MA and colons from WT and BK β1-KO mice In SM tissues, APC-036 (Alomone Labs) and PA5-28284 (Thermo Scientific-Pierce) anti-BK β1-subunit antibodies detected protein bands at ~28 kDa (manufacturer suggested molecular weight of BK β1-subunit protein) in MA and colonic tissues from WT and BK β1-KO mice, (Figure 5.1A, 5.1B). Ab3587 (Abcam) anti-BK β1-subunit antibody detected a protein band at ~38 kDa (manufacturer’s suggested molecular weight of BK β1-subnit protein) in MA and colons from WT and BK β1-KO mice (Figure 5.1C). Signals were blocked after preincubation of the Ab3587 antibody with its competing peptide (Ab5023) (Figure 5.1C). PA1924 (Thermo Scientific-Pierce) anti-BK β1-subunit antibodies detected a protein band at ~40 kDa in MA and colons from WT and BK β1-KO mice, but not at ~28 kDa (manufacturer suggested molecular weight of BK β1-subunit protein) (Figure 5.1D). The sc-14749 (Santa Cruz Biotechnology) and Orb-101774 (Biorbyt) antibodies detected multiple protein bands in MA and colonic tissues from WT and BK β1-KO mice. However, none of these protein bands corresponded to a protein with a molecular weights of ~28 kDa or ~21 kDa (the manufacturer’s suggested molecular weight of BK β1-subunit protein) in tissues from WT mice, even when gels were loaded with 80 mg of protein and incubated with high concentration of primary antibody (1:100) (Figure 5.1E, 5.1F). Protein loading and transfer were confirmed by β-actin immune blot (Figure 5.1E). I repeated sc-14749 and Orb-101774 immunoblotting using a variety of protocols (See Methods). Results obtained using modified approaches were similar to the results described above. Tissues from 6-8 animals were tested in each group. 149       Figure 5.1 Representative western blot obtained using commercially available anti-BK β1subunit antibodies. (A) Alomone Labs (APC-036), (B) Pierce (PA5-28284), and (C) Abcam (Ab3587) antibodies detected a protein band at ~28 kDa or ~38 kDa in colons or MA from both WT and BK β1-KO mice. The bands in Abcam sets were diminished after preincubation of the primary antibody with the competing peptide. (D) Pierce (PA1-924), (E) Santa Cruz (sc-14749), and (F) Biorbyt (orb-101774) antibodies did not detect any band at ~28 kDa or ~21 kDa in MA or colons from WT mice. Arrows indicate the manufacturer's recommended molecular weight of BK β1-subunit protein. Reprinted from Physiological reports, Volume 2, Bhattarai et.al., Western blot analysis of BK channel β1-subunit expression should be interpreted cautiously when using commercially available antibodies, copyright (2014). 5.4.2 BK β1-subunit mRNA expression in tissues from BK β1-KO mice To confirm the BK β1-subunit gene has been deleted in BK β1-KO mice, I measured expression of BK β1-subunit mRNA levels in MA and colons from WT and BK β1-KO mice using real time RT-PCR. After 40 PCR cycles, the Ct values of BK β1-subunit mRNA levels in MA and colon from WT mice were ~25 and ~24 respectively (Figure 5.2A, 5.2B) but BK β1- 150       subunit mRNA was undetectable in tissues from BK β1-KO mice (Figure 5.2C). Tissues from 68 animals were tested in each group. Figure 5.2 Real-time RT-PCR analysis of BK β1-subunit and GAPDH in MA and colons from WT and BK β1-KO mice. (A and B) The expression threshold was set at 0.22 for amplification plots, a level above background fluorescence but within the linear phase of the amplification plot. The intersection between the threshold level and the amplification plot is the Ct value, which correlates with the amount of template in the sample. Ct values over 35 are excluded, as these values approach the sensitivity limits of the Taqman assay. (C) Agarose gel separation of RT-PCR products. Amplification of real-time RT-PCR products was seen at 75 bp in tissues from WT animals only. Reprinted from Physiological reports, Volume 2, Bhattarai et.al., Western blot analysis of BK channel β1-subunit expression should be interpreted cautiously when using commercially available antibodies, copyright (2014). Abbreviation: Con, non-template control. 5.4.3 Expression of BK α-subunit in the colons APC-107 anti BK α-subunit detected a protein band at ~100 kDa (manufacturer suggested molecular weight) in colonic tissues from WT and BK β1-KO mice. Western blot 151       signals were blocked after pre-incubation of the APC-107 antibody with its competing peptide (Figure 5.3). Figure 5.3 BK α-subunit western blot in colonic tissues from WT and BK β1-KO mice. Antibody detected a protein band at ~100 kDa in all tissues from WT and BK β1-KO mice. The signals are blocked by pre-incubation with the antibody competing peptide (CP). Arrows indicate the manufacturer's recommended molecular weight of BK α-subunit protein. Reprinted from Physiological reports, Volume 2, Bhattarai et.al., Western blot analysis of BK channel β1subunit expression should be interpreted cautiously when using commercially available antibodies, copyright (2014). 152       5.5 Discussion I tested the specificity and sensitivity of commercially available BK β1-subunit antibodies using tissues from WT and BK β1-KO mice, and found that each antibody lacked specificity and the sensitivity for BK β1-subunit in BK β1-subunit enriched tissues from C57BL/6 mice. The antibodies evaluated in this paper have been used extensively in published work (Table 5.2). However, this study is the first to determine their specificity for detection of the BK β1-subunit using BK β1-subunit KO mice. 153       Table 5.2 Publications that performed BK β1-subunit western blot analysis using commercially available antibodies. Antibody Alomone APC-037 References # Model (Zheng et al., H, R, M 2013),(Shi et al., 2013a),(Zhang et al., 2010a),(Shi et al., 2013b),(Kunduri et al., 2013),(Rueda et al., 2013),(Pabbidi et al., 2014) Abcam (Leo et al., M Ab-3587 2014),(Nystoriak et al., 2014), (Evanson et al., 2014) Santa Cruz (Grimm et al., H, R, M SC-14749 2007),(Ahn et al., 2012), (Xie et al., 2010),(Loot et al., 2012),(Grimm et al., 2009) Pierce, (Chang et al., H, R, M PA1-924 2006),(Evseev et al., 2013),(Yang et al., 2013),(Albarwani et al., 2010) Customer (Yi et al., H, M Made 2014),(Lu et al., 2012) Merck (Yang et al., H, R 2009),(Howitt et al., 2011),(MatharooBall et al., 2003), H, human; R, rat; M, mouse. Tissue Type Reported MW Blood Vessel 24-28 kDa Used BK β1 KO mice NO Blood Vessel 35-45 NO Blood Vessel, Adrenal Gland 22-28 kDa Yes (Grimm et al., 2007), (Loot et al., 2012), (Grimm et al., 2009) Blood Vessel, SMC from Tracheal 37 kDa NO Blood Vessel 32kDa NO Blood vessel, Myometrium 32-36 NO 154       The BK β1-KO mice used in our study were generated by using a viral vector (pPNT) to completely delete exon 2 of gene 27, which also includes a transcriptional terminator after the lacZ gene to prevent downstream expression of β1-subunits (Brenner et al., 2000b). Absence of BK β1-subunit expression in the BK β1-KO mice has been confirmed by RT-PCR previously (Brenner et al., 2000b) and in this study. BK β1-subunit specific antibodies should identify a protein band with a molecular weight of 21 - 35 kDa (depending on the supplier’s recommended molecular weight) in BK β1-subunit enriched tissues obtained from WT mice. BK β1-subunit protein expression should either be absent, or it would be detected as a truncated lower molecular weight protein in tissues from BK β1-KO mice, if the terminator is not fully functional, and the immunogenic site for each antibody was expressed. However, all tested antibodies either failed to detect proteins at the appropriate molecular weight in tissues from WT mice, or the antibodies detected identical protein bands in tissues from WT and BK β1-KO mice. These results suggest that the proteins detected by these antibodies are not specific for the BK β1-subunit; even though some of bands could be blocked by pre-incubation of the primary antibodies with its competing peptide. I tested six antibodies not only in BK β1-subunit specific SM tissues (arteries and colons), but also tested in kidneys, a BK β1-subunit enriched non-SM tissue (Grimm et al., 2007). Results from all tissues indicate that none of antibodies can reliably detect the BK β1subunit. I confirmed protein loading and transfer performance on these blots by detection of βactin on the same membrane. I also reliably detected the BK a-subunit protein in these tissues by western blot. Finally I confirmed BK β1-subunit gene deletion in the BK β1-KO mice using real time RT-PCR on the same tissue samples used for western blot analysis. 155       Two groups previously western blot data using the sc-14749 antibody in tissues from BK β1-KO mice. Loot et al. studied BK β1-subunit expression in cultured pulmonary arterial SMCs from WT and BK β1-KO mice (Loot et al., 2012). However, the mouse strain used to generate the BK β1-KO mice used in their study differs from ours (Brenner et al., 2000b; Plüger et al., 2000). Those BK β1-KI mice were generated on B6CBAF1/J (female) and deleted exon 1 of BK β1-subunit gene. In addition, the protein bands obtained from extracts of SMCs from WT mice were not well resolved, and molecular weight of the protein detected was lower than the manufacturer’s recommendation (<25 kDa). Grimm et al. reported a ~28 kDa protein band in extracts from kidney cortex and adrenal glands from WT mice, which was absent in tissues from BK β1-KO mice (Grimm et al., 2007; Grimm et al., 2009). The BK β1-KO mice used in this study were generated from the same mouse strain used in our studies. When I used the Sc-14749 antibody I failed to detect a protein band at the predicted molecular weight (~28 kDa) in protein extracts from MA or colons from WT mice. Studies have been done using custom-made antibodies (Lu et al., 2012; Yi et al., 2014) or an antibody produced by the Merck pharmaceutical company (Matharoo-Ball et al., 2003; Yang et al., 2009; Howitt et al., 2011) (also see Table 5.2). The antibody from Merck is no longer available and I was unable to test this antibody. 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Regulation of Large Conductance Ca2+-activated K+ (BK) Channel β1 Subunit Expression by Muscle RING Finger Protein 1 in Diabetic Vessels. J Biol Chem 289, 10853-10864. Zhang DM, He T, Katusic ZS, Lee HC & Lu T. (2010). Muscle-specific f-box only proteins facilitate bk channel β(1) subunit downregulation in vascular smooth muscle cells of diabetes mellitus. Circ Res 107, 1454-1459. Zheng YM, Park SW, Stokes L, Tang Q, Xiao JH & Wang YX. (2013). Distinct activity of BK channel β1-subunit in cerebral and pulmonary artery smooth muscle cells. Am J Physiol Cell Physiol 304, C780-789. 161       CHAPTER 6 KEY FINDINGS, SIGNIFICANCE AND FUTURE DIRECTIONS   162     Key findings In the first part of my dissertation I advanced the field by identifying neurotransmitters, receptors, and ion channels involved in mediating IJPs in the mouse distal colon. I found that ATP and NO ultimately act on SK channels and CaCCs to mediate inhibitory neuromuscular transmission in the mouse distal colon. The opposing effect of K+ and Cl- ion regulates both the IJP and the membrane potential of the smooth muscle. Figure 6.1 Mechanism of IJP production in the mouse distal colon. ATP acts on GQ coupled P2Y1 receptors to increase intracellular Ca2+concentration [Ca2+i] while NO mediates PKG activation. Increase in [Ca2+i] and PKG activation causes opening of SK channels and closure of CaCCs to produce IJPs.   163     In the second part of the study, I found that β1BK channel regulates the RMP and the action potential firing in the mouse distal colon. Disruption or genetic manipulation leading to loss of β1 subunit increases smooth muscle excitability and impairs colonic motility (France et al., 2012). Figure 6.2 Regulation of colonic smooth muscle excitability via β1BK channel function. Depolarizing stimulus in the smooth muscle activate voltage gated Ca2+ channels. (1) Influx of Ca2+ into the cell and efflux of K+ from the cell causes smooth muscle action potential generation. Continuous influx of Ca2+ in the cell activates ryanodine receptors in the SER to produce Ca2+ sparks thereby increasing subplasmalemmal Ca2+ concentration above ≥ 1µM. Such increase in Ca2+ concentration activates β1BK channels. (3) β1BK channels activation causes quick efflux of K+ out of the cell causing membrane hyperpolarization. Membrane hyperpolarization leads to closure of voltage gated Ca2+ channels and abolishes action potential generation causing cessation of burst in WT DC.   164     In the third part of my dissertation I investigated in detail the etiology of colonic dysmotility caused by HF diet. I found that HF diet causes oxidative stress, loss of inhibitory NOS neurons, and decreases basal NO availability in the mouse colonic myenteric ganglia. This causes disruption in neuromuscular transmission, impairment in colonic smooth muscle relaxation and alterations in colonic smooth muscle excitability. NOS neuron loss in HF fed mice alters mouse colonic smooth muscle excitability in such a way that it recapitulates changes in smooth muscle excitability seen in β1BK KO mice. This suggests that HF diet might affect β1BK channel function. However, I could not detect whether β1 subunit expression is down regulated in HF diet fed mice due to lack of commercially available β1 subunit specific antibodies. Figure below summarizes the details of the investigation. Figure 6.3 Mechanisms of colonic dysmotility due to HF diet consumption. Dotted line indicates that β1BK channel function/expression might be altered in HF diet fed mice.   165     Significance My study provides a comprehensive overview how IJPs are regulated in the mouse distal colon. Using a novel ectoneuclotidase, POM-1, I verified that ATP is the primary neurotransmitter involved in mediating IJP. My study also suggests that development of drugs that target CaCCs could benefit patients with impaired colonic motility. My study demonstrates that β1BK channels and NO regulate smooth muscle excitability by regulating the RMP and action potential firing activity in the mouse distal colon. I show HF diet causes oxidative stress induced loss of NOS neurons and lower NO availability in mouse colonic myenteric ganglia. Loss of NOS neurons contributes to impaired inhibitory neuromuscular transmission, disrupted smooth muscle relaxation and alterations in smooth muscle excitability. This suggests that antioxidants or drugs that target the NO synthesis pathway might have beneficial effects on patients with colonic motility complications. I also show that disruption in smooth muscle relaxation, electrical activity and colonic dysmotility seen in HF fed mice is similar to β1BK KO mice. These results suggest that β1BK KO mice could serve as an excellent model for testing of prokinetic drugs for obese patients with altered colonic motility. Finally, my study cautions investigators to use appropriate positive and negative controls when using commercially available antibodies in western blot experiments to avoid erroneous conclusion. I highlight that identifying protein bands at manufacturers recommended molecular weight and using blocking peptide is not sufficient to conclude that a protein of interest has been detected. Using β1BK KO mice I emphasized that, appropriate negative control such as a KO animal models must be used to test the specificity of the commercially available antibodies. This is especially important in study of diseases where altered protein expression might have a pronounced effect.   166     Future directions In the first part of the dissertation, I concluded that the CaCCs and the SK channels mediate inhibitory neuromuscular communication and smooth muscle RMP in the mouse distal colon. These conclusions are based on results obtained from electrophysiolgical studies where pharmacologic application of apamin and NFA blocked IJPs and disrupted the RMP in the colonic smooth muscles. Although NFA is a CaCC blocker, some studies suggests that it also blocks nonspecific cation channels (Gögelein et al., 1990), L type Ca2+ channels (Doughty et al., 1998) and can activate Ca2+ activated K+ channels (Toma et al., 1996; Cruickshank et al., 2003). Thus it is possible that results obtained from NFA could be due to an indirect effect on cation or Ca2+ activated K+ channels. It would therefore be necessary to see how modulation of extracellular Cl- concentration in the presence of NFA impacts the IJPs and the membrane potential of the colonic smooth muscle in order to verify that disruption in IJPs and the RMP are due to closure of CaCCs. My studies show that HF diet causes oxidative stress concomitant with nNOS neuron loss, disrupted neuromuscular communication and altered smooth muscle excitability. However, it is important to investigate if feeding HF mice with antioxidants can reverse NO neuron loss and disrupted neuromuscular transmission to conclusively verify if oxidative stress causes NO neuron loss. It would also be interesting to investigate whether neuromuscular transmission and smooth muscle excitability in nNOS KO mice replicates our HF mice model. Besides smooth muscle, NO also acts on interstitial cells of Cajal (ICC) to modulate IJPs (Lies et al., 2014). Therefore, it is possible that disruptions in function/expression of the ICCs, but not the NOS neurons causes impairment in neuromuscular transmission seen in HF mice. One way to tackle this issue is by measuring Anoctamin-1 (Ano1, ICC specific CaCCs protein) protein expression and by recording IJPs from HF mice in presence of NFA.   167     Lastly, most of my conclusions on the role of β1BK channels are based on electrophysiological studies. These data needs to be verified using proper expression studies such as rt-PCR and western blot before important conclusions can be drawn. Continued development and exploration of β1BK antibodies for studies of β1BK channel expression and function is therefore crucial.   168     BIBLIOGRAPHY   169     BIBLIOGRAPHY Cruickshank SF, Baxter LM & Drummond RM. (2003). The Cl(-) channel blocker niflumic acid releases Ca(2+) from an intracellular store in rat pulmonary artery smooth muscle cells. Br J Pharmacol 140, 1442-1450. Doughty JM, Miller AL & Langton PD. (1998). Non-specificity of chloride channel blockers in rat cerebral arteries: block of the L-type calcium channel. J Physiol 507 ( Pt 2), 433-439. France M, Bhattarai Y, Galligan JJ & Xu H. (2012). Impaired propulsive motility in the distal but not proximal colon of BK channel β1-subunit knockout mice. Neurogastroenterol Motil 24, e450-459. Gögelein H, Dahlem D, Englert HC & Lang HJ. (1990). Flufenamic acid, mefenamic acid and niflumic acid inhibit single nonselective cation channels in the rat exocrine pancreas. FEBS Lett 268, 79-82. Lies B, Gil V, Groneberg D, Seidler B, Saur D, Wischmeyer E, Jiménez M & Friebe A. (2014). Interstitial cells of Cajal mediate nitrergic inhibitory neurotransmission in the murine gastrointestinal tract. Am J Physiol Gastrointest Liver Physiol 307, G98-106. Toma C, Greenwood IA, Helliwell RM & Large WA. (1996). Activation of potassium currents by inhibitors of calcium-activated chloride conductance in rabbit portal vein smooth muscle cells. Br J Pharmacol 118, 513-520.   170