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Adenosine A₁ and A₂₄ receptors in the rat stomach : biological actions, cellular localization, structure.. Yip, Linda 2004

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ADENOSINE A, AND A2A RECEPTORS IN THE RAT STOMACH: BIOLOGICAL ACTIONS, CELLULAR LOCALIZATION, STRUCTURE AND GENE EXPRESSION by LINDA YIP B.Sc, The University of British Columbia, 1997 A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF THE FACULTY OF GRADUATE STUDIES (Department of Physiology) We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH COLUMBIA April 2004 DOCTOR OF PHILOSOPHY in © Linda Yip, 2004 Abstract Adenosine has been shown to inhibit gastric acid secretion in the rat stomach, likely by regulating gastrin and somatostatin release. The current study demonstrates that Ai and A2A receptors may play significant roles in this action. Specifically, in the isolated vascularly perfused rat stomach, the effect of various selective adenosine agonists and antagonists suggests that the inhibition of immunoreactive gastrin (IRG) and somatostatin-like immunoreactivity (SLI) release is mediated by Ai receptors, while the stimulation of SLI release is mediated by A2A receptors. To determine the site at which adenosine acts to elicit these actions, immunohistochemistry was performed. Both Ai receptor immunoreactivity (AiR-IR) and A2A receptor immunoreactivity (A2AR-IR) were observed on mucosal somatostatin-containing D-cells. AiR-IR was also observed on gastrin-containing G-cells, while neither AiR-IR nor A2AR-IR was observed on parietal cells. These results suggest that adenosine does not act directly on the parietal cells to inhibit gastric acid secretion. Instead, adenosine may act on Ai receptors of G-cells and D-cells to inhibit IRG and SLI release, respectively, and may act on A2A receptors of D-cells to stimulate SLI release. The localization of AiR-IR and A2AR-IR in the gastric plexi suggests that adenosine may also act indirectly on the gastric plexi by altering enteric neural transmission to regulate IRG and SLI release. The coding regions of the gastric Ai and A2A receptors were also examined and found to be structurally identical to those in the rat brain, but expressed in extremely low levels. Gene expression was measured using Real-Time reverse-transcriptase polymerase chain reaction (RT-PCR) assays developed in the current study. Using these assays, Ai and A2A receptor gene expression was found to be altered by fasting and omeprazole treatment. Ill These treatments produced reciprocal changes in Ai receptor and gastrin gene expression, and similar changes in A2A receptor and somatostatin gene expression. Further studies demonstrate that omeprazole treatment also altered adenosine agonist-induced changes in SLI release, suggesting that changes in adenosine receptor gene expression may result in actual changes in receptor expression. These results suggest that changes in the gastric state may alter Ai and A2A receptor expression, which in turn regulates the synthesis and release of gastrin and somatostatin, and consequently gastric acid secretion. Table of contents Abstract ii Table of contents v List of tables viii List of figures ix Acknowledgements xiii Chapter 1: Introduction 1 I. Background 2 1. Regulation of endogenous adenosine levels 2 1.1 The de novo synthesis pathway 3 1.2 Intracellular and extracellular salvage1.3 Presynaptically and postsynaptically released ATP as a source of adenosine 8 1.4 Cellular release and reuptake of adenosine by adenosine transporters 9 1.5 Metabolism of adenosine 10 1.6 Endogenous adenosine levels 2 2. The adenosine receptors 5 2.1 Classification of the adenosine receptors 12.2 Pharmacological characterization of the adenosine receptors 16 2.3 Molecular characterization of the adenosine receptors 22 2.4 Regulation of adenosine receptor gene and protein expression 25 2.5 Structural characterization of the adenosine receptors 29 2.6 Signal transduction mechanism 31 3. Biological actions of adenosine in the gastrointestinal tract 33 3.1 Gastroprotective actions of adenosine 35 3.2 Effect of adenosine on gastric acid secretion 37 II. Hypothesis 50 III. RationaleIV. Objectives and specific aims 51 Chapter 2: Methods 52 I. Effect of adenosine analogs on gastric SLI and IRG release 52 1. Isolated vascularly perfused rat stomach preparation 52. Drug administration and sample collection 53 3. Radioimmunoassay (RIA) for the measurement of SLI release 54 4. Radioimmunoassay for the measurement of IRG release 57 5. Data analysis 60 II. Adenosine receptor mRNA: distribution, structure, and quantification 61 1. Distribution of gastric adenosine receptor mRNA 61.1 Primer design and synthesis1.2 Optimization of RT-PCR conditions 63 1.3 Tissue extraction 61.4 Total RNA extraction and quantification 64 1.5 Formaldehyde gel electrophoresis 66 1.6 DNase I treatment1.7 First strand cDNA synthesis 7 1.8 PCR 61.9 Agarose gel electrophoresis 68 1.10 Restriction enzyme digestion2. Structure of the adenosine receptors 9 2.1 Detection of Ai and A2A receptor coding regions by RT-PCR 70 2.2 Cloning and DNA sequence analysis of the adenosine receptors 70 3. Quantification of gastric Aj and A2A receptor gene expression 73 3.1 Quantification by competitive RT-PCR 73.1.1 RNA competitor design and synthesis 3 3.1.2 Competitive RT-PCR assay 77 3.1.3 Competitive RT-PCR data analysis 78 3.2 Quantification by Real-Time RT-PCR 80 3.2.1 Design and synthesis of primers and probes 82 3.2.2 Synthesis of Ai and A2A receptor RNA standards 83.2.3 Two step Real-Time RT-PCR assay 86 3.2.4 Real-Time RT-PCR data collection and analysis 87 3.2.5 Optimization of Real-Time RT-PCR 88 III. Regulation of adenosine receptor gene expression and function in altered physiological states of the stomach 89 1. Quantification of A i and A2A receptor, somatostatin and gastrin gene expression 92 1.1 Animal treatment 91.2 Tissue extraction 3 1.3 Sample preparation1.4 Real-Time RT-PCR 94 1.5 Data collection and analysis 95 2. Effect of omeprazole on Ai and A2A receptor-induced changes in SLI and IRG release 97 IV. Cellular localization of the gastric Ai and A2A receptors 98 1. Tissue preparation for immunohistochemistry (IHC) 92. Adenosine receptor antibodies and control peptides 9 3. Antibodies used for double-staining 101 4. Confocal microscopy 102 5. Quantification of co-localization 103 6. Control studiesChapter 3: Results 105 I. Effect of adenosine on gastric IRG and SLI release 105 1. Effect of adenosine analogs on gastric IRG release 102. Effect of adenosine receptor antagonists on gastric IRG release 108 3. Effect of adenosine receptor agonists on gastric SLI release 108 4. Effect of adenosine receptor antagonists on gastric SLI release 115 5. Involvement of endogenous adenosine on SLI release 117 II. Distribution and sequence analysis of adenosine receptor mRNA 121 1. Distribution of adenosine receptor mRNA in various regions of the stomach 122. Ai and A2A receptor coding region and A2B and A3 receptor partial cDNA sequence 129 III. Quantification of gastric Adenosine Ai and A2A receptor mRNA expression 133 1. Competitive RT-PCR2. Real-Time RT-PCR 135 IV. Effect of fasting on adenosine receptor, somatostatin and gastrin gene expression 144 1. Effect of fasting on body weight and Total RNA concentration 144 2. Effect of fasting on 18S rRNA and GAPDH gene expression 144 3. Effect of fasting on adenosine Ai and A2A receptor expression 146 4. Effect of fasting on gastrin and somatostatin gene expression 149 V. Omeprazole treatment on adenosine receptor and gastric peptide gene expression 149 1. Effect of omeprazole treatment on 18S rRNA and GAPDH gene expression 152 2. Effect of omeprazole treatment on adenosine Ai and A2A receptor expression 152 3. Effect of omeprazole treatment on gastrin and somatostatin gene expression 155 VI. Omeprazole on adenosine agonist-induced changes in IRG and SLI release 155 1. Effect of omeprazole on CPA-induced changes in IRG release 157 2. Effect of omeprazole on agonist-induced changes in SLI release 157 VII. Cellular localization of the gastric adenosine A\ and A2A receptors 163 1. Cellular localization and distribution of AiR-IR 162. Double staining with the Ai receptor antibody 7 3. Cellular localization and distribution of A2AR-IR 171 4. Double staining with the A2A receptor antibody 8 Chapter 4: Discussion 18I. Alteration of IRG and SLI release by selective adenosine analogs 189 1. Effect of adenosine receptor-selective analogs on IRG release 189 2. Effect of adenosine receptor-selective analogs on gastric SLI release 191 3. Regulation of gastric SLI and IRG release by adenosine 196 II. Distribution and structure of gastric adenosine receptor mRNA 199 III. Quantification of Aj and A2A receptor gene expression 202 1. Competitive RT-PCR and Real-Time RT-PCR assay development 202 2. Changes of gastric Ai and A2A receptor gene expression and function by altered states of the stomach 208 IV. Cellular localization of the adenosine Ai and A2A receptor 220 1. Adenosine Ai and A2A receptors of the gastric mucosa 222 1.1 Adenosine Ai receptors on gastrin-secreting G-cells 224 1.2 Adenosine Ai and A2A receptors on somatostatin secreting D-cells 225 2. Adenosine Ai and A2A receptors of the enteric plexi 226 3. Adenosine Ai and A2A receptors of the gastric vasculature 230 V. Conclusion 232 References 4 Appendix 1: Table of abbreviations 280 List of tables Table 1: Adenosine receptor G protein coupling, signal transduction, and regulation of expression 32 Table 2: Effect of adenosine and adenosine agonists on gastric acid secretion in the rat 41 Table 3: Adenosine receptor RT-PCR primers and PCR conditions 62 Table 4: Ai and A2A receptor Real-Time RT-PCR assay: primers and probes 83 Table 5: Optimization of primer and probe concentrations for Real-Time RT-PCR 90 Table 6: Optimization of MgCl2 concentrations for Real-Time RT-PCR 91 Table 7: Gastrin and somatostatin Real-Time RT-PCR assay: primers and probes 96 Table 8: Effect of fasting and omeprazole treatment on body weight, tissue weight and total RNA concentration 145 List of figures Fig. 1. The de novo synthesis and metabolism of adenosine 4 Fig. 2. The molecular structure of adenosine 5 Fig. 3. The intracellular and extracellular synthesis and metabolism of adenosine 6 Fig. 4. Adenosine agonists and antagonists 18 Fig. 5. Schematic representation of the human adenosine Ai, A2A, and A3 receptors 30 Fig. 6. Typical elution profile for the separation of labeled gastrin 58 Fig. 7. Design and synthesis of the adenosine Ai and A2A receptor RNA competitors 75 Fig. 8. Analysis of competitive RT-PCR data 79 Fig. 9. Cleavage of the fluorogenic probe during Real-Time RT-PCR 81 Fig. 10. Orientation of Ai and A2A receptor transcript insertion into pGEM-T vector 85 Fig. 11. Effect of CPA on gastric IRG release 106 Fig. 12. Effect of various concentrations of CPA, and 0.1 uM of CGS 21680 and IB-MECA on IRG release 107 Fig. 13. Effect of DPCPX and DMPX on gastric IRG release 109 Fig. 14. Effect of DPCPX and DMPX on adenosine-induced changes in IRG release 110 Fig. 15. Effect of 0.1 uM and 1 uM CPA on gastric SLI release Ill Fig. 16. Effect of 0.1 uM CGS 21680 and IB-MECA on gastric SLI release 112 Fig. 17. Effect of various concentrations of adenosine agonists on gastric SLI release 114 Fig. 18. Effect of DMPX on basal, adenosine, and adenosine agonist-stimulated SLI release 116 Fig. 19. Effect of ZM 241385 on basal and stimulated SLI release 118 Fig. 20. Effect of ZM 241385 and DPCPX on basal and adenosine and agonist-stimulated SLI release 119 Fig. 21. Effect of EHNA on basal and adenosine-induced SLI release 120 Fig. 22. Results of A i, A2A, A2B and A3 RT-PCR performed using rat gastric tissues 122 Fig. 23. Amplification of the Ai and A2A receptor coding region by RT-PCR 123 Fig. 24. Restriction enzyme digestion of the adenosine A] receptor PCR amplicon 124 Fig. 25. Restriction enzyme digestion of the adenosine A2A receptor PCR amplicon 125 Fig. 26. Restriction enzyme digestion of the adenosine A2B receptor PCR amplicon 126 Fig. 27. Restriction enzyme digestion of the adenosine A3 receptor PCR amplicon 127 Fig. 28. Restriction enzyme digestion of the full length adenosine A2A receptor amplicon 128 Fig. 29. Alignment of the rat mucosal and brain Ai receptor coding regions 130 Fig. 30. Alignment of the rat mucosal and brain A2A receptor coding regions 131 Fig. 31. Alignment of the rat mucosal and brain A2B and A3 cDNA sequences 132 Fig. 32. Striatal Ai and A2A receptor gene expression measured by competitive RT-PCR. 134 Fig. 33. Effect of template concentrations on quantification of Ai and A2A receptor mRNA by competitive RT-PCR 136 Fig. 34. Quantification of gastric Ai receptor gene expression by competitive RT-PCR 137 Fig. 35. Quantification of gastric A2A receptor gene expression by competitive RT-PCR 138 Fig. 36. Competitive RT-PCR quantification of Ai and A2A receptor gene expression in the rat stomach 139 Fig. 37. Results of a representative Ai Real-Time RT-PCR assay 140 Fig. 38. Results of a representative A2A Real-Time RT-PCR assay 142 Fig. 39. Real-Time RT-PCR quantification of gastric and striatal adenosine A| and A2A receptor gene expression 143 Fig. 40. Effect of fasting on gastric housekeeping gene expression 147 Fig. 41. Effect of fasting on adenosine receptor gene expression 148 Fig. 42. Effect of fasting on gastric peptide gene expression 150 Fig. 43. Comparison between fasting-induced changes in somatostatin and adenosine A2A receptor gene expression 151 Fig. 44. Effect of omeprazole treatment on gastric housekeeping gene expression 153 Fig. 45. Effect of omeprazole treatment on adenosine Ai and A2A receptor gene expression 154 Fig. 46. Effect of omeprazole treatment on gastric peptide gene expression 156 Fig. 47. Effect of CPA on gastric IRG release from perfused stomachs of 3 day omeprazole-treated and control animals 158 Fig. 48. Effect of omeprazole treatment on CPA-induced changes in IRG release 159 Fig. 49. Effect of CGS 21680 on gastric SLI release in 3 day omeprazole-treated and control animals 160 Fig. 50. Effect of omeprazole treatment on CGS 21680-induced changes in SLI release 161 Fig. 51. Effect of omeprazole treatment on CPA-induced changes in SLI release 162 Fig. 52. Representative AiR-IR and A2AR-IR in various regions of the rat stomach 164 Fig. 53. Confocal images of AjR-IR in the corpus of the rat stomach 165 Fig. 54. Confocal images of AiR-IR in the antrum of the rat stomach 166 Fig. 55. Double staining of AiR-IR with gastrin-IR 168 Fig. 56. Double staining of AjR-IR with somatostatin-IR in the gastric mucosa 169 Fig. 57. Double staining of AiR-IR and somatostatin-IR in the gastric muscle 170 Fig. 58. Double-staining of AiR-IR with H+K+-ATPase P-IR 172 Fig. 59. Double staining of AiR-IR with VWF-IR 173 Fig. 60. Localization of AiR-IR relative to VWF-IR in the corporeal muscle 174 Fig. 61. Double staining of AiR-IR with PGP 9.5-IR 175 Fig. 62. Confocal images of A2AR-IR in the corpus of the rat stomach 176 Fig. 63. Confocal images of A2AR-IR in the antrum of the rat stomach 177 Fig. 64. Double staining of A2AR-IR with somatostatin-IR in the gastric mucosa 180 Fig. 65. Double staining of A2AR-IR with somatostatin-IR in the gastric muscle of the corpus 181 Fig. 66. Double staining of A2AR-IR with somatostatin-IR in the gastric muscle of the antrum 182 Fig. 67. Double staining of A2AR-IR with gastrin-IR 183 Fig. 68. Double-staining of A2AR-IR with H+K+-ATPase p-IR 184 Fig. 69. Double staining of A2AR-IR with VWF-IR 185 Fig. 70. Localization of A2AR-IR relative to VWF-IR in the corporeal muscle 186 Fig. 71. Double staining of A2AR-IR with PGP 9.5-IR 187 Acknowledgements I am most grateful to my supervisor, Dr. Kenny Y. N. Kwok for his many years of guidance and support, and for his extraordinary kindness and generosity. The academic skills I've acquired from him will certainly serve me well throughout my career. My sincerest gratitude to Dr. Ross MacGillivray for his support and encouragement throughout my Ph.D. Many thanks also to Jeff Hewitt for teaching me the techniques of molecular biology that have figured so prominently in my work, and for useful technical discussion. I would also like to acknowledge Dr. Alison Buchan for her patience and guidance during our collaboration, through which I acquired my knowledge of immunohistochemistry. Thanks also to Dr. Ray Pederson for carefully reading over this thesis and making graduate studies in this department a more social experience. I am also grateful to other members of my supervisory committee, Dr. Steve Vincent and Dr. Chris Mcintosh, for direction in preparation for my comprehensive exam. In addition, I would like to express my appreciation for the students who have worked in my laboratory, particularly Henry Leung, who contributed to the immunohistochemistry and perfusion studies. Thanks also to Zaira Khan for her administrative support, and finally to Stephan Gudmundson, my family, and friends, who supported me throughout this experience. Financial support from The University of British Columbia, in the form of several University Graduate Fellowship awards, from the Cordula and Gunter Paetzold Fellowship, and from the Wah-Sheung Prize was greatly appreciated. Chapter 1: Introduction Adenosine is found ubiquitously throughout the body, and has been shown to act as a neurotransmitter, neuromodulator, and local mediator in various biological systems (Ralevic and Burnstock 1998). Adenosine exerts its effect by activating purinergic receptors belonging to the PI receptor family. These receptors are classified into the adenosine Ai, A2A, A2B, and A3 receptor subtypes (Fredholm et al., 2001). In the stomach, adenosine has been shown to protect against stress-, indomethacin-, and ethanol-induced ulcerative formations (Glavin et al., 1987; Cho and Ogle, 1990; Cho et al., 1991, 1995; Bozkurt et al., 1998). This gastroprotective action is likely due to the inhibitory effect of adenosine on gastric acid secretion. Although adenosine has been shown to exert its acid-inhibitory effect in dogs and guinea pigs by acting directly on parietal cells (Gerber et al., 1985; Heldsinger et al., 1986), this does not occur in the rat (Puurunen et al., 1987). Instead, adenosine inhibits the release of gastrin, a major secretagogue of gastric acid, and stimulates the release of somatostatin, a potent inhibitor of acid secretion (Kwok et al., 1990). Adenosine elicits these actions by activating adenosine receptors, but the receptor subtypes involved have not been determined. This thesis examines the adenosine receptor subtypes involved in regulating gastrin and somatostatin secretion in the rat stomach, and characterizes the cellular localization, abundance, and structure of these receptors. Various physiological states of the stomach have been shown to alter gastric acid secretion by regulating the release and synthesis of gastrin and somatostatin. Another objective of this thesis is to examine whether adenosine may be involved in this regulatory process. Thus, changes in adenosine receptor gene expression were examined under altered physiological states of the stomach that are known to alter gastrin and somatostatin synthesis and release. This chapter provides background information pertinent to the current study, and is organized into three sections. The first section discusses the regulation of endogenous adenosine levels by processes controlling the synthesis, release, reuptake and metabolism of adenosine. The second section describes the classification and pharmacological and molecular characterization of each adenosine receptor subtype. This section also reviews the regulation of adenosine receptor gene and protein expression. Since this thesis focuses on the Aj and A2A receptors, emphasis is placed on background literature related to these two subtypes. The third section discusses the gastric actions of adenosine, with particular attention given to the role of adenosine in regulating gastric secretion. This section also describes key experiments that this study is based upon. This chapter concludes by presenting the experimental hypothesis, rationale and specific aims of the thesis research. I. Background 1. Regulation of endogenous adenosine levels Since adenosine is the endogenous ligand for all four adenosine receptor subtypes, it is essential to understand how the level of this nucleoside is regulated. Several processes are involved in the production, release, reuptake and metabolism of adenosine. In the stomach, adenosine is produced by intracellular and extracellular salvage pathways that produce adenosine from sources already containing an intact purine ring. De novo synthesis occurs mainly in cells of the liver, and is responsible for the complete formation of adenosine from sources that do not contain a purine ring. Diet is not a source of significant amounts of adenosine since most purines absorbed by the intestines are catabolized by xanthine oxidase in the esophageal, gastric, small intestinal and large intestinal epithelium (Moriwaki et al., 1996). When adenosine is present in the extracellular fluid, it is rapidly removed by cellular reuptake through specialized nucleoside transporters (discussed in section 1.4), degraded to inosine by adenosine deaminase (ADA) or converted to AMP by adenosine kinase (AK). The next section reviews the regulation of endogenous adenosine levels by the synthesis, release, reuptake and metabolism of adenosine. 1.1 The de novo synthesis pathway Although de novo synthesis of adenosine has not been shown to occur in the stomach, this pathway is included in this overview to illustrate the biosynfhetic origin of the purine ring. A simplified pathway for the de novo synthesis and metabolism of adenosine is described in Fig. 1. Components of the purine ring are donated by glutamine, glycine, formyl-tetrahydrofolic acid, and aspartate during this process (see Fig. 2). De novo synthesis occurs mainly in the liver. In other tissues, including the stomach, adenosine is produced through intracellular and extracellular salvage pathways. 1.2 Intracellular and extracellular salvage Under normoxic conditions, intracellular adenosine is formed mainly by the conversion of S-adenosyl homocysteine (SAH) to adenosine (Lloyd et al., 1988) (See Fig. 3). SAH is formed when S-adenosyl methionine (SAM) donates a methyl group to methylate DNA, mRNA, RNA, and rRNA. SAH is hydrolyzed by the enzyme S-adenosyl 10-step de novo purine ring formation Glutamine, Glycine, •4 i Aspartate t RibOSe 5-PhOSphate + ATP • PRPP formyl-tetrahydrofolic acid ||\/|p 1 2 Aspartate 5 4 i Inosine -4 Adenosine A AMP -4—- AMPS i hypoxanthine —-—• xanthine —-—• H202+Urate Fig. 1. The de novo synthesis and metabolism of adenosine. The de novo synthesis of adenosine occurs intracellularly through the actions of the following enzymes: 1) PRPP synthase; 2) adenylosuccinate synthetase ; 3) adenylosuccinate lyase; and 4) endo-5' nucleotidase. Intracellular metabolism of adenosine utilize the following enzymes: 5) adenosine deaminase; 6) phosphorlase; and 7) xanthine oxidase. In the initial step, two phosphates from ATP are transferred to ribose 5-phosphate to form PRPP. The purine ring is formed by a 10-step reaction which converts PRPP to IMP. IMP is converted to AMPS by the addition of aspartate. AMP is hydrolyzed by 5'endonucleotidase to form adenosine (Stone and Simmonds, 1991). [Abbreviations: 5'phospho-a-D-riobsyl 1 pyrophosphate (PRPP), inosine monophosphate (IMP), adenylosuccinate (AMPS), and adenosine monophosphate (AMP)]. Adenosine Source Glutamine [ Glycine 1 Formyl-tetrahydrofolic acid J Aspartate j Ribose 5-phosphate Fig. 2. The molecular structure of adenosine. The ribose moiety is donated by ribose 5-phosphate. The purine ring is formed by de novo synthesis and consists of components donated by glutamine, glycine, formyl-tetrahydrofolic acid and aspartate. extracellular ATP E-NTPase E-NPP AP CAMP ADP E-NTPase E-NPP AP AMP Ecto-5' NT Inosine I ADA -• Adenosine Adenosine Transporter T ATP ATPase ^ . MlK • ADP ATPas V AMP ^ Encfo 5-WT 4K intracellular SAM -> SAH > Adenosine SAHH Inosine Fig. 3. The intracellular and extracellular synthesis and metabolism of adenosine. Adenosine is produced intracellularly and extracellularly through salvage pathways using sources which already contain an intact purine ring. Enzymes responsible for the extracellular synthesis (blue), intracellular synthesis (red), and metabolism (green) of adenosine are shown. Cellular release and reuptake of adenosine occur through the adenosine transporters. [Abbreviations: ecto-nucleoside triphosphate diphoshohydrolase (E-NTPase), ecto-nucleotide pyrophosphatase/phosphodiesterase (E-NPP), alkaline phosphatase (AP), ecto-5' nucleotidase (ecto-5' NT), adenosine deaminase (ADA), adenosine kinase (AK), cyclic AMP (cAMP), nicotinamide adenine dinucleotide (NAD), S-adenosyl methionine (SAM), S-adenosyl homocysteine (SAH), S-adenosyl homocysteine hydrolase (SAHH)] homocysteine hydrolase (SAHH) to form adenosine (Lloyd et al., 1988). Under hypoxic or ischemic conditions, the contribution of SAH to adenosine levels is negligible (Deussen et al., 1989). Instead, intracellular adenosine is formed primarily by the dephosphorylation of adenosine triphosphate (ATP) (Geiger et al., 1997). ATP is converted to adenosine diphosphate (ADP) and then adenosine monophosphate (AMP) by ATPase, and AMP is then dephosphorylated by either cytosolic AMP-specific or IMP-specific 5' nucleotidase to produce adenosine (Sala-Newby et al., 1999). In the stomach, extracellular adenosine may be synthesized though the hydrolysis of AMP by 5' nucleotidase. The membrane-bound form of 5' nucleotidase has been localized in the corpus and antrum of the stomach (Sakai et al., 1981; Grover et al., 1983, 1985; Borkje et al., 1986). The stomach is divided into three morphological and functionally distinct regions, the fundus, corpus and antrum, as shown in Fig. 36. The activity of 5' nucleotidase was found on plasma membrane enriched fractions extracted from canine antral and corporeal smooth muscle cells (Sakai et al., 1981; Grover et al., 1983), and rat corporeal smooth muscle cells (Grover et al., 1985). The activity of 5' nucleotidase has also been measured in biopsy samples extracted from human antral and corporeal gastric mucosa (Borkje et al., 1986). Thus, the activity of this enzyme may regulate gastric adenosine levels. The state of the stomach has been shown to alter 5' nucleotidase activity. The activity of this enzyme is reduced in the gastric juice of patients suffering from gastric cancer, gastric ulcers, and gastritis (Durak et al., 1994). This may be due to changes in gastric acidity since 5' nucleotidase activity has been shown to be altered by changes in acidity in the rat skeletal muscle. In particular, increased acidity was shown to increase extracellular adenosine levels by stimulation of 5' nucleotidase activity (Cheng et al., 2000). Extracellular adenosine is synthesized from AMP as shown in Fig. 3. AMP levels are regulated by a number of enzymes which hydrolyze ATP, ADP, nicotinamide adenine dinucleotide (NAD+) and cyclic AMP (cAMP). These enzymes are classified into the ecto-nucleoside triphosphate diphoshohydrolase (E-NTPase) family, the ecto-nucleotide pyrophosphatase/phosphodiesterase (E-NPP) family, and the alkaline phosphatase family. The characteristics of these enzymes have been reviewed by Zimmerman (Zimmermann, 2000). The enzymes are not as important as 5' nucleotidase in regulating extracellular adenosine levels since the dephosphorylation of AMP to adenosine is the rate-limiting step in the production of extracellular adenosine (Dunwiddie et al., 1997). Therefore, 5' nucleotidase activity may be an important regulator of extracellular adenosine production in the stomach. 1.3 Presynaptically and postsynaptically released ATP as a source of adenosine Extracellular adenosine can also be derived from the degradation of ATP released from purinergic nerve terminals (Burnstock et al., 1970; Burnstock, 1975). Purinergic nerves were proposed by Burnstock based on studies performed in the guinea pig and toad stomach (Burnstock et al., 1970). In the stomach, the cell bodies of purinergic nerves are found in the myenteric plexus, and the axons of these nerves project to the circular and longitudinal muscles (Burnstock, 1975). ATP is released from these nerves and acts as a neurotransmitter (Su et al., 1971; Burnstock, 1975). ATP is stored in synaptic vesicles with other neurotransmitters, and is released by nerve depolarization (Su et al., 1971). In the brain, ATP is released presynaptically from a number of neurons, including cholinergic, glutamatergic and catecholaminergic neurons, and from glial cells (Ralevic and Burnstock, 1998). In the gut, ATP was also suggested to be co-released with nitric oxide (Selemidis et al., 1997), pituitary adenylate cyclase-activating polypeptide (Furness et al., 1995; Imoto et al, 1998), and vasoactive intestinal peptide (VIP) (Furness et al., 1995). Vizi and colleagues have also suggested that acetylcholine and norepinephrine may elicit ATP release from postsynaptic sites through a process that they referred to as cascade transmission (Vizi et al., 1992). This type of release has been observed in the gut. Specifically, acetylcholine was shown to induce ATP release from smooth muscle cells of the guinea pig ileum via activation of postsynaptic M3 receptors (Nitahara et al., 1995). Thus, in - the stomach, adenosine may be derived from the hydrolysis of presynaptically or postsynaptically released ATP. Although cAMP may also be released from neurons and can be metabolized to form adenosine (see Fig. 3), cAMP was not shown to significantly alter extracellular adenosine levels unless there is continuous release over an extended period of time (Brundege et al., 1997). 1.4 Cellular release and reuptake of adenosine by adenosine transporters Adenosine transporters are responsible for the cellular release and reuptake of intracellular and extracellular adenosine, respectively (Geiger et al., 1997). Adenosine transporters are classified into two groups, the equilibrative or facilitated-diffusion carriers and the concentrative or sodium-dependent nucleoside transporters (Thorn and Jarvis, 1996). The equilibrative nucleoside transporters are found in most cell types and are divided into two groups, the es (equilibrative, sensitive) or ei (equilibrative, insensitive) carriers, based on whether they are sensitive to blockade by nitrobenzylthioinosine, a synthetic nucleoside analogue (Lee and Jarvis, 1988). Both classes translocate adenosine bidirectionally across the plasma membrane, and are blocked by inhibitors, such as dipyridamole, dilazep and lidoflazine (Thorn and Jarvis, 1996). Our laboratory has suggested that these transporters may be involved in regulating the endogenous level of extracellular adenosine in the rat stomach (Kwok et al., 1990). The administration of dipyridamole was shown to significantly enhance basal and adenosine-induced augmentation of gastric somatostatin-like immunoreactivity (SLI) release in the isolated perfused stomach preparation (Kwok et al., 1990). Other studies have also suggested that the ei transporters function as an adenosine-scavenging system to maintain the extracellular adenosine level at the basolateral membrane of the intestinal epithelia (Tally et al., 1996). Concentrative nucleoside transporters have not been studied extensively in the stomach, but have been detected on epithelial cells of the intestines (Ritzel et al., 2001). Currently, four concentrative nucleoside transporters (N1-N4) have been identified and characterized (Thorn and Jarvis, 1996). All four can transport adenosine bidirectionally across the membrane, but are generally responsible for the influx of adenosine. Currently, no selective inhibitors exist for the concentrative nucleoside transporters (Geiger et al., 1997). Like 5' nucleotidase, the expression of equilibrative adenosine transporters may be altered to regulate endogenous adenosine concentrations (Lorbar et al., 1999). Although this has not been examined in the stomach, alterations in adenosine transporter activity have been observed in the hearts of aged rats. A decrease in adenosine transporter expression was observed with increased levels of extracellular adenosine in these animals (Lorbar et al., 1999). 1.5 Metabolism of adenosine Extracellular adenosine can be regulated by the action of ADA, an enzyme that converts adenosine to inosine (See Fig. 3). ADA is expressed in the cytoplasm and on cell membranes of all tissues (Franco et al., 1997), and can be inhibited by erythro-9-(2-hydroxy-3-nonyl)adenine hydrochloride (EHNA) (Mendelson et al., 1983). ADA is highly expressed in the thymus and spleen in humans (Moriwaki et al., 1999), and the lung, spleen, and intestines in rats (Centelles et al., 1987). The expression of the ADA gene has also been detected in the epithelia of the mouse esophagus, forestomach and duodenum (Witte et al., 1991). The high expression of ADA throughout the gastrointestinal tract suggests that much of the adenosine that is absorbed through the gut is metabolized. In ADA-deficient mice, a significantly elevated level of adenosine was observed in various regions of the gastrointestinal tract, including the forestomach, hindstomach, esophagus, duodenum, jejunum and ileum (Xu and Kellems, 2000). Thus, ADA plays a significant regulatory role in modulating endogenous adenosine levels in the gastrointestinal tract. In humans, changes in gastric ADA activity have been shown to occur with changes in gastric acidity (Namiot et al., 1990, 1991, 1993). In particular, hypersecretors of acid were shown to exhibit increased ADA activity, while patients suffering from achlorhydria, a condition characterized by a lack of gastric acid secretion, were shown to exhibit lowered ADA activity (Namiot et al., 1990). Thus, ADA may play a role in regulating gastric acid secretion by modulating endogenous adenosine levels. Mucosal ADA activity has also been shown to decrease in the vicinity of a healed ulcer (Namiot et al., 1993) and in the gastric mucosa after treatment with ranitidine, a gastric acid inhibitor used for treatment of ulcers (Namiot et al., 1991). These findings suggest that increased adenosine levels may be involved in ulcer healing. As shown on Fig. 3, the cytosolic enzyme adenosine kinase (AK) is responsible for catalyzing the phosphorylation of adenosine to AMP (Geiger et al., 1997), and is found in various gastrointestinal tissues, including the stomach, duodenum, ileum, and large intestine (Andres and Fox, 1979). This enzyme has been shown to regulate endogenous adenosine levels. For example, inhibition of AK activity in the hippocampus has been shown to increase intracellular adenosine levels (Pak et al., 1994). During oxidative and metabolic stress, decreased AK activity has also been shown to significantly increase adenosine levels in DDT1 MF-2 smooth muscle cells (Sinclair et al., 2000). 1.6 Endogenous adenosine levels As described in the previous sections, the endogenous adenosine level is tightly regulated by processes controlling its synthesis, release, reuptake and metabolism. Although adenosine can be synthesized intracellularly and can be produced extracellularly, under well oxygenated conditions, the majority of adenosine (92%) is produced intracellularly and is released into the extracellular space, as illustrated in the isolated guinea pig heart (Deussen, 2000). In plasma, adenosine has a short half-life of only a few seconds because it is rapidly degraded by ADA or removed by cellular reuptake (Geiger et al., 1997). Under normoxic conditions, basal adenosine levels are consistently found in the nanomolar range (see below). Although the adenosine concentration has not been measured in the stomach, the venous adenosine concentration in the rat jejunum has been shown to be 62 nM (Sawmiller and Chou, 1990). This concentration is consistent with levels measured elsewhere, such as in the rat cerebrospinal fluid (160 nM) (Meno et al., 1991), cerebral cortex (30-50 nM) (Phillis et al., 1987), and striatum (41- 210 nM) (Ballarin et al., 1991; Pazzagli et al., 1995; Melani et al., 1999). In whole blood and plasma of rats, adenosine concentrations were found to be 79 nM (Phillis et al., 1992) and 320 nM (Conlay et al., 1997), respectively. Arterial adenosine concentrations have also been measured in the rat and found to be 59 nM (Sawmiller and Chou, 1990). In human blood, the concentration was shown to be 35 nM (Geiger et al., 1997). During periods of stress, adenosine levels can rise to micromolar concentrations. In rats, ischemia induced by medial cerebral artery occlusion was shown to increase adenosine concentrations in the striatum and hippocampus to 3 \iM (Melani et al., 1999) and 30 uM (Latini et al., 1999), respectively. The increase in adenosine concentration is due to enhanced adenosine production caused by the degradation of ATP. The adenosine production rate in the rat heart has been shown to increase by approximately 70-fold during ischemia, from a normoxic production rate of 6.4-7.7 nmol min"1 g"1 (Lorbar et al., 1999) to an ischemic production rate of 410 nmol min"1 g"1 (Meghji et al., 1988). The plasma adenosine level has also been shown to increase to micromolar concentration after chronic blockade of adenosine receptors (Conlay et al., 1997). In rats treated with caffeine for two weeks, the plasma adenosine concentration was found to increase from 320 nM to 3.2 uM (Conlay et al., 1997). Thus, endogenous adenosine levels are clearly elevated by oxidative stress and by treatment with adenosine antagonists. The concentration of adenosine exposed to target tissues may be important because adenosine has been shown to elicit concentration-dependent physiological effects. For example, our laboratory has shown that gastric somatostatin release is inhibited by low concentrations of adenosine (0.01 [xM), but is stimulated by higher concentrations (>0.6 uM) (Kwok et al., 1990). The physiological actions of adenosine are determined, in part, by the adenosine receptor subtype stimulated. These subtypes are described in detail in the next section. However, it is worth mentioning here that the activation of each adenosine receptor subtype depends on the local extracellular adenosine concentration. The high affinity adenosine Ai and A2A receptor subtypes may be stimulated by relatively low concentrations of adenosine, while the low affinity A2B and A3 receptors may be activated by extremely high concentrations of adenosine, such as those present during oxidative stress (Ralevic and Burnstock, 1998). This concentration-dependent activation of adenosine receptors is suggested by studies showing that adenosine elicits Ap, A2A-, A2B-, and A3- mediated effects with EC50 values of 73, 150, 5100, and 6500 nM, respectively (Daly and Padgett, 1992; Zhou et al., 1992; Peakman and Hill, 1994). In certain tissues, adenosine has been shown to elicit distinct effects depending on the concentration in which it is administered, and the adenosine receptor subtype that is activated. For example, in the renal vasculature, the administration of low micromolar concentrations of adenosine was shown to vasoconstrict renal afferent arterioles via activation of Ai receptors, while higher concentrations of adenosine (>10 uM) resulted in vasodilation of these vessels via activation of A2A receptors (Tang et al., 1999; Tabrizchi and Bedi, 2001). In the lung, low adenosine concentrations mediate anti inflammatory actions by activating the A2A receptors, while high adenosine concentrations mediate pro-inflammatory actions by activation of the A3 receptors (Blackburn, 2003). These observations raise the possibility that the concentration-dependent effect of adenosine on somatostatin release in the isolated rat stomach is mediated by the activation of different adenosine receptor subtypes. 2. The adenosine receptors 2.1 Classification of the adenosine receptors The concept of purinergic receptors was introduced in the late 1970s by Burnstock (Burnstock, 1978). These receptors were classified into the Pi and P2 purinoceptors, based mainly on their binding preference for adenosine or ATP, respectively. The PI receptors (or adenosine receptors) were initially characterized by the rank order of binding preference (adenosine > AMP > ADP > ATP), by their sensitivity to blockade by methylxanthines, and by their ability to alter cAMP levels through the modulation of adenylate cyclase activity (Burnstock, 1978). The Ai and A2 receptor subtypes were proposed in 1979 by van Calker to describe the two distinct plasma membrane receptors on cultured brain cells that were capable of inhibiting and stimulating adenylate cyclase activity, respectively (van Calker et al., 1979). Two A2 receptor subtypes were later identified: a high-affinity A2 receptor (A2A) on rat striatal membranes, and a low affinity A2 receptor (A2B) on rat cerebral cortical membranes (Daly, 1983). After binding studies demonstrated that these receptors represented distinct receptors subtypes (Bruns et al., 1986), the A2A and A2B nomenclature was proposed by Bruns to refer to these high and low affinity A2 receptors, respectively. While the Ai, A2A and A2B receptors were initially identified and named based on pharmacological studies, the fourth adenosine receptor subtype, the A3 receptor, was cloned and its molecular structure was determined before it was identified as a distinct subtype using pharmacological means (Zhou et al., 1992). Currently, the nomenclature and classification of the adenosine receptors adhere to the principles of receptor nomenclature adopted by the International Union of Pharmacology (IUPHAR). Each distinct receptor subtype is identified based on both molecular and pharmacological evidence. The existence of the four adenosine receptor subtypes is now well established (Ralevic and Burnstock, 1998; Fredholm et al., 2001), and the gene expression of all four subtypes has been demonstrated in various tissues, including the rat stomach (Dixon et al., 1996). Although additional adenosine receptor subtypes have been proposed, the structural and pharmacological evidence required by the IUPHAR to support their existence is lacking (Cornfield et al., 1992). 2.2 Pharmacological characterization of the adenosine receptors The adenosine receptor subtypes can be characterized pharmacologically by the rank order of potency of various adenosine analogs in eliciting certain responses. In the past two decades, numerous adenosine agonists and antagonists have been developed. Nearly all known agonists are based on the structure of adenosine, while antagonists are based on both xanthine and non-xanthine compounds (Fredholm et al., 2001). Due to the large number of ligands available for these receptors, the following section focuses on ligands that are commonly used to characterize the four adenosine receptors subtypes, and on the ligands which are used in the present study. Adenosine agonists are based on the structure of adenosine, and require an intact ribose moiety (Fredholm et al., 2001; Jacobson et al., 2001). Certain structural modifications have been shown to increase the metabolic stability of adenosine, resulting in longer half lives and/or resistance to degradation and cellular reuptake (Pavan 1998). Adenosine can be modified at three sites to increase metabolic stability and confer receptor subtype selectivity (Jacobson, 1996; Klotz, 2000): the 5' position of the ribose ring, the C2 position and the N6 position of the purine ring (see Fig. 4). Substitutions at the N6 position generally increase Ai and A3 receptor selectivity, while substitutions at the C2 position generally increase A2A receptor selectivity (Bruns et al., 1986; Klotz, 2000). Agonists containing modifications at more than one site are generally more potent (Jacobson, 1996). The binding affinity and the structure of some of the most common adenosine receptor agonists are shown in Fig. 4. Analogs containing bulky substitutions at the N6 position, such as N6-cyclopentyladenosine (CPA), 5'-N6-cyclohexyladenosine (CHA), R(-)-N6-(2-phenylisopropyl)adenosine (R-PIA), and (S)-N6-(2-phenylisopropyl)adenosine (S-PIA), preferentially bind to the Ai receptor (Bruns et al., 1986; Klotz et al, 1998). CPA is one of the most potent and selective Ai receptor agonists (see Fig. 4), and has been used extensively to characterize Ai receptor-mediated responses in the gut. In the gastric antrum of the guinea pig, CPA was used to demonstrate the involvement of the Ai receptor in the inhibition of fast excitatory postsynaptic potentials (Christofi et al., 1992). CPA has also been used to demonstrate the presence of Ai receptors in the rat duodenum (Nicholls et al., 1992; Brownhill et al., 1996; Peachey et al., 1996), jejunum (Hancock and Coupar, 1995), and colon (Peachey et al., 1999). A2A receptors prefer agonists with bulky substitutions at the C2 position, such as 2-phenylaminoadenosine (CV1808) and the potent A2A receptor agonist 2-p-(2-carboxyethyl)phenethylamino-5'N-ethylcarboxamidoadenosine (CGS 21680) (Hutchison et al., 1989; Lupica et al., 1990). CGS 21680 is highly selective for A2A receptors, and exhibits a 140-fold selectivity for A2A receptors over Ai receptors (Hutchison et al., 1989). CGS 21680 has low affinity for the A2B and A3 receptors (Klotz et al., 1998) (see Fig. 4), and was used to characterize the first A2A receptors cloned from the rat striatum (Fink et al., 1992). In the rat isolated perfused superior mesenteric arterial bed, CGS 21680 was used to demonstrate A2A receptor-mediated vasodilation (Hiley et al., 1995). In our laboratory, CGS NH, u OH OH Adenosine N* position (purine) C2 position (purine) 5' position (ribose) PIA R-PIAfL-PIAI A,: 2 V 860 A„: 3800-33700 A,:16 S-PIA (D-PIA1 A,: 75 V7800 A„: 62,800 A,: 45 00 CPA a. 00 A,: 2.3 OH OH Mm ^21,000-34,400 A,: 43 CGS 21680 A,: 290 M* A,: 38,100 A,: 87 NECA A,: 14 A^: 330-360 A,: 6 NH, 00 NH2 IB-MECA A,: 3.7 V 2,500 A„: 54,000 A,: 1.2 9' NH 0 f*7 I R3 Xanthine Caffeine A,: 29,100 A^; 48,100 Theophylline i 1 A,: 6.800 V 1700 Aj,: 7.900 A, 86,000 8-PT ^x3 A,: 86 A»: 848 DPCPX r-0 r1 A,: 3 M M A 129 51 3960 DMPX x n A,: 12,000 A,,: 9.000 ZM 241385 (non-xanthine) IC A,: 260 A»: 0.8 A_: 32 A,: 10.000 Fig. 4. Adenosine agonists (A) and antagonists (B). Adenosine agonists are based on the structure of adenosine. These compounds contain an intact ribose moiety, and may be modified at the N6 and C2 position of the purine ring or at the 5' position of the ribose moiety. The Ki values (nM), which indicate the relative binding affinities at different receptor subtypes, are shown (Bruns et al., 1986; Klotz et al., 1998;Ralevic and Burnstock, 1998; Fredholm et al., 2001). Many adenosine antagonists are xanthine-based, containing possible modifications at the 1,3, 7, or 8 position of the xanthine ring. Caffeine and theophylline are naturally occurring xanthines. 8-PT, DPCPX, and DMPX are xanthine-based derivatives, and ZM 241385 is a non-xanthine A2A receptor-selective antagonist 21680 was also shown to decrease perfusion pressure in the isolated vascularly perfused rat stomach in a concentration-dependent manner (Yip and Kwok, 2004). Selective agonists have not been developed for the A2B receptor, despite intense efforts (Cristalli et al., 1998; de Zwart et al., 1998). Currently, one of the most potent agonists for this receptor is the non-selective 5' modified agonist N-ethylcarboxamidoadenosine (NECA; see Fig. 4) (Ralevic and Burnstock, 1998). NECA and CGS 21680 bind to A2A receptors with similar affinity, but NECA binds to the A2B receptors with higher affinity than CGS 21680 (Brackett and Daly, 1994; Klotz et al., 1998). Thus, in various tissues, including the gut, A2A and A2B receptor-mediated effects are often distinguished by the relative potency of NECA and CGS 21680. In the rat duodenum (Brownhill et al., 1996; Nicholls et al, 1996) and distal colon (Peachey et al., 1999), A2B receptor-mediated actions were demonstrated by the potent effect of NECA and by the lack of an effect by CGS 21680. Actions mediated by A2A and A2B receptors can also be distinguished by the EC50 values of NECA and CGS 21680 (Brackett and Daly, 1994). In particular, in PC12 cells that express A2A receptors, activation of adenylyl cyclase elicited by CGS 21680 and NECA were characterized by EC50 values of 0.07 and 0.13 p.M, respectively (Brackett and Daly, 1994). In fibroblast 3T3 cells expressing A2B receptors, the activation of adenylyl cyclase activity elicited by NECA was characterized by an EC50 of 1.9 \xM, and CGS 21680 was unable to elicit a maximal response (Brackett and Daly, 1994). Although CPA, CGS 21680, and NECA can also bind to the A3 receptors, N6-substituted adenosine-5'-uronamides, such as l-deoxy-l-[6-[[(3-iodophenyl)methyl]amino]-9H-purin-9-yl]-N-methyl-B-D-ribofuranuronamide (IB-MECA), are more selective for this receptor subtype (Gallo-Rodriguez et al., 1994) (see Fig. 4). In the gut, IB-MECA has been used to examine the involvement of A3 receptors in the relaxation of the guinea pig taenia caeci (Prentice and Hourani, 1997). This agonist has also been used to demonstrate the involvement of A3 receptors in protecting against colitis in the mouse (Mabley et al., 2003). The adenosine receptor involved in eliciting a specific effect can be determined pharmacologically by comparing the relative potency of various adenosine agonists. Based on the selectivity and binding affinity of various agonists at each adenosine receptor subtype (see Fig. 4), the following rank order of potency can be used to characterize Ai, A2A, A2B and A3 receptor mediated actions: Ai: CPA > CHA > R-PIA > NECA > IB-MECA > CGS 21680; A2A: CGS 21680 = NECA = CV1808> CPA > IB-MECA; A2B: NECA > CGS 21680 = CV1808 > CPA > IB-MECA; A3: IB-MECA > NECA > CPA > CGS 21680. Since adenosine agonists are not specific, and are only selective for a given adenosine receptor subtype, the use of selective antagonists is also necessary to characterize the adenosine receptor subtype involved in eliciting various actions. Adenosine receptors, with the exception of the A3 receptor in some species, are blocked by methylxanthines such as caffeine and theophylline (Zhou et al., 1992; Klotz, 2000). A number of selective xanthine-based antagonists have been developed by modification of the 1,3, 7, and/or 8 positions of the xanthine ring. Unlike human A3 receptors, rodent A3 receptors are not blocked by xanthine derivatives (Zhou et al., 1992; Salvatore et al., 1993; van Galen et al., 1994). Several classes of non-xanthine derivatives have also been developed as Ai, A2A, A2B and A3 receptor-selective antagonists. The characteristics of each class have been reviewed by Baraldi (Baraldi et al., 2000). The structure and binding affinities of several common xanthine and non-xanthine antagonists are shown in Fig. 4. One of the most potent and widely used Ai receptor antagonists, 8-cyclopentyl-l,3-dipropylxanthine (DPCPX) (Lohse et al, 1987; Klotz, 2000; Muller, 2001), is produced by the substitution of a cycloalkyl group at position 8 of the xanthine ring (See Fig. 4). This antagonist has previously been used to demonstrate the involvement of Ai receptors in various actions in the gastrointestinal tract. For example, DPCPX was shown to abolish adenosine-induced inhibition of gastrin release from canine antral G-cells (Schepp et al., 1990), inhibit CPA-induced contraction in the rat colonic muscularis mucosae (Bailey et al., 1992; Reeves et al., 1993), and inhibit CPA-induced relaxation in the rat duodenal longitudinal muscle (Nicholls et al., 1996). The first A2 receptor antagonist, 3,7-dimethyl-l-propargylxanthine (DMPX), was generated by substitutions at the 1, 3, and 7 positions of the xanthine ring (Seale et al., 1988). Although this compound does not exhibit extremely high selectivity or affinity for the A2 receptors, it remains one of the most widely used A2 receptor antagonists due to its high solubility in water (Muller et al., 1997). More recently, a potent and highly selective non-xanthine A2A receptor antagonist, 4-(2-[7-Amino-2-(2-furyl)[l,2,4]triazolo[2,3-a][l,3,5]triazin-5-ylamino]ethyl)phenol (ZM 241385), was developed (Palmer et al., 1995b; Poucher et al., 1995, 1996). This antagonist, shown in Fig. 4, binds with high affinity to membranes expressing the A2A receptor (Palmer et al., 1995b; Poucher et al., 1995), but binds with low affinity to membranes expressing the Ai and A3 receptors (Poucher et al., 1995). ZM 241385 was also shown not to antagonize A2B receptor-mediated actions in the guinea pig aorta (Poucher et al., 1995). While selective A2B receptor agonists are not available, several selective A2B antagonists have been developed. These include 1,3-dialkylxanthine derivatives, and 8-phenylxanthine carboxylic congener derivatives (Kim et al., 2000). Some A2B receptor antagonists demonstrate species-dependent differences in ligand binding affinity (Fozard et al., 2003). However, these differences are not as apparent as in the A3 receptor. In humans, A3 receptors are blocked by high concentrations of xanthine derivatives such as DPCPX (Salvatore et al., 1993). In rats, these receptors were not blocked by xanthine derivatives, including DPCPX (Zhou et al., 1992), but instead were blocked by a number of non-xanthine derivatives such as 9-chloro-2(2-furyl)[l,2, 4]triazolo[l,5-c]quinazolin-5-amine (CGS 15943) (Baraldi and Borea, 2000). Since a number of selective adenosine agonists and antagonists are available, a pharmacological approach may be used to characterize the adenosine receptor subtype(s) involved in mediating various physiological actions. 2.3 Molecular characterization of the adenosine receptors The four adenosine receptors subtypes have been cloned and characterized in various tissues of different species, including the human brain (Furlong et al., 1992; Libert et al., 1992; Pierce et al., 1992; Salvatore et al., 1993) and rat brain (Reppert et al., 1991; Fink et al., 1992; Stehle et al., 1992; Zhou et al., 1992). In this section, the cloning and characterization of the human and rat adenosine receptor genes are examined. In addition, the regulation of adenosine receptor gene and receptor expression is also reviewed. Emphasis is placed on studies related to the Ai and A2A receptors since these two receptors are the focus of the current study. Both the Ai and A2A receptors were first cloned from a canine thyroid library (Libert et al., 1989), and thought to represent new members of the G protein-coupled family before they were recognized as clones of the Ai and A2A receptor (Furlong et al., 1992; Libert et al., 1992). The human Ai receptor is localized on chromosome lq32.1 (Townsend-Nicholson et al., 1995), and contains six exons and five introns (Libert et al., 1992). All introns are present in the 5' untranslated region (UTR) except for one, which interrupts the coding region at a position corresponding to the second intracellular loop. The coding region of the human Ai receptor consists of 987 bp (Libert et al., 1992). The coding region of the rat brain receptor was similar and consists of 981 bp (Mahan et al., 1991; Reppert et al., 1991). Two distinct Ai receptor transcripts, a 3.5 kb and a 5.5 kb transcript, were revealed by Northern blot analysis in rat tissues (Reppert et al., 1991). Both transcripts are expressed throughout the body in various tissues, including the brain and stomach. Another study by Mahan and co-workers has also identified two distinct transcripts, 3.1 kb and 5.6 kb in size, which have been detected in various tissues, including the rat brain and stomach (Mahan et al., 1991). The 3.5 and 3.1 kb species were found to be the predominant forms in the stomach (Mahan et al., 1991; Reppert et al., 1991). Two human Ai receptor transcripts have also been demonstrated in the brain (Ren and Stiles, 1994a). These transcripts differ in the 5' untranslated region (UTR) by 54 bp, and were initially thought to occur through alternative splicing (Ren and Stiles, 1994a). However, two different promoters, promoters A and B (Ren and Stiles, 1995), were later found to control the synthesis of the two transcripts. For the rat A\ receptor, Reppert has suggested that the two transcripts are generated by posttranscriptional modifications (Reppert et al., 1991). However, the promoter regions and 5' UTR of the rat Ai receptor gene have not been identified. Thus, the origin of the two rat Ai receptor transcripts remains unclear. The human and rat A2A receptor genes exhibit some similarities with each other (Chern et al., 1992; Fink et al., 1992; Furlong et al., 1992). The human A2A receptor gene is 9-10 kb in size, localized on chromosome 22ql 1.2 (Le et al., 1996), and contains 2 exons separated by a 6-6.4 kb intron, localized at a position corresponding to the second intracellular loop (Le et al., 1996; Peterfreund et al., 1996). Northern blot analysis reveals the expression of only one human A2A transcript that is 2.6 kb in size (Peterfreund et al., 1996). The rat A2A receptor gene was also shown to contain 2 exons, separated by a 7.2 kb intron (Chu et al., 1996). The 5' UTR of the rat A2A receptor has also been examined, and contains two independent promoters situated 338 bp apart (Chu et al., 1996). These two promoters were shown to drive the synthesis of two distinct transcripts that differ in the 5' UTR but encode for the same polypeptide sequence. The rat A2B receptor was first isolated from a rat brain cDNA library (Stehle et al., 1992). Extremely high levels of two distinct A2B receptor transcripts, a 1.8 kb and a 2.2 kb transcript, were found by Northern blot analysis and were expressed in the large intestine and caecum in rats (Stehle et al., 1992). In humans, two A2B receptor transcripts, a 1.8 kb and a 2.4 kb transcript, were also shown to be expressed in VA 13 fibroblast cells (Rivkees and Reppert, 1992). The origin of the two human and rat A2B receptor transcripts is unclear since the promoter regions and 5' UTR of these genes have yet to be determined. The human A2B receptor has been shown to contain a single intron at a position corresponding to the second intracellular loop (Pierce et al., 1992; Jacobson et al., 1995), and is localized on chromosome 17p 12 (Jacobson et al., 1995). The A3 receptor was first discovered by its cloning and characterization in the rat brain (Zhou et al., 1992). This receptor also contains a single intron at a position corresponding to the second intracellular loop. A variant of the rat A3 receptor, the A3i, has also been cloned and characterized (Sajjadi et al., 1996). This transcript is identical to the A3 receptor cloned by Zhou, except for a 57 bp insertion in the coding region, which occurs from alternative splicing of the intron. This variant was detected in a number of tissues, including the heart, brain, liver and testis (Sajjadi et al., 1996). However, it is unclear whether this variant is also present in the stomach. Evidence suggests that alternative splicing does not occur in the human A3 receptor gene (Murrison et al., 1996) and only one transcriptional start site was identified when the 5' UTR of this gene was analyzed (Murrison et al., 1996). The human A3 receptor gene is localized on chromosome lpl3.3 (Atkinson et al., 1997), and like the other adenosine receptor genes, it contains an intron (~2.2 kb) in the second intracellular loop. Of the four adenosine receptor subtypes, only the rat A3 receptor has been shown to be alternatively spliced within its coding region. The presence of a variant A3 receptor may have functional implications. 2.4 Regulation of adenosine receptor gene and protein expression The synthesis and expression of adenosine receptors can be regulated at a transcriptional or post-transcriptional level, and may be altered by changes in the physiological environment such as during periods of stress (see Table 1). The genomic structure of the Ai, A2A, and A3 receptor has been examined and the promoter regions of these genes contain putative binding sites for various transcription factors. A number of binding sites have been identified in the two promoters of the human Ai receptor gene (Ren and Stiles, 1995, 1998; Rivkees et al., 1999). Some of these sites were shown to interact with glucocorticoids in the regulation of Ai receptor gene transcription (Ren and Stiles, 1999). The promoter region of the rat Ai receptor gene has not been defined. However, as in humans, the transcription of this gene is regulated by glucocorticoids (Svenningsson and Fredholm, 1997), suggesting that the promoter region of the rat Ai receptor may also contain binding sites for glucocorticoids. The A2A receptors also respond to glucocorticoid treatment. Dexamethasone, a synthetic glucocorticoid, has been shown to attenuate the action of NECA in stimulating cAMP production (Gerwins and Fredholm, 1991), suggesting that glucocorticoids decrease A2A receptor expression. However, it is unclear if glucocorticoids act at a transcriptional or post-transcriptional level to regulate A2A receptor expression. A transcriptional repressor of the A2A receptor gene has recently been identified in the rat and has been shown to bind to specific nuclear factor-1 proteins to repress gene transcription (Lee et al., 2003). The A2A receptor has also been shown to regulate the expression of its own mRNA (Saitoh et al., 1994). Activation of the A2A receptor has been shown to transiently increase and then decrease A2A receptor gene and protein expression in PC 12 cells (Saitoh et al., 1994). The translation of Aj and A2A receptor mRNA can also be regulated by the presence of out-of-frame AUG codons in the 5' UTR (Ren and Stiles, 1994a; Chu et al., 1996; Lee et al., 1999). Translational regulation of the A2A receptor may occur in other species as well, since the 5' UTR of the rat A2A receptor gene exhibits high interspecies homology (Chu et al., 1996). Stress-induced changes in Ai and A2A receptor gene and protein expression have also been demonstrated. During periods of oxidative stress, Ai and A2A receptor gene and/or protein expression is upregulated (Nie et al., 1998; Kobayashi and Millhorn, 1999). Ai receptor mRNA and receptor expression were increased in DDT MF-2 cells during oxidative stress. This was mediated by enhanced transcription of the Ai receptor by the transcription factor nuclear factor KB (NFKB) (Nie et al., 1998). In the gut, oxidative stress induced by chronic ileitis has also been shown to increase Ai receptor gene expression (Sundaram et al., 2003). In PC 12 cells, stress induced by hypoxia was shown to increase A2A receptor mRNA and to increase A2A receptor protein expression through the synthesis of new A2A receptors (Kobayashi and Millhorn, 1999). These changes were mediated by PKC and may result from an increase in A2A receptor transcription. This is consistent with another study showing that the transcription of A2A receptors was enhanced in human neuroblastoma SH-SY5Y cells by the activation of PKC (Peterfreund et al., 1997). Other forms of stress have also been shown to alter A\ receptor expression in the rat. Stress induced by sustained performance, such as repeated restraint stress, was shown to exhibit increased Ai receptor agonist binding in the rat hypothalamus (Anderson et al., 1988). Rats suffering from acute renal failure were also shown to exhibit increased Ai receptor binding in renal membranes and increased Ai receptor mRNA expression (Gould et al., 1997). Gould has suggested that the increased Ai receptor expression was due to enhanced Ai receptor transcription. Stress induced by sleep deprivation has also been shown to increase Ai receptor mRNA expression in rats (Basheer et al., 2000). Prolonged wakefulness was shown to increase extracellular adenosine levels, which increased Ai receptor activation. Ai receptor activation was then shown to enhance the activity of the transcription factor NFKB, which initiates transcription of the Ai receptor (Basheer et al., 2001). Changes in Ai and A2A receptor expression can also be induced pharmacologically. Treatment of DDT 1 MF-2 cells with R-PIA was shown to decrease Ai receptor expression in membrane preparations within hours (Ramkumar et al., 1991). Chronic treatment with R-PIA has also been shown to decrease Ai receptor expression in isolated adipocytes (Green et al., 1992). In the presence of caffeine, an adenosine antagonist, both Ai and A2A receptor expression are increased. Chronic daily intraperitoneal treatments with caffeine also increase A] receptor expression (Fredholm, 1982), although long-term oral treatment with caffeine did not alter Ai receptor mRNA or receptor expression (Johansson et al., 1993). Similarly, in the mouse brain, striatal A2A receptor expression was upregulated by chronic intraperitoneal caffeine treatment (Traversa et al., 1994), while long-term oral caffeine treatment did not alter A2A receptor mRNA or receptor expression (Johansson et al., 1993). These contradictory findings may result from the different route of drug administration and the preferential exposure of these compounds to different sites of the body. The regulation of A2B receptor gene and protein expression is not as well understood. A study in chicken melanocytes has shown that A2B receptor mRNA expression can be upregulated by c-MyB (Worpenberg et al., 1997), a transcription factor that is encoded by the c-myb proto-oncogene. In human melanocytes, cell damage induced by 4-tertiary butylphenol, a phenolic agent, was shown to upregulate A2B receptor expression (Le Poole et al., 1999). In mouse bone marrow-derived macrophages, A2B mRNA expression can also be upregulated by IFN-y, a cytokine involved in activating macrophages (Xaus et al., 1999). The genomic structure of the A3 receptor has been examined, and a number of binding motifs have been identified in the 5' UTR of the human A3 gene (Murrison et al., 1996; Atkinson et al., 1997). These transcriptional binding sites may be important in regulating the promoter activity of the human A3 receptor. Possible binding sites have also been identified in the promoter of the mouse A3 receptor gene (Zhao et al., 1999; Yaar et al., 2002). In particular, the GATA-6 and CRE binding sites are involved in enhancing and inhibiting A3 receptor promoter activity, respectively (Yaar et al., 2002). The regulation of A3 receptor gene expression has some similarities to that of the Ai receptor. As with the Ai receptor, glucocorticoids and oxidative stress upregulate A3 receptor mRNA expression (Ramkumar et al., 1995; Sundaram et al., 2003). A3 receptor mRNA expression was significantly increased by dexamethasone treatment (Ramkumar et al., 1995) and by oxidative stress during chronic ileitis (Sundaram et al., 2003). Alterations in A3 receptor expression can also be induced pharmacologically. For example, the exposure of CHO cells to NECA for 24 h has been shown to significantly decrease A3 receptor expression (Palmer et al., 1995a). The studies presented in this section demonstrate that adenosine receptor gene and protein expression may be regulated through transcriptional and post-transcriptional mechanisms. In addition, the expression of these receptors can be altered by stress-induced changes and by treatment with various adenosine analogs. 2.5 Structural characterization of the adenosine receptors All four adenosine receptor subtypes belong to the G protein-coupled receptor family, which signal by activating various heterotrimeric G proteins (Olah and Stiles, 1995). The adenosine receptors are characterized by seven hydrophobic transmembrane domains, an extracellular N-terminal, an intracellular C-terminal, and three intracellular and three extracellular loops (Ralevic and Burnstock, 1998). In the rat, the Aj, A2A, A2b and A3 receptors are comprised of 326-7, 410, 332 and 320 amino acids, respectively (Mahan et al., 1991; Reppert et al, 1991; Stehle et al., 1992; Zhou et al., 1992; Chu et al., 1996). The large size of the A2A receptor results from its extended carboxyl tail (see Fig. 5). Each transmembrane domain consists of approximately 21-28 amino acids and all four receptors are glycosylated by asparagine (N)-linked glycoproteins at the second extracellular loop. Presently, the precise three-dimensional conformation of the adenosine receptors is unknown. Structural analysis identified receptor domains that participate in ligand recognition and binding. Most of these studies concentrate on the Ai, A2A, and A3 receptors, due to the lack of highly selective A2B receptor analogs. Molecular modeling of the A] I s «•** I V v v _ - • » fi N P S L Q T svvi M V i r L V A t g V1»Y r - 1 v Dfl R V K L M S '/A • M 1 v"c\ 6 B V p VLl "e i M M », 1 V ' V R Y K UlLSIS ^JT" Y rim I E T Y L I. I L cvvcn B P S D S L D T IIIIII B Fig. 5. Schematic representation of the human adenosine A, (A), A2A (B), and A3 (C) receptors. Transmembrane domains are numbered from 1 to 7, and amino acids shaded in black have been demonstrated, by site-directed mutagenesis, to participate in ligand binding, (figures are adapted from Olah and Stiles, 2000, and Gao et al., 2002). g (IJzerman et al., 1992), A2A (IJzerman et al., 1994; Kim et al., 1995) and A3 (van Galen et al., 1994) receptors has also been performed based on the structure of bacteriorhodopsin and/or rhodopsin, two structurally well characterized 7 transmembrane-spanning receptors. Based on these modeling studies and additional site-directed mutation studies, several amino acids were found to be involved in ligand binding at the Ai, A2A and A3 receptors (See Fig. 5). 2.6 Signal transduction mechanism Table 1 shows that multiple signal transduction mechanisms and second messengers are involved in adenosine receptor signaling (Ralevic and Burnstock, 1998; Fredholm et al., 2001). The Ai receptor couples to the G;/G0 family of G proteins (Freissmuth et al., 1991; Jockers et al., 1994). Stimulation of Ai receptors inhibits adenylate cyclase activity through the release of Gai, which decreases cAMP production and modulates cAMP-dependent protein kinase activity (Londos et al., 1980; Ralevic and Burnstock, 1998). This receptor also signals through a number of other messengers, such as phospholipase C (PLC), PKC, Ca2+ channels, K+ channels, and mitogen activated protein kinases (MAPK), as described in Table 1. The A2A receptor is coupled to the Gs protein in most tissues (Olah, 1997), and to G0if, a member of the Gs subfamily of G proteins, in the striatum (Kull et al., 2000). Activation of the A2A receptor can increase adenylate cyclase activity and cAMP production (Ralevic and Burnstock, 1998). However, these receptors can also mediate changes in the activity of other second messengers such as Ca2+ channels, KATp channels, and MAPK (Table 1). Like the A2A receptors, the A2B receptors also stimulate adenylate cyclase activity and increase cAMP production (Ralevic and Burnstock, 1998) through activation of the Gs protein (Pierce et al., 1992). The A2B receptors can also couple to Gq/Gi i to regulate PLC Table 1: Adenosine receptor G protein coupling, signal transduction, and regulation of expression Receptor Subtype G protein Coupling Effectors Second Messengers Receptor Expression A, Gj, G0 4-AC TPLC IPLC 1 cAMP (van Calker et al., 1979) T IP3, DAG, PKC (Gerwins and Fredholm, 1995) 1IP3, DAG, PKC (Delahunty et al., 1988) T PLD (Gerwins and Fredholm, 1995) TK+ channels (Belardinelli et al., 1995) iCa2+ channels (Dolphin et al., 1986) t MAPK (ERK1/2, P42/p44, p38) (Dickenson et al., 1998; Schulte and Fredholm, 2000; Robinson and Dickenson, 2001) upregulated by glucocorticoid treatment (Svenningsson and Fredholm, 1997), oxidative stress (Nie et al., 1998), nuclear factor KB binding (Nie et al., 1998), caffeine treatment (i.p.) (Fredholm, 1982), and stress [sustained performance stress (Anderson et al., 1988), acute renal failure (Gould et al., 1997), sleep deprivation (Basheer et al., 2000)] suppressed by agonist treatment (PIA) (Green et al., 1992), and out of frame AUG codons (Ren and Stiles, 1994) A2A Gs, Goif t AC t cAMP (Furlong et al., 1992) t Ca2+ channels (Gubitz et al., 1996) f K+ channels (Akatsuka et al., 1994) t MAPK (ERK1/2) (Sexl et al., 1997; Schulte and Fredholm, 2000) upregulated by oxidative stress (Kobayashi et al., 1998), PKC activation (Peterfreund et al., 1997), and treatment with caffeine (i.p.) (Traversa et al., 1994). suppressed by glucocorticoid treatment (Gerwins and Fredholm, 1991), binding of nuclear factor-1 proteins (Lee et al., 2003), A2A receptor activation (Saitoh et al., 1994), and out of frame AUG codons (Chu et al., 1996). A2B Gs, Gq/Gn TAC tPLC tcAMP (Feoktistov and Biaggioni, 1995) t IP3, DAG, PKC (Feoktistov and Biaggioni, 1995) TMAPK (ERK 1/2, JNK, p38) (Feoktistov et al., 1999; Schulte and Fredholm, 2000) upregulated by c-MyB transcription factor (Worpenberg et al., 1997), cell damage (Le Poole et al., 1999), and IFN-y (Xaus et al., 1999). A3 Gj, Gq/Gn iAC TPLC lcAMP(Zhou et al., 1992) t IP3, DAG, PKC (Abbracchio et al., 1995) TMAPK (ERK 1/2) (Schulte and Fredholm, 2000) upregulated by GATA-6 binding (Yaar et al., 2002), oxidative stress (Sundaram et al., 2003), and glucocorticoid treatment (Ramkumar et al., 1995). suppressed by CRE binding (Yaar et al., 2002), and treatment with agonist (NECA)(Palmer et al., 1995) and MAPK activity (Yakel et al., 1993; Feoktistov and Biaggioni, 1995; Schulte and Fredholm, 2000) (Table 1). The A3 receptor is linked to the inhibition of adenylate cyclase activity (Zhou et al., 1992) through activation of the Gj protein (Palmer et al., 1995a), and can alter PLC and MAPK activity via activation of the Gq/Gn protein (Abbracchio et al., 1995; Schulte and Fredholm, 2000), as shown in Table 1. Since each adenosine receptor subtype is coupled to specific signal transduction mechanisms, adenosine can mediate specific actions throughout the body by binding to different adenosine receptor subtypes. 3. Biological actions of adenosine in the gastrointestinal tract In the gastrointestinal tract, adenosine has been shown to modulate neurotransmitter release (Christofi, 2001), to alter gastric mucosal blood flow (Nagata et al., 1996; Burnstock, 2001), and to regulate gastric endocrine and exocrine secretion (Burnstock, 2001). The objectives of this thesis were to examine the role of the adenosine receptors in modulating gastrin and somatostatin secretion and to examine adenosine receptor gene expression under different physiological states of the stomach. A detailed review of the gastrointestinal actions of adenosine is presented in this section with emphasis on the role of adenosine in regulating gastric hormone secretion. Adenosine modulates gastrointestinal motility, neurotransmission, and secretion in various species by acting on different adenosine receptors (Burnstock, 2001). Although all four adenosine receptors have been localized in the enteric nervous system of humans (Christofi et al., 2001), few biological studies have studied the gastrointestinal effects of adenosine in humans. The majority of biological studies have been performed in animal models such as the guinea pig and rat (Burnstock, 2001; Christofi, 2001), where adenosine modulates gastrointestinal neurotransmitter release and motility. In the guinea pig ileum, adenosine has been shown to inhibit peristalsis (Shinozuka et al., 1985) and to inhibit cholinergic (Wiklund and Gustafsson, 1987) and adrenergic neurotransmission (Barajas-Lopez et al., 1991) via activation of presynaptic Ai receptors (Barajas-Lopez et al., 1991; Nitahara et al., 1995). Adenosine also depolarizes submucosal neurons of the ileum through activation of A2A receptors (Barajas-Lopez et al., 1991). In the guinea pig colon, adenosine was shown to inhibit contraction through activation of A2B receptors on smooth muscle cells (Kadowaki et al., 2000). Thus, adenosine can act on different adenosine receptor subtypes to exert distinct actions on various regions of the guinea pig gastrointestinal tract. In the rat, adenosine also modulates gastrointestinal neurotransmission, motility, and secretion. Adenosine was shown to induce relaxation of the gastric fundus (Matharu and Hollingsworth, 1992), enhance lower gastrointestinal propulsion (Suzuki et al., 1995), and inhibit peristalsis in the jejunum (Coupar and Hancock, 1994). These actions may be mediated by the activation of various adenosine receptor subtypes. In the rat ileum, the Ai subtype was shown to modulate the myenteric reflex response by eliciting ascending contraction and descending relaxation (Storr et al., 2002). In the rat duodenum, Ai and A2B receptors were found to inhibit contraction of the longitudinal muscle and stimulate contraction of the muscularis mucosae, respectively (Nicholls et al., 1996). In the rat stomach, adenosine has been shown to alter gastric mucosal blood flow (Cho and Ogle, 1990; Cho et al., 1991; Cho et al., 1995), and to regulate ethanol-induced gastric dilation in submucosal arterioles (Nagata et al., 1996). Our laboratory demonstrated that the administration of the A2A receptor agonist, CGS 21680, significantly decreases perfusion pressure in the isolated vascularly perfused rat stomach (Yip and Kwok, 2004), suggesting that the A2A receptor subtype may be involved in mediating gastric vasodilation. Adenosine also regulates gastric endocrine and exocrine secretion. In particular, adenosine and its analogs can inhibit (Gerber et al., 1984; Glavin et al., 1987; Scarpignato et al., 1987), stimulate (Puurunen et al., 1986), or have no effect (Puurunen et al., 1987; Puurunen and Huttunen, 1988) on gastric acid secretion, depending on the route of drug administration. This nucleoside can also inhibit gastrin release (DeSchryver-Kecskemeti et al., 1981; Harty and Franklin, 1984; Kwok et al., 1990) and stimulate or inhibit gastric somatostatin release, depending on the concentration administered (Kwok et al., 1990). The gastric secretory actions of adenosine are discussed in section 3.2. 3.1 Gastroprotective actions of adenosine In humans, caffeine has been implicated in promoting gastric ulcer formation (Cooke, 1976). Although the mechanism remains unclear, adenosine antagonists such as caffeine and theophylline were shown to stimulate gastric acid secretion (Krasnow and Grossman, 1949), possibly by inhibiting the action of adenosine. Studies have suggested that the gastric juice of ulcer patients contains decreased levels of adenosine (Durak et al., 1994) since xanthine oxidase activity and 5'nucleotidase activity in these patients were shown to be increased and decreased, respectively. As reviewed in Section 1,5' nucleotidase is responsible for the synthesis of adenosine (see Fig. 3), while xanthine oxidase is involved in the metabolism of adenosine (see Fig. 1). Ulcer patients also exhibit decreased ADA activity in the vicinity of a healed ulcer after ranitidine treatment (Namiot et al., 1993) and increased ADA activity in the area surrounding an unhealed ulcer (Namiot et al., 1993). ADA is a metabolic enzyme of adenosine (see Fig. 3). Thus, it is likely that increased and decreased adenosine concentrations are also found surrounding a healed and unhealed ulcer, respectively. These observations suggest that adenosine may be involved in the healing process and may be important in protecting the stomach against ulcer formation. In rats, adenosine can act as a gastroprotective agent. Adenosine protects against indomethacin- (Bozkurt et al., 1998), ethanol- (Cho and Ogle, 1990; Cho et al., 1991), cold restraint stress- (Geiger and Glavin, 1985; Westerberg and Geiger, 1987), and ischemia/reperfusion- (Konturek et al., 2001; Pajdo et al., 2001) induced gastric ulcer formation. The ability of the non-selective adenosine antagonists 8-phenyltheophylline (8-PT) or 8-(p-sulfophenyl)theophylline (8-SPT) to block the protective action of adenosine in some of these studies confirms that adenosine receptors are involved (Geiger and Glavin, 1985; Westerberg and Geiger, 1987; Konturek et al., 2001; Pajdo et al., 2001). In indomethacin- and ethanol-treated rats, adenosine was also shown to inhibit indomethacin-induced gastric acid secretion (Bozkurt et al., 1998) and basal acid secretion (Cho and Ogle, 1990), respectively. In ethanol- and ischemia-induced ulcer studies, an increase in gastric mucosal blood flow was also observed (Cho et al., 1991; Konturek et al., 2001; Pajdo et al., 2001). Thus, the ability of adenosine to inhibit gastric acid secretion and to increase gastric mucosal blood flow may play an important role in the gastroprotective action of adenosine. Although the action of adenosine is mainly protective, studies by Ushijima have shown that adenosine may also exacerbate gastric mucosa damage (Ushijima et al., 1985, 1992). Contrary to Westerberg (Westerberg and Geiger, 1987), the administration of adenosine analogs increased restraint stress-induced gastric ulcer formation in rats. The cause of this functional difference is currently undetermined. 3.2 Effect of adenosine on gastric acid secretion In humans, indirect evidence suggests that adenosine is involved in regulating gastric acid secretion. Studies have shown that fundic ADA activity increases in parallel with gastric acid output (Namiot et al., 1990). Since ADA is a metabolic enzyme of adenosine, an increase or decrease in the activity of this enzyme should decrease or increase adenosine levels, respectively. Patients suffering from hypersecretion of acid exhibited higher fundic ADA activities (Namiot et al., 1990), while those suffering from the absence of gastric acid secretion, a condition known as achlorhydria, exhibited lower ADA activities. The inhibition of gastric acid secretion by ranitidine also decreased ADA activity (Namiot et al., 1991). Thus, in humans, it is possible that gastric acid secretion is regulated, in part, by ADA activity, and consequently by the adenosine level. The role of adenosine in regulating gastric acid secretion is also supported by the gastric action of caffeine and theophylline, two non selective adenosine receptor antagonists. Both of these compounds stimulate gastric acid secretion (Krasnow and Grossman, 1949; Cohen and Booth, 1975), and may mediate this action by blocking the inhibitory effect of endogenously released adenosine. Studies using experimental animal models indicate that the effect of adenosine on gastric acid secretion is likely to be species-specific, and may depend on the site(s) at which adenosine acts. Gastric acid is secreted by parietal cells which reside in the corporeal mucosa of the stomach (Sachs et al., 1997). The regulation of gastric acid secretion is complex and is controlled by neural, paracrine, and endocrine pathways that modulate the release of stimulatory mediators such as gastrin, acetylcholine, and histamine, and the release of inhibitory compounds such as somatostatin and prostaglandins (Hersey and Sachs, 1995). Adenosine has been shown to inhibit gastric acid secretion in the dog, guinea pig, and rat, but in rabbits, adenosine stimulates gastric acid secretion. In isolated gastric glands (Gil-Rodrigo et al., 1990) and enriched parietal cells (Ota et al., 1989; Ainz et al., 1993) of the rabbit, adenosine and its analogs were shown to significantly stimulate basal and potentiate histamine-stimulated aminopyrine uptake. [14C]aminopyrine uptake was used as an index of gastric acid secretion since it can become protonated and trapped in the acidic canaliculi of the parietal cell (Berglindh et al., 1976). The rank order of potency of different agonists in stimulating acid secretion was shown to be NECA > 2-chloroadenosine (2-CA) > adenosine, which suggests the involvement of adenosine A2 receptor (Ota et al., 1989). The stimulatory action of adenosine on isolated gastric glands (Gil-Rodrigo et al., 1990) and NECA on enriched parietal cells (Ainz et al., 1993) were also inhibited by theophylline. Since activation of A2 receptors increases cAMP production, this proposal is consistent with findings demonstrating the ability of adenosine to increase cAMP levels in the enriched rabbit parietal cell preparation (Ota et al., 1989). In these experiments, the effect of adenosine was not altered by the endogenous release of the acid secretagogue histamine (Ainz et al., 1993) or by the production of the acid inhibitor prostaglandin E2 (Ota et al., 1992). Thus, in the rabbit, adenosine may act directly on the A2 receptors of parietal cells to stimulate gastric acid secretion. In this species, ATP was shown to inhibit histamine-stimulated aminopyrine uptake in the isolated gastric gland (Gil-Rodrigo et al., 1990). Further experiments demonstrated that this is likely due to the activation of ATP receptors, of the P2y subtype, on the parietal cells (Gil-Rodrigo et al., 1996). The inhibitory action of ATP is consistent with the localization of P2y receptors on gastric glands of the rabbit fundus (Vallejo et al., 1996). Thus, in rabbits, purinergic compounds are able to directly alter gastric acid secretion. In particular, adenosine can act on A2 receptors and ATP can act on P2y receptors to stimulate and inhibit gastric acid secretion, respectively, from parietal cells. In other species, such as the dog, adenosine has been shown to inhibit histamine- and methacholine-stimulated gastric acid secretion in vivo (Gerber et al., 1984). This inhibitory effect may be due to the direct action of adenosine on the parietal cells since 2-chloroadenosine (2-CA) and R-PIA have been shown to inhibit histamine-stimulated aminopyrine uptake from isolated canine parietal cells (Gerber et al., 1985). Activation of Ai receptors is likely involved in this action since the Ai selective analog R-PIA was more potent than 2-CA in inhibiting histamine-stimulated cAMP accumulation. Additional studies have also shown that the inhibitory effect of adenosine is unlikely to be due to changes in the release of the acid secretagogue histamine (Payne and Gerber, 1997), or the activity of phosphodiesterase (Gerber et al., 1984, 1985). Phosphodiesterase metabolizes cAMP, and inhibitors of this enzyme, such as high concentrations of theophylline and caffeine, may stimulate gastric acid secretion (Barnette et al., 1995). Furthermore, adenosine does not mediate its effect by altering the synthesis of prostaglandin E2, an inhibitor of gastric acid secretion (Gerber et al., 1984; Geiger and Glavin, 1985). These findings suggest that, in the dog, adenosine may act directly on Ai receptors of the parietal cells to inhibit gastric acid secretion. In dogs, adenosine has also been shown to modulate the release of gastrin (Schepp et al., 1990), a potent gastric acid secretagogue (Hersey and Sachs, 1995). In isolated canine antral cells, R-PIA was shown to significantly inhibit forskolin-stimulated gastrin release, while CV 1808 had no effect on this action (Schepp et al., 1990). CV1808, however, slightly stimulated basal gastrin release and potentiated bombesin-stimulated gastrin release (Schepp et al., 1990). Since R-PIA and CV 1808 preferentially bind to the Ai and A2 receptors, respectively, these results suggest that A\ receptors may be involved in inhibiting gastrin release, while the A2 receptors are involved in stimulating gastrin release. In guinea pigs, adenosine was shown to inhibit histamine and gastrin-stimulated acid secretion from oxyntic cells (Heldsinger et al., 1986). Adenosine and its analogs inhibited histamine-stimulated hydroxyl ion production with the following rank order of potency: R-PIA > NEC A > adenosine. The preferential action of R-PIA on Ai receptors suggests that this subtype is involved in the inhibitory action of adenosine on gastric acid secretion (Heldsinger et al., 1986). This adenosine Ai receptor-mediated inhibitory action is similar to that in the dog (Gerber et al., 1985). The gastric effects of adenosine have been studied extensively in the rat. Various studies have shown that adenosine inhibits, stimulates, or does not alter gastric acid secretion (see Table 2), depending on the route of drug administration, the state of anesthesia, and whether or not the vagus is intact. Intraperitoneal (i.p.), subcutaneous (s.c), intraduodenal (i.d.), oral, intragastric, and intracerebroventricular (i.c.v.) administration of adenosine and its analogs inhibited gastric acid secretion (Puurunen et al., 1986; Scarpignato et al., 1987; Puurunen and Huttunen, 1988; Westerberg and Geiger, 1989; Cho and Ogle, 1990), while intravenous (i.v.) administration of these drugs stimulated gastric acid secretion (Puurunen et al., 1986). The localization of the adenosine receptors and their preferential exposure to adenosine through different routes of administration may account for these functional differences. The various effects of adenosine suggest that this nucleoside may regulate gastric acid secretion through more than one site of action. Possible sites of action are addressed in the following discussion. Table 2: Adenosine and adenosine agonists on gastric acid secretion in the rat Administration Drug State Effect Reference i.p. R-PIA, S-PIA, 2-CA, NECA conscious •1 basal acid secretion i i.d adenosine anaesthetized no effect 2 i.d. R-PIA anaesthetized •I basal acid secretion 2 oral R-PIA anaesthetized i basal gastric secretion 2 intragastric R-PIA anaesthetized I basal gastric secretion 2 s.c. R-PIA anaesthetized 1 basal acid secretion 2 s.c. adenosine anaesthetized •I basal gastric secretion 2-5 s.c. R-PIA conscious 4- basal gastric secretion 6 s.c. S-PIA conscious no effect 6 s.c. adenosine conscious I indomethacin-induced acid secretion 7 i.v. R-PIA vagotomized & anaesthetized no effect 8 RPIA & NECA isolated parietal cells no effect 8 i.v. adenosine, NECA, R-PIA, CHA, S-PIA, 2-CA anaesthetized T basal acid secretion 8 i.v. R-PIA vagotomized & anaesthetized no effect 9 i.c.v. R-PIA anaesthetized no effect 9 i.c.v. R-PIA conscious I basal acid secretion 9 i.c.v. adenosine, NECA, R-PIA, S-PIA, conscious I basal acid secretion 10 i.c.v. NECA vagotomized & conscious no effect 10 i.c.v. NECA anaesthetized no effect 10 1 Westerberg and Geiger, 1989;2 Scarpignato et al., 1987;3-5 Cho and Ogle, 1990; Cho et al., 1991; Cho et al, 1995; 6Glavin et al., 1987; 7Bozkurt et al, 1998;8 Puurunen et al, 1987;9 Puurunen et al, 1986; 10 Puurunen and Huttunen, 1988. In conscious rats, the i.e.v. administration of adenosine has been shown to inhibit gastric acid secretion. This effect may be mediated by the action of adenosine on the central nervous system. The regulation of gastric acid secretion by the central nervous system involves the dorsomotor nucleus of the vagus (DMNV), the hypothalamus and the nucleus tractus solitarius (NTS) (Hersey and Sachs, 1995). The hypothalamus and NTS receive sensory input from the periphery and regulate the activity of the DMNV. Activation of the DMNV can stimulate gastric acid secretion. Central stimuli from the hypothalamus and the NTS are integrated at the DMNV, and efferent fibers originating from the DMNV travel to the stomach via the vagus nerve. The vagus also contains fibers originating from the nucleus ambiguus, but these nerves are mainly involved in controlling gastric motility (Hersey and Sachs, 1995). The i.e.v. administration of adenosine analogs to conscious rats with intact vagi inhibited gastric acid secretion with the following rank order of potency: NECA > R-PIA > S-PIA (Puurunen and Huttunen, 1988). The higher potency of NECA suggests that the A2 receptor is involved in this action. The i.c.v. administration of NECA, however, did not alter gastric acid secretion in vagotomized rats (Puurunen and Huttunen, 1988) or anaesthetized rats with intact vagi (Puurunen and Huttunen, 1988). Anesthetics can suppress vagal tone (Murthy et al., 1982; Yamamura et al., 1983). Thus, the central A2 receptors may inhibit gastric acid secretion by suppressing central stimulatory vagal impulses to the stomach. Furthermore, the inhibitory effect of NECA was attenuated by the removal of brain monoamines such as noradrenaline, dopamine and serotonin, but was not affected by opioid or prostaglandin inhibitors (Puurunen and Huttunen, 1988). Thus, in the brain, the A2 receptors may mediate the inhibitory effect on acid secretion by altering the release of biogenic monoamines. Gastric acid secretion was also inhibited by s.c, i.p., i.d., orally and intragastrically administered adenosine agonists (see Table 2). However, it is unclear whether this action is mediated centrally or peripherally. In one study, the s.c. administration of R-PIA was able to inhibit gastric acid secretion, while the s.c. administration of S-PIA had no effect (Glavin et al., 1987). In another study, the potency of NECA in inhibiting acid secretion was equal to that of R-PIA when administered i.p. (Westerberg and Geiger, 1989). Thus, both the Ai and A2 receptors may be involved in the inhibition of gastric acid secretion. Gastric acid secretion is regulated peripherally through several mechanisms. Acetylcholine, gastrin, and histamine stimulate gastric acid secretion by acting on the M3 receptor, CCKB receptor, and H2 receptor, respectively, which are expressed on parietal cells (Hersey and Sachs, 1995). Somatostatin and prostaglandins can also act directly on parietal cells to inhibit acid secretion (Schubert and Shamburek, 1990). Thus, adenosine may modulate gastric acid secretion by regulating the release of these substances. Unlike in other species, adenosine does not act directly on the parietal cells to alter gastric acid secretion in the rat. NECA and R-PIA were shown to have no effect on basal, histamine-stimulated, or carbachol-stimulated aminopyrine accumulation in the rat isolated enriched parietal cell preparation (Puurunen et al., 1987). Histamine and carbachol were shown to stimulate acid secretion by increasing cAMP and inositol phosphate levels, respectively, but neither NECA nor R-PIA elicited this effect (Puurunen et al., 1987). Histamine is released from enterochromaffm-like (ECL) cells of the stomach (Sachs et al, 1997). Its release is stimulated by gastrin, Ach, and P-adrenergic agonists, and is inhibited by somatostatin (Hersey and Sachs, 1995). Although the effect of adenosine on histamine has not been examined in the rat, adenosine was shown not to alter histamine release in both the dog and rabbit (Ainz et al., 1993; Payne and Gerber, 1997). Acetylcholine is released from the enteric nervous system and can stimulate gastric acid secretion either by acting directly on the parietal cells or indirectly by stimulating gastrin secretion or inhibiting somatostatin secretion (Hersey and Sachs, 1995). Studies on gastrointestinal motility have demonstrated that adenosine inhibits acetylcholine release from the myenteric plexus of several species. Adenosine was found to inhibit nicotinic neurotransmission in the antrum (Christofi et al., 1992) and ileum (Gustafsson, 1981). In the submucosal plexus, adenosine inhibits the presynaptic release of acetylcholine through activation of the Ai receptor (Barajas-Lopez et al., 1991). Although adenosine can regulate acetylcholine release, this action has only been implicated in the modulation of gastric motility. It is unclear whether alterations in acetylcholine release are involved in the adenosine-induced inhibition of gastric acid secretion. Adenosine may inhibit gastric acid secretion by stimulating the release of the gastric acid inhibitor, somatostatin. Somatostatin is released from D-cells of the gastric mucosa and from enteric nerves (Mcintosh, 1985). The stomach contains two distinct populations of D-cells, one residing in the corpus and the other in the antrum. In the present study, the term "corpus" refers to the acid-secreting portion of the stomach (see Fig. 36). However, a number of studies also refer to this region as the "fundus". Thus, in this chapter, both "corpus" and "fundus" are used to describe the acid-secreting region of the stomach. In Chapter 3 (Results), the term "fundus" will be used exclusively to describe the non-glandular region of the stomach. Somatostatin-containing D-cells of the fundus are localized at the basal mucosa and are characterized as small rounded or pyramid-shaped cells with long processes (Keast et al., 1985). In the antrum, D-cell density is higher than in the fundus (Keast et al., 1985). Antral D-cells are also localized at the base of the gastric gland (Buchan et al., 1985a) and are characterized as flask-shaped cells with long processes (Keast et al, 1985). The fundic and antral D-cells of the rat are distinct and respond differently to gastric acidity and the presence of food (Schwarting et al., 1986). Antral D-cells are open to the luminal environment (Hersey and Sachs, 1995). These cells detect changes in the intraluminal pH, and respond to decreases in gastric pH by releasing somatostatin (Schusdziarra et al., 1983). Increases in gastric pH, such as those induced by omeprazole treatment, have been shown to decrease somatostatin gene expression (Brand and Stone, 1988; Wu et al., 1990a; Wu et al., 1990b; Sandvik et al., 1995), plasma somatostatin concentration, antral somatostatin concentration, and antral D-cell density (Allen et al., 1986; Lee et al., 1992; Pawlikowski et al., 1992). Omeprazole is a drug that inhibits gastric acid secretion by binding irreversibly to the proton pump, H+K+ATPase, on parietal cells (Hirschowitz et al., 1995). The prandial state of the stomach also alters somatostatin synthesis and release. After prolonged fasting (96 h), antral D-cell density has also been shown to decrease (Schwarting et al., 1986). Fundic D-cells are closed cells that do not interact with the luminal environment. Unlike antral D-cells, changes to the intraluminal pH and prolonged fasting do not alter fundic somatostatin release (Schusdziarra et al., 1983) and D-cell density (Schwarting et al., 1986), respectively. Instead, somatostatin release from fundic D-cells may be stimulated by intrinsic (Schubert et al., 1991) and extrinsic (Sandvik et al., 1993) nerve stimulation. The distribution of gastric somatostatin-containing nerve fibers differs among species. In the rat stomach, these fibers are not present in the mucosa, external muscle or myenteric plexus, but a few single somatostatin-containing fibers have been observed in the muscularis mucosae (Keast et al., 1985). In another study, somatostatin-containing fibers were absent from the mucosa and submucosal plexus; however, a few somatostatin-containing fibers were demonstrated in the smooth muscle and in the myenteric ganglia (Ekblad et al., 1985). Somatostatin is released from the long processes extending out from the D-cells, and has been shown to act in a paracrine manner to inhibit gastrin release from G-cells, acid secretion from parietal cells (Park et al., 1987; Kusumoto et al., 1995) and histamine release from ECL cells (Prinz et al., 1993). Fundic D-cells have also been shown to be in close proximity to the vasculature (Kusumoto et al., 1995), and antral D-cell processes have been shown to project to the blood capillaries (Kusumoto and Grube, 1987). Thus, somatostatin may be released into the blood and act in an endocrine manner. Somatostatin release is inhibited by cholinergic agents (Koop et al., 1982), enkephalins (Mcintosh et al., 1983), substance P (Kwok et al., 1985; Mcintosh et al., 1987a), neurokinins and tachykinins (Kwok et al., 1988) and is stimulated by p-adrenergic agents (Koop et al., 1982; Koop et al., 1983), histamine (Mcintosh et al., 1987a), cholecystokinin (CCK) (DelValle et al., 1996), secretin/VIP (Chiba et al., 1980; Saffouri et al., 1984), calcitonin gene related peptide (CGRP) (Manela et al., 1995; Rasmussen et al., 2001), glucagon (Chiba et al., 1980) and bombesin/gastrin releasing peptide (GRP) (Martindale et al., 1982). Our laboratory has demonstrated that basal somatostatin release can also be stimulated by adenosine in a concentration-dependent manner (Kwok et al., 1990). The stimulatory effect of adenosine on SLI was not altered by atropine and hexamethonium, cholinergic antagonists, or by propranolol, a P-adrenergic antagonist. Therefore, adenosine-induced attenuation of gastric SLI release does not involve changes in cholinergic or adrenergic neurotransmission (Kwok et al., 1990). Adenosine was also shown not to mediate its stimulatory action on SLI through its vasodilatory effects since nitroprusside, a potent vasodilator, did not alter gastric SLI release (Yip and Kwok, 2004). It is possible that adenosine acts directly on A2A receptors on D-cells to stimulate somatostatin release. Activation of A2A receptors has been shown to stimulate adenylate cyclase activity, cAMP production, PKC activity, and to activate L-type Ca2+ channels (see Table 1). The same second messengers were shown to be involved in mediating somatostatin release from D-cells. In human cultured D-cells, somatostatin release was stimulated by activation of PKC, production of cAMP, and increases in intracellular Ca2+ concentration (Buchan et al, 1990). In rat fundic D-cells, bombesin-induced release of somatostatin was also found to be mediated through a calcium-phospholipid-dependent mechanism (Schaffer et al, 1997). Furthermore, in canine fundic D-cells, CCK-stimulated somatostatin release was mediated by an increase in intracellular Ca2+ induced by activation of L-type Ca2+ channels (DelValle et al, 1996). Since the A2A receptors are linked to second messengers which are involved in stimulating somatostatin release, it is possible that activation of A2A receptors on D-cells stimulates somatostatin release. Experiments have shown that somatostatin gene transcription is regulated by cAMP (Montminy et al, 1996a, 1996b). A cAMP response element (CRE), which confers cAMP inducibility, has been identified in the somatostatin gene (Comb et al, 1986) and can mediate the transcriptional induction of somatostatin. Activation of the A2A receptors has been shown to increase the expression of genes that contain a cAMP response element (CRE), such as the tyrosine hydroxylase gene (Chae and Kim, 1997). Adenosine may therefore regulate the synthesis and release of somatostatin by activating A2A receptors on D-cells. The cellular localization of these receptors, however, has not been determined in the stomach. Gastrin is a major hormonal stimulant of gastric acid secretion (Edkins, 1905; Kovacs et al., 1989). In the stomach, it is synthesized in G-cells of the antral mucosa. These cells are open to the luminal environment, and their activity is partially regulated by the gastric contents. Gastrin secretion is enhanced by a number of chemical and physical factors such as acetylcholine (Matsuno et al., 1997), P-adrenergic agonists (DeSchryver-Kecskemeti et al., 1981; Koop et al., 1983), bombesin/GRP (Martindale et al., 1982; Guo et al., 1988; Campos et al., 1990), vagotomy (Stening and Grossman, 1970; Pederson et al., 1984), proteins and amino acids (Guo et al., 1988; DelValle and Yamada, 1990), gastric distension (Lloyd et al., 1992), and intraluminal pH (Hoist et al., 1983). Gastric pH serves as a negative feedback mechanism since luminal acidification (pH < 3.5) has been shown to inhibit the further release of gastrin (Magee, 1996). Increased gastric pH induced by omeprezole treatment, conversely, has been shown to increase gastrin mRNA, antral gastrin and plasma gastrin levels (Brand and Stone, 1988; Inauen et al., 1988; Larsson et al., 1988; Dockray et al., 1991). Gastrin release is inhibited by the antral somatostatin-secreting D-cells (Saffouri et al., 1980). D-cells may act as chemoreceptors for intraluminal pH and modulate somatostatin secretion to control gastrin secretion (Hersey and Sachs, 1995). An inhibition of basal somatostatin release may stimulate gastrin secretion since basal gastrin secretion was shown to be tonically restrained by somatostatin (Saffouri et al., 1980). Somatostatin acts on somatostatin 2 receptors located on the G-cells (Zaki et al., 1996), and inhibits gastrin secretion by inhibiting adenylate cyclase activity (DelValle et al., 1990). Our laboratory has shown that adenosine can inhibit immunoreactive gastrin (IRG) release from the isolated vascularly perfused rat stomach in a concentration-dependent manner (Kwok et al, 1990). Adenosine was also found to inhibit carbachol- and norepinephrine-stimulated gastrin secretion in rat antral mucosa cells (DeSchryver-Kecskemeti et al, 1981), and to inhibit basal and carbachol-stimulated gastrin secretion in the rat antral mucosal fragments (Harty and Franklin, 1984). The effect of adenosine on gastrin secretion is probably not centrally mediated since the administration of NECA (i.c.v.) did not alter serum gastrin concentrations (Puurunen and Huttunen, 1988). The adenosine receptor subtype involved in inhibiting gastrin release in the rat has not been examined. However, in the dog, adenosine was suggested to inhibit gastrin secretion by activating Ai receptors localized on G-cells (Schepp et al, 1990). Activation of Ai receptors inhibits adenylate cyclase activity, cAMP production, and Ca2+ channel activity, and has also been shown to alter PKC activity (See Table 1) (Ralevic and Burnstock, 1998). The same second messengers are involved in regulating gastrin secretion. In isolated rat and pig parietal cells, cAMP is required to elicit aminopyrine accumulation (Cabero et al, 1993; Li et al, 1995). Cyclic AMP has also been shown to potentiate bombesin-evoked gastrin release from isolated perfused rat stomachs (Guo et al, 1988) and to stimulate gastrin release from isolated human G-cells in culture (Campos et al, 1990). Activation of PKC (Campos et al, 1990) and an elevation in intracellular Ca2+ concentration have also been shown to stimulate gastrin release (Campos et al, 1990; Buchan and Meloche, 1994; Seensalu et al, 1997). The release of gastrin from isolated G-cells was also shown to be mediated by the cytosolic Ca concentration (Campos et al, 1990; Buchan and Meloche, 1994; Seensalu et al, 1997). Thus, it is possible that activation of Ai receptors on G-cells leads to an inhibition of gastrin release through an inhibition of cAMP production and Ca2+ channel activity. The Ai receptor may also be involved in regulating gastrin gene expression since the expression of this gene has been shown to be cAMP-dependent (Shiotani and Merchant, 1995). II. Hypothesis I hypothesize that activation of gastric adenosine receptors plays a role in the regulation of gastric exocrine and endocrine secretion. In particular, adenosine may inhibit gastric acid secretion by inhibiting gastrin release and stimulating somatostatin release. I propose that adenosine acts on adenosine Ai receptors on G-cells and A2A receptors on D-cells to inhibit gastrin and stimulate somatostatin release, respectively. I also hypothesize that changes to the gastric state, such as those induced by fasting and omeprazole treatment, may alter adenosine receptor gene and protein expression to regulate gastrin and somatostatin release. III. Rationale Gastric acid secretion may be inhibited by adenosine through the regulation of gastrin and somatostatin release since adenosine has previously been shown to alter IRG and SLI release in the isolated vascularly perfused stomach. Adenosine was shown to inhibit basal and carbachol-stimulated gastrin release from isolated G-cell preparations (Schepp et al., 1990). Thus, this compound may act directly on Ai receptors localized on G-cells to inhibit gastrin release. Adenosine may also stimulate somatostatin release by acting on A2A receptors localized on the gastric D-cells since previous studies performed in our laboratory have shown that the effect of adenosine was not altered by cholinergic and adrenergic agonists. In addition, gastric adenosine receptor expression may be altered under different physiological conditions since changes to the prandial and acid secretory state of the stomach have been shown to regulate the synthesis and release of gastrin and somatostatin (Brand and Stone, 1988; Wu et al, 1991; Sandvik et al, 1993). IV. Objectives and specific aims The objectives of this thesis were to determine the adenosine receptor subtypes that are involved in regulating SLI and IRG release, to provide information about the distribution, localization, and structure of these receptors, and to study how adenosine receptors are regulated under different physiological states of the stomach. The specific aims of this study are outlined below: 1. Determine the adenosine receptor subtypes involved in regulating IRG and SLI release in the isolated vascularly perfused rat stomach, using selective adenosine receptor agonists and antagonists. 2. Determine the distribution of adenosine receptor gene expression in morphologically and functionally distinct regions of the rat stomach using RT-PCR, and determine the structure of the gastric Ai, A2A, and A3 receptor by cloning and sequencing. 3. Develop a sensitive (competitive RT-PCR and Real-Time RT-PCR) assay to measure adenosine Ai and A2A receptor gene expression in various regions of the rat stomach. 4. Examine whether adenosine receptor gene expression and function are regulated by different physiological states of the stomach, in particular, fasting and omeprazole treatment. 5. Investigate the cellular localization and distribution of gastric A\ and A2A receptors in the stomach by immunohistochemistry. Chapter 2: Methods Animals were treated in accordance with the guidelines of the University of British Columbia Committee on Animal Care. I. Effect of adenosine analogs on gastric SLI and IRG release 1. Isolated vascularly perfused rat stomach preparation Male Wistar rats (250 to 325 g) were housed in light and temperature controlled rooms with free access to food and water. Animals were deprived of food, but had free access to water for at least 14 h prior to experiments. Rats were anesthetized with an i.p. injection (60 mg/kg) of sodium pentobarbital (Somnotol®, MTC Pharmaceuticals, Cambridge, ON, Canada). The surgical procedures used to isolate the stomach for perfusion have previously been described (Kwok et al., 1990; Kwok and Mcintosh, 1990). The stomach was exposed by an abdominal midline incision. The superior mesenteric artery and vasculature supplying the left and right adrenal glands and kidneys were tied or cut between double ligatures. The pancreas and spleen were carefully dissected out from the stomach without damaging the gastroepiploic artery. To empty the content of the stomach, a drainage cannula was inserted into the stomach through the gastroduodenal junction. The pancreas, spleen, small and large intestines were then removed. Arterial perfusion was achieved by inserting a cannula into the aorta with the tip lying adjacent to the celiac artery. Perfusate was introduced into the stomach via this arterial cannula. Six hundred units of heparin (Sigma-Aldrich, St. Louis, MO) in 2 ml saline solution were immediately injected. The portion of the body above the diaphragm was removed. The vena cava was tied, and the portal vein was cannulated to allow for the collection of venous effluent. The stomach was perfused at a rate of 3 ml/min using a peristaltic pump (Cole-Parmer Instrument Co. Chicago, IL). The perfusate was composed of Krebs' solution (120 mM NaCl, 4.4 mM KC1, 2.5 mM CaCl2, 1.2 mM MgS04-7H20, 1.5 mM KH2P04, 25 mM NaHC03, and 5.1 mM dextrose) containing 0.2% bovine serum albumin (BSA, RIA grade; Sigma-Aldrich) and 3% dextran (Clinical grade; Sigma-Aldrich). The perfusate was continuously gassed with a mixture of 95% 02 and 5% C02 to maintain a pH of 7.4. Both 0 the perfusate and the stomach preparation were kept at 37 C by thermostatically-controlled heating units throughout the experiment. 2. Drug administration and sample collection Venous effluent was collected via the portal cannula. After a 30 min equilibration period, samples were collected at 5 min intervals. Three basal samples were obtained before drugs were introduced. Drugs were administered into the perfusate via side-arm infusion at a rate calculated to give the final perfusion concentrations. The following drugs were purchased from Sigma-Aldrich: adenosine hemisulphate salt, N6-cyclopentyladenosine (CPA), 2-p-(2-carboxyethyl) phenethylamino-5'N-ethylcarboxamidoadenosine HC1 (CGS 21680), 1 -deoxy-1 - [6- [[(3 -iodophenyl)methyl] amino] -9H-purin-9-yl] -N-methyl-B-D-ribofuranuronamide (IB-MECA), 8-cyclopentyl-l,3-dipropylxanthine (DPCPX), and 3,7-dimethyl-l-propargylxanthine(DMPX). 4-(2-[7-amino-2-(2-furyl)[l,2,4]triazolo[2,3-a][l,3,5]triazin-5-ylamino]ethyl)phenol (ZM 241385) was procured from Tocris Cookson Inc. (Ellisville, MO). Adenosine analogs were first dissolved in a small volume of dimethylsulfoxide (DMSO) (BDH, Toronto, ON) and subsequently diluted with saline or perfusate to 0.03 or 0.5% before perfusing into the stomach. The perfusion of DMSO alone, at these concentrations, did not alter basal IRG or SLI release. Five min samples were collected into ice-cold polyethylene scintillation vials containing 0.3 ml of Trasylol® (aprotinin, 10,000 KIU/ml; Miles Labs., Etobicoke, ON). For the measurement of SLI release, 0.5 ml aliquots of each sample were transferred into ice-cold test-tubes containing 0.05 ml Trasylol. For the measurement of IRG release, 1 ml aliquots of each sample were transferred into ice-cold test tubes. Aliquots of samples were frozen and stored at -20°C until assayed. 3. Radioimmunoassay (RIA) for the measurement of SLI release The RIA used to measure SLI has previously been described (Mcintosh et al;, 1978; Mcintosh et al., 1987a; Mcintosh et al., 1987b), and has been used routinely in our laboratory (Kwok et al., 1988; Kwok et al, 1990). Iodination of somatostatin: Somatostatin was iodinated using chloramine-T. In neutral buffer, chloramine-T oxidizes iodide and allows it to bind, by electrophilic substitution, to residues containing an aromatic group, such as tyrosine. Five pg of !Tyr-somatostatin (Penninsula Laboratories Inc., Belmont, CA) was dissolved in 10 jul of distilled water, followed by the addition of 10 pi 0.5 M phosphate buffer (pH 7.5). Then 10 pi of chloramine-T (2 pg/pl 0.05 M phosphate buffer, pH 7.5, EM Sciences, Darmstadt, Germany) and 1 mCi of Na[125I] (in 10 p.1 of NaOH) were added to the peptide solution. The mixture was incubated for 30 sec, and the reaction was terminated by the addition of 10 pi sodium metabisulphite (5 pg/pl 0.05 M phosphate buffer, pH 7.5, EM Sciences). One milliliter of hormone free plasma and 20 mg of QUSO G32 (Philadelphia Quartz Co., Valley Forge, PA) was then added. The mixture was vortexed, and centrifuged for 2-3 min. The supernatant (fraction A) was removed and kept. The pellet was resuspended in 1 ml of water. The mixture was again vortexed and centrifuged. The supernatant (fraction B) was removed, and the pellet was resuspended in 1 ml distilled water. The procedure was repeated. The supernatant (fraction C) was removed, and the pellet was resuspended in 1 ml of solution containing glacial acetic acid/acetone/water (1:39:40). This solution is used to elute the labeled peptide. The mixture was centrifuged for 20 min, and the supernatant (fraction D) was removed. The pellet was re-suspended in 1 ml of water (fraction E). Fractions A-E were counted, and fraction D, containing the labeled protein, was diluted to 1 x 106 cpm/10 pi in 0.1 M acetic acid containing 0.5% BSA (Miles, Kankake, IL). Aliquots of 100 pi were lyophilized, and stored at -20°C. The somatostatin label was purified on the day of the assay. The lyophilized label was re-dissolved in 2 mM ammonium acetate buffer (pH 4.6) and then applied to a column (12 cm x 1 cm) containing CM-cellulose CM 52 (Whatman Biosystem Ltd., Kent, England) which was previously equilibrated with the same buffer. The column was washed with approximately 50 ml of this buffer, before the label was eluted with 200 mM ammonium acetate buffer (pH 4.6). Buffer was introduced into the column at a rate of ~1 ml/min using a peristaltic pump. Two min fractions were collected during elution. Aliquots (10 pi) from each fraction were counted. The peak fraction was neutralized with 5 M NaOH, diluted to 3,500 ± 500 cpm/100 pi, and used for the assays. Standards: To prepare the standards, somatostatin-14 (Bachem California, Torrance, CA) was dissolved in 0.1 M acetic acid containing 0.05% BSA (Miles, Kankake, IL) to a final concentration of 5 p.g/50 (0.1. These 50 p.1 aliquots were lyophilized and stored at -20°C. On the day of the assay, the lyophilized standard was re-dissolved in 100 pi of distilled water and 400 JU.1 of assay buffer to yield a stock of 10 ug/ml. This stock was then serially diluted with assay buffer to obtain standard concentrations of: 500, 250, 125, 62.5, 31.25, 15.625, 7.8, and3.9pg/ml. Assay: Each assay tube contained 100 ul of standard or sample, 100 p.1 of label, and 200 p.1 of somatostatin antibody (ascites SOMA 03 monoclonal antibody, final dilution of 1:4,000,000). The specificity of this antibody has previously been described (Mcintosh et al, 1987b). The somatostatin antibody and label were diluted in assay buffer [23.77 mM Sodium barbital, 3.9 mM sodium acetate, 43.6 mM sodium chloride, and 0.247 mM ethylmercurithiosalicylic acid sodium salt (Eastman Kodak Co, Rochester, NY)]. The concentration of the drugs used in the present experiments did not cross-react with the antibody. Non-specific binding (NSB) tubes for standards and samples were prepared and contained buffer in place of the antibody, while zero standard tubes contained buffer in place of the standard. Total count (TC) tubes contained only 100 p.1 of labeled somatostatin. Triplicates were prepared for total count, standard, NSB standard, and zero tubes, while duplicates were prepared for the NSB sample and each unknown sample. Assay tubes were incubated for 72 h at 7°C. Free and bound labeled somatostatin were separated by adding 1 ml of dextran-coated activated charcoal [1.25% activated charcoal (Fischer Scientific, Fairlawn, NJ), 0.25% dextran T-70 (Pharmacia, Uppsala, Sweden), and 0.1% hormone free plasma in 0.05 M phosphate buffer, pH 7.5]. Tubes were incubated for 10 min, and centrifuged at 3,000 rpm for 30 min at 10°C. The supernatant was discarded and the charcoal pellet was air dried for >6 h, before the radioactivity was counted. The percentage bound of sample was calculated using the following equation: % Bound = [(TC-standard)/TC - (TC-NSB)/TC)]cpm x 100. The standard curve of the % bound vs. Log(standard concentration) was plotted using GraphPad Prism (v. 3.0, GraphPad software, San Diego, CA), and unknown somatostatin concentrations were determined by interpolation using the standard curve. Inter- and intra-assay variations were found to be <3% and <6.5%, respectively. 4. Radioimmunoassay for the measurement of IRG release IRG was measured by specific RIA as previously described (Jaffe and Walsh, 1978; Fujimiya and Kwok, 1997). Iodination of Gastrin: Gastrin was iodinated using chloramine-T. Five pg of synthetic human gastrin-1 (Penninsula Laboratories, Merseyside, England) dissolved in 10 pi 0.4 M phosphate buffer (pH 7.4) was combined with 10 pi choloramine-T, (5 pg/pl in 0.04 M phosphate buffer, pH 7.4) and 0.3 mCi Na [l25I]. After a 30 sec incubation, the reaction was terminated with the addition of 10 pi sodium metabisulphite (5 pg/pl in 0.04 M phosphate buffer, pH 7.4). The iodinated gastrin was purified using a DEAE-Sephadex A25 (Amersham Biosciences, Uppsala, Sweden) column (12 cm x 0.5 cm) equilibrated with 0.05 M imidazole buffer (pH 7.5). The label was eluted with 1 M NaCl in a 0.05 M imidazole buffer (pH 7.5) gradient at a flow rate of 1.0 ml/min. A typical elution profile for the labeled gastrin is shown in Figure 6. The four fractions, collected after the second peak, were screened by RIA. The fraction of labeled gastrin that provided the best standard curve, as indicated by measuring control samples of 50 and 100 pg/ml gastrin was further diluted and used. 1500-1 1 1 1 1 1 1 1 1 1 1 1 l Sample Fig. 6. Typical elution profile for the separation of labeled gastrin. 125I-conjugated synthetic human gastrin-I was purified on a DEAE-Sephadex A25 column. The label was eluted with 1 M NaCl in a 0.05 M imidazole buffer gradient (pH 7.5) at a flow rate of 1.0 ml/min. Fractions in the second peak have been shown to exhibit higher specific activity (Jaffe and Walsh, 1978). Thus, the four fractions collected after peak II (# 61-64) were selected for screening by radioimmunoassay. Standards: Human synthetic gastrin-1(2000 pg/ml; Penninsula Laboratories, San Carlos, CA) was serially diluted with assay buffer to obtain 1,000, 500, 250, 125, 62.5, 31.25, 15.625, 7.8, and 3.9 pg/ml standard samples. Assay: The assay buffer was composed of 0.02 M sodium barbital (pH 8.4) containing 0.5% BSA (RIA grade, Sigma-Aldrich). Each assay tube contained 700 pi assay buffer, 100 pi standard (0 to 1000 pg/ml) or sample, 100 pi gastrin antibody (PM1, rabbit polyclonal, final dilution of 1:90,000) and 100 pi labeled gastrin (-2000 counts/tube). The gastrin antibody (PM-1) was kindly provided by Dr. R. Pederson (Physiology, UBC). NSB samples contained buffer in place of the antibody, while zero standard tubes contained buffer in place of the sample, and total count tubes contained only labeled somatostatin. Triplicates were prepared for total count, standard, NSB standard, and zero tubes, while duplicates were prepared for the NSB sample, and each unknown sample. Assay tubes were incubated for 48 h at 7°C. Free and bound labeled gastrin were separated by the addition of 200 pi dextran-coated activated charcoal (1.25% activated charcoal, 0.25% dextran T-70, and 6.5% charcoal extracted plasma in 0.04 M phosphate buffer, pH 6.5). Samples were incubated for 10 min, and centrifuged at 3,000 rpm for 30 min at 10°C. The supernatant was discarded and the charcoal pellet was dried for >6 h, before the radioactivity was counted. The % bound was calculated as described for the SLI RIA. The IRG of unknown samples were determined by interpolation using the standard curve. The inter- and intra-assay variations of this assay were found to be <5.9% and <3.2%, respectively. 5. Data analysis SLI measurements: Although the basal release rate of SLI varied among animals, previous experiments have demonstrated that basal SLI release is well maintained during a 50 min perfusion period (Saffouri et al, 1980; Kwok et al, 1988, 1990). Therefore, results are expressed as mean ± SEM of SLI release (%), and calculated as follows: [SLI release (pg/min) during a 5 min period + SLI release (pg/min) during period 1] x 100. To compare the effect of various analogs, results are also expressed as percent change (SLI release), which is calculated as follows: [mean SLI release in the presence of drug - mean basal SLI release (periods 1-3)] pg/min + [mean basal SLI release (periods 1-3)] pg/min x 100. IRG measurements: Basal IRG release is also maintained in the perfused stomach preparation (Saffouri et al, 1980; Pederson et al, 1984; Kwok et al, 1990). The mean ± SEM of IRG release (%) was also calculated as described above. The percentage inhibition of IRG release by various analogs was calculated as follows: [mean basal IRG release (periods 1-3) - mean IRG release in the presence of drug] pg/min + [mean basal IRG release (periods 1-3)] pg/min x 100. Statistics: Statistical significance (P < 0.05) was determined using one-way ANOVA followed by Dunnett's multiple comparison test and the paired or unpaired Student's t-test when appropriate. Statistics were performed using GraphPad Prism, and the EC50 for different drugs were also estimated using the same software. II. Adenosine receptor mRNA: distribution, structure, and quantification 1. Distribution of gastric adenosine receptor mRNA RT-PCR was performed to determine whether the mRNA of all four adenosine receptor subtypes are expressed in various regions of the rat stomach, including the fundus, corpus, antrum, and the mucosa. 1.1 Primer design and synthesis Adenosine Ai, A2A, A2B, and A3 receptor PCR primers were designed based on previously published cDNA sequences of these receptors in the rat brain (See Table 3). Primers were designed with the assistance of Mr. Jeff Hewitt (Department of Biochemistry and Molecular Biology, UBC) using the software program PCGene. Upper and lower primer regions were selected based on several criteria. Regions with similar melting temperature (Tm) were selected. Primers had a GC content of 40-60%. Complementarity between the upper and lower primers, runs of >3 Gs or Cs at the 3' end, and the occurrence of a T at the 3' end were avoided. Primer sets, designated as short length (SL) primers, were used to amplify only a portion of the adenosine receptor cDNA. These were designed for all four adenosine receptors. Full length (FL) primers, which amplify the entire coding region of the receptors, were designed for the Ai and A2A receptors. The sequences of these primers are listed in Table 3. The primers were synthesized by the Nucleic Acids Protein Services (NAPS) Unit at UBC and were diluted in autoclaved distilled water. Since single stranded DNA at a concentration of 33 Table 3: Adenosine receptor RT-PCR primers and PCR conditions Primer Set' Accession No.2 Position (brain sequences) Primer Sequence Amplicon Length Annealing Temp. (°C)3 [MgCl2] mM4 A,SL M64299 373-392 bp 636-655 bp Upper: 5'TTC CAG GCT GCC TAC ATT GG3' Lower: 5'CAA TGG CGA GCA GAG CCA GA3' 283 bp 64 1.5 AiFL M64299 322-341 bp 1388-1407 bp Upper: 5' GTC TGC TGA TGT GCC CAG CT3' Lower: 5' AC A GGG TGG GAC AGG GAG AA 3' 1086 bp 58 1.5 A2ASL S47609 99-118 bp 520-539 bp Upper: 5' GTG GAG CTG GCC ATC GCT GT 3' Lower: 5' GCC GCA GGT CTT CGT GGA GT 3' 441 bp 60 1.5 A2AFL S47609 41-61 bp 1355-1377 Upper: 5'CTG CTG AGC CTG CCC AAG TGT3' Lower: 5'CCC TTC TCT TTG GGT TAC CCG3' 1335 bp 58 4.5 A2BSL M91466 223-242 bp 583-602 bp Upper: 5'AGA CCC CCA CCA ACT ACT TT 3' Lower: 5' GCT CTT ATT CGT GAT GCC AT 3' 380 bp 53 1.5 A3SL M94152 319-338 bp 788-807 bp Upper: 5' AAA GCC AAC AAT ACC ACG AC 3' Lower: 5' GGA GCT GTT TTG AGA GAG CT 3' 489 bp 58 3.0 Primer sets designated as "SL" (short length) span only part of the coding region of the receptor. length) span the entire coding region of the receptor. Accession number of adenosine receptor sequences in the rat brain Optimal annealing temperatures used for PCR Optimal MgCh concentration used for PCR hose designated as "FL" (full-(j,g/ml has an A260 reading of 1, concentrations were determined spectrophotometrically using the following equation: Concentration (ug/ml) = A260 x 33 pg/ml x 100 (dilution factor). 1.2 Optimization of RT-PCR conditions Several variables were examined to determine the optimal conditions for the Ai, A2A, A2B, and A3 PCR reactions, including different MgCl2 concentration (0.5 mM-7 mM) and annealing temperature during PCR (50-70°C). The optimum conditions for all RT-PCR reactions performed in this study are listed in Table 3. 1.3 Tissue extraction Male Wistar rats (200-250g) were housed in light and temperature controlled rooms, with free access to food and water. Rats were anaesthetized with Somnotol® (60 mg/kg, i.p.). The fundus, corpus, and antrum were dissected out, rinsed in sterile ice-cold saline, flash frozen in liquid nitrogen, and stored at -80°C until total RNA was extracted. The total RNA from the gastric mucosa (containing the gastrin secreting G-cells and somatostatin secreting D-cells) was obtained as follows: The whole stomach was rinsed in sterile ice-cold saline and pinned onto a cork board with the luminal surface facing up. The mucosa was gently scraped off using a sterile glass slide, and total RNA was extracted immediately, as described in the next section. The striatum was also dissected from the brain and used as a positive control since this tissue has been shown to express the mRNA of all four adenosine receptors (Dixon et al, 1996). 1.4 Total RNA extraction and quantification Total RNA was initially extracted using the acid guanidinium thiocyanate-phenol-chloroform extraction method, previously described by Chomczynski and Sacchi (Chomczynski and Sacchi, 1987). Since this method is fairly time consuming, total RNA extractions were subsequently performed using Trizol® reagent (Invitrogen, Carlsbad, CA). This reagent produced higher yields of total RNA at greater speed. The choice of extraction method did not affect RT-PCR results, and both methods are described in the next section. All materials and reagents used in the preparation of RNA and first strand cDNA were DNase- and RNase-free. Solutions were prepared using filtered distilled water which has been treated with diethylpyrocarbonate (DEPC) (Sigma-Aldrich), as previously described (Sambrooket al., 1989a). For the guanidinium fhiocyanate-phenol-chloroform method (Chomczynski and Sacchi, 1987), tissues were ground to fine particles in a sterile mortar containing liquid nitrogen. Samples were transferred to sterile tubes, and homogenized on ice in Solution D [4 M guanidinium thiocyanate, 25 mM sodium citrate (pH 7), 0.5% sarcosyl and 0.1 M 2-mercaptoethanol] (1 ml/100 mg tissue) using the Tissumizer® tissue homogenizer (Tekmar Co. Cincinnati, OH). The following reagents were then added, 2 M sodium acetate (0.1 ml/100 mg), water-saturated phenol (1 ml/100 mg), and chloroform-isoamyl alcohol [49:1] (0.2 ml/100 mg). The mixture was vortexed for 10 sec and incubated on ice for 15 min. The homogenate was centrifuged at 10,000 g for 20 min at 4°C. The aqueous phase was removed and RNA was precipitated from this phase by the addition of isopropanol (1 ml/100 mg). Samples were incubated for 1 h at -20°C, and then centrifuged at 10,000 g for 20 min at 4°C. The resulting pellet was dissolved in Solution D (0.3 ml/100 mg) at 37°C. An equal volume of isopropanol was added and the mixture was allowed to reprecipitate at -20°C for 1 h. The mixture was centrifuged at 10,000 g for 10 min at 4°C. Isopropanol was then removed and the RNA pellet was resuspended in 75% ethanol (1 ml/100 mg). After removal of the ethanol, the RNA pellet was vacuum-dried for 15 min, dissolved in DEPC-treated water (100 pl/100 mg), aliquoted, and stored at -80°C. To extract total RNA using Trizol® reagent, tissues were homogenized on ice in Trizol® (1 ml/100 mg tissue) using the Tissumizer® tissue homogenizer. Samples were incubated at room temperature for 5 min, followed by the addition of chloroform (0.2 ml/ml Trizol®). Samples were shaken vigorously by hand for 15 sec, incubated in room temperature for 3 min and then centrifuged at 12,000 g for 15 min at 4°C. The aqueous phase was removed, and the total RNA was precipitated from this phase by the addition of isopropanol (0.5 ml/ml Trizol®). Samples were incubated at room temperature for 10 min and then centrifuged at 12,000 g for 10 min at 4°C. The resulting RNA pellet was washed in 75% ethanol (lml/ml Trizol®), and centrifuged at 7,500 g for 5 min at 4°C. The ethanol was removed and the RNA pellet was air-dried for 10 min at room temperature. The total RNA was finally rediluted in DEPC-treated water (100 pl/ml Trizol®), aliquoted, and stored at -80°C. RNA purity and concentration: The total RNA purity was determined by measuring the absorbance at 260 nm (A260) and 280 nm (A280), while the concentration was estimated by measuring the A260- The spectrophotometer was calibrated with DEPC-treated water, and total RNA samples were diluted 100-fold in DEPC-treated water prior to measurement. Pure RNA gives an A26o/A28o ratio of 2 (Chomczynski and Sacchi, 1987). The A260/A280 ratio for all our samples exceeded 1.8. Since total RNA at a concentration of 40 pg/ml has an A260 of 1, total RNA concentrations were determined using the following equation: concentration (pg/ml) = A260 x 40 pg/ml x 100 (dilution factor). 1.5 Formaldehyde gel electrophoresis The integrity of the RNA samples was examined by formaldehyde gel electrophoresis to ensure that the samples were not degraded during total RNA extraction. Formaldehyde gel electrophoresis was performed using a method modified from the Molecular Cloning Laboratory Manual (Sambrook et al, 1989a). RNA samples were prepared on ice by mixing 10 pg of total RNA (in a volume of 4.5 pi) with 10 pi deionized formamide and 2 pi 5x formaldehyde gel running buffer [0.1 M MOPS, 40 mM sodium acetate, and 5 mM EDTA]. The mixture was incubated for 15 min at 55°C. This was followed by the addition of 1 pi ethidium bromide (1 mg/ml) and 2 pi formaldehyde gel loading buffer [50% glycerol, 1 mM EDTA (pH 8.0), 0.25 % bromphenol blue, 0.25% xylene cyanol]. Samples were electrophoresed through a 1% (w/v) agarose gel containing 5x formaldehyde gel running buffer and formaldehyde in a final concentration of lx and 2.2 M, respectively. The gel was submerged in lx gel loading buffer. The samples were loaded along with a 0.24-9.5 kb RNA ladder (Invitrogen), and run at 100 V for 1 h. Gels were viewed under UV light using the Stratagene Eagle Eye II system (La Jolla, CA). Images were acquired and saved using the Eagle Sight image analysis software (version 3.1, 1997, Stratagene). 1.6 DNase I treatment RNA samples were treated with DNase I (Amersham Pharmacia, Piscataway, NJ) to remove any residual DNA contamination that may remain in samples after total RNA extraction. DNase I treatment was performed at room temperature in lx first strand buffer [50 mM Tris-HCl (pH 8.3 at 25°C), 75 mM KC1, 3 mM MgCl2] containing 1 U DNase I/pg total RNA. The reaction was allowed to proceed for 15 min, before 1 pi of 25 mM EDTA was added to stop the reaction (Sanyal et al., 1997). Samples were then incubated for an additional 10minat65°C. 1.7 First strand cDNA synthesis First strand cDNA was synthesized from 5 pg of DNase I treated total RNA using Superscript II RNase H- Reverse Transcriptase (Invitrogen). Total RNA (5 pg) was reverse transcribed in lx first strand buffer [50 mM Tris-HCl (pH 8.3 at 25°C), 75 mM KC1, 3 mM MgCl2] containing 200 ng random hexamers, 10 mM DTT, 0.5 mM dNTP mix, and 200 U Superscript II RNase H- Reverse Transcriptase, in a total volume of 20 pi. As a negative control for PCR, a sample was prepared using autoclaved distilled water in place of total RNA. 1.8 PCR The PCR reaction mixture for all PCR reactions consisted of 2 pi cDNA in 1 x PCR buffer [20 mM Tris-HCl (pH 8.4), 50 mM KC1], containing 0.2 mM dNTP mix, 1.5-4.5 mM MgCl2 (see Table 3 for MgCl2 concentrations), 100 ng forward primer, 100 ng reverse primer (see Table 3 for primer sequences), and 1 U Platinum Taq DNA polymerase (Invitrogen), in a total volume of 50 pi. For each experiment, a positive control sample containing striatal cDNA, and a negative control sample were included. The PCR was performed as follows using the Robocycler Temperature cycler (Stratagene, La Jolla, CA): cDNA was completely denatured (2 min at 94°C) during the first step of PCR. Samples were then subjected to 30 cycles of amplification. Each cycle consisted of a 45 sec denaturation period at 94°C, a 1 min annealing period at 53°C - 60°C (See Table 3), and a 1 min extension period at 72°C. The sample was then subjected to a final extension period for 5 min at 72°C. PCR products were visualized by gel electrophoresis as described below. 1.9 Agarose gel electrophoresis To determine the size of the amplicons produced by PCR, agarose gel electrophoresis was performed as previously described (Sambrook et al, 1989b). The PCR products (10 pi) were mixed with 3 pi gel loading dye [0.25% bromphenol blue in 50% glycerol, 10 mM Tris, 1 mM EDTA, pH 8], and loaded on a 2.5% agarose gel (w/v) containing lx Tris acetate EDTA (TAE) buffer [40 mM Tris acetate and 1 mM EDTA (pH 8.5)] and ethidium bromide (0.5 pg/ml). The gel was placed in lx TAE buffer and run at 100 V for 45 min. Bands were visualized and photographed under UV light using the Stratagene Eagle Eye II System and the Eagle Sight Image Analysis software program. 1.10 Restriction enzyme digestion To confirm the identities of the PCR amplicons, restriction enzyme digestion was performed. PCR products generated by the Ai, A2A, A2B and A3 RT-PCR reactions were concentrated, and cleaved with restrictive enzymes that were selected based on their cleavage sites. Cleavage sites within the PCR amplicons were determined using PCGENE. The following restriction enzymes were used to cleave the following PCR amplicons: Nar I and Sac I for the A, receptor; EcoRI, Mbo I, Pst I, and Rsa I for the A2A receptor; Ava II and Hpa II for the A2B receptor; Ava II and Hinf I for the A3 receptor. All restriction enzymes were purchased from Amersham Pharmacia (Uppsala, Sweden). To concentrate the DNA, PCR products (300 pi) were mixed with 150 pi of 7.5 M ammonium acetate and 600 pi 95% ethanol. Samples were incubated for 1 h at -20°C and then centrifuged at 10,000 g for 20 min at 4°C. The resulting pellet was washed in 1 ml 70% ethanol, and centrifuged at 10,000 g for 5 min at 4°C. The ethanol was removed and the pellet was re-dissolved in 50 pi of autoclaved distilled water. The restriction enzyme digestion was performed using the buffer provided with the enzyme, according to the manufacturer's instructions. For these reactions, 5 pi of the concentrated PCR product was digested in a final volume of 10 pi. Samples were electrophoresed through a 1.5-2 % agarose gel along with a 1 kb ladder (Invitrogen), as described earlier. The Eagle Sight Image analysis software was used to determine the size of the fragments. This measurement is based on the rate of DNA migration through the agarose gel. The software produces a standard curve of migration distance vs. Log[fragment size (bp)] using the fragments of the 1 kb ladder. The size of the fragments generated by restriction enzyme digestion was interpolated by comparing the migration distance of the fragment against the standard curve. 2. Structure of the adenosine receptors To determine if the gastric Ai and A2A receptors undergo alternative splicing and if the structure of these receptors are the same as those in the brain, the entire coding region of the gastric Ai and A2A receptors were amplified by RT-PCR, cloned, and sequenced. In addition, part of the gastric A2B and A3 receptor gene sequence was also determined. These A2B and A3 gene sequences are located between the short length upper and lower primers listed in Table 3. This portion of the A3 receptor gene is of particular interest since its sequence can reveal whether the A3i variant of the A3 receptor is expressed in the stomach. 2.1 Detection of Ai and A2A receptor coding regions by RT-PCR Primers: The primers used for this experiment are listed in Table 3. The AiFL and A2AFL primers span the entire coding region of the Ai and A2A receptor, respectively. RT-PCR: To examine whether alternative splicing occurs for the Ai and A2A receptors, RT-PCR was performed in all gastric tissues, including the fundus, corpus, antrum and mucosa using the AiFL and A2AFL primers. RT-PCR was performed as previously described using the PCR conditions listed in Table 3. Fundus, corpus, antrum and mucosa RNA was used as the template for these reactions. A sample containing striatal RNA and a sample containing no template was also included as positive and negative controls, respectively. 2.2 Cloning and DNA sequence analysis of the adenosine receptors PCR products and purification: RT-PCR was performed for all four receptors using rat gastric mucosal RNA as the template and the AiFL, A2AFL, A2BSL, and A3SL primers. The PCR products were concentrated using the QIAquick PCR purification kit (Qiagen, Mississauga, ON). According to manufacturer's instructions, 1.5 ml Buffer PB was mixed with 300 pi of the PCR product. The DNA was bound to a QIAquick column by centrifugation and washed with PE buffer. The DNA was finally eluted with 50 pi of autoclaved distilled water. The purified DNA was electrophoresed through a 2% agarose gel, as described earlier, and extracted from the gel using the QIAquick gel extraction kit (Qiagen), according to the manufacturer's instructions. The fragment was excised from the gel using a sterile razor blade and weighed. Buffer QX1 (300 jal/100 mg gel) was added, and the mixture was vortexed for 30 sec. QIAEX II buffer (5 pi) was then added and the sample was incubated at 50°C for 10 min, and vortexed every 2 min, and the sample was then centrifuged at 10,000 g for 30 sec. The pellet was washed with Buffer QX1 (500 pi), followed by two washes in Buffer PE (500 pi). The pellet was air dried for 10-15 min, and rediluted in autoclaved distilled water. Ligation and transformation: The purified adenosine receptor transcripts were ligated into the pGEM-T vector (Promega, Madison, WI) by combining 5 pi of the purified A,FL, A2AFL, A2BSL or A3SL PCR product with 1 pi lOx T4 DNA Ligase Buffer, 1 pi pGEM-T vector (50 ng), 1 pi T4 DNA ligase and 2 pi autoclaved distilled water. The mixture was incubated overnight at 4°C, and then heated to 70°C for 10 min, and cooled to room temperature. DHot-competent E. coli cells (Invitrogen) were transformed with the ligated PCR product/pGEM-T vector by combining 2 pi of this sample with 50 pi of cells. The mixture was incubated on ice for 20 min, heat shocked at 37°C for 50 sec, and placed on ice. The volume of Luria-Bertani (LB) broth [0.1% NaCl containing tryptone (lOg/L) and yeast extract (5g/L)] added to cells depended on the size of the PCR products, which were transformed into the cells. Cells containing the A[FL, A2AFL, A2BSL, and A3SL transcript were combined with 900 pi, 1 ml, 1.5 ml, and 1.5 ml of LB, respectively. Cells were incubated for 1 h at 37°C, and then plated on LB plates containing ampicillin (100 pg/ml), isopropyl P-D-thiogalactopyranoside (0.5 mM) and 5-bromo-4-chloro-3-indolyl-B-D-galactoside (80 pg/ml). Cells were grown overnight at 37°C. Colonies containing inserts appear white since the inserts interrupts the lacZ gene in the pGEM-T vector. White colonies were screened by RT-PCR. Colonies containing the correct insert were grown in 5 ml of LB broth containing ampicillin (100 pg/ml). Plasmid DNA was purified from these cells using the QIAprep Miniprep kit (Qiagen). The sample was centrifuged at 3,000 rpm for 5 min at 4°C to pellet the cells. Cells were resuspended in 250 pi Buffer PI, followed by the addition of 250 pi of Buffer P2. Cells were allowed to lyse for 5 min prior to the addition of 350 pi N3 Buffer. The sample was then centrifuged at 10,000 g for 10 min at 4°C. The supernatant was applied to a QIAprep column and centrifuged for 30 sec. The column was washed with 750 pi PE buffer, and plasmid DNA was eluted from the column with 50 pi of Buffer EB [10 mM Tris-Cl (pH 8.5)] DNA sequence analysis: The DNA sequences were determined at the NAPS Unit (UBC) using both the T7 and SP6 primers. This procedure was successful in obtaining the AiFl, A2BSL and A3SL sequences. However, due to the length of the A2AFL sequence, 2 additional primers located within the A2AFL transcript were required to complete the sequence analysis. These primers were based on the rat brain A2A receptor gene (accession #S47609). Their sequences are as follows: 5' TTG TCC TGG TCC TCA CGC 3' (position 313-330 bp) and 5' AGG GCC GGG TGA CCT GTC 3' (position 541-558 bp). The sequence of the rat brain Ai and A2A receptor coding regions, and the partial cDNA sequence of the A2B and A3 receptors were aligned with the published sequences of these receptors in the rat brain using the "University of Southern California (USC) Sequence Alignment Server" (www-hto.usc.edu/software). These sequences were submitted to GenBank, and assigned accession numbers (see Results). 3. Quantification of gastric Ai and A2A receptor gene expression 3.1 Quantification by competitive RT-PCR Competitive RT-PCR was performed to quantify the level of adenosine A\ and A2A receptor gene expression in various regions of the stomach, including the fundus, corpus, antrum, and mucosa. The general steps for measuring Ai and A2A receptor gene expression using competitive RT-PCR are as follows: 1) synthesize, purify, and quantify the Ai and A2A receptor RNA competitors; 2) perform the competitive RT-PCR assay by preparing samples containing a fixed amount of tissue total RNA and various amounts of competitor RNA as the template; 3) separate and visualize the PCR products by agarose gel electrophoresis; 4) determine the relative intensities of the tissue template and competitor bands; 5) calculate the tissue template level based on the relative intensity of the competitor vs. the template band. The intensities of both bands are approximately equal when the concentration of the template is equal to the concentration of the competitor. Each step is described in detail below. 3.1.1 RNA competitor design and synthesis Previous experiments have shown that the reverse transcription step can vary considerably between assays (Ferre, 1992). Thus, to control for the efficiency of the reverse transcription step, RNA competitors were synthesized for this experiment instead of DNA competitors. The RNA competitors were synthesized by deleting a small portion (< 15%) of the adenosine receptor transcripts. The deletion allows for differentiation between the template and competitor bands when PCR products are visualized by electrophoresis. The small deletion also controls for differences in PCR amplification efficiency that may occur due to differences in RNA length and secondary structure. The adenosine receptor RNA competitors were synthesized as described in the following section and in Figure 7. The general method has previously been described by Tang et al. (1996). Vectors containing the Ai and A2A competitor: To synthesize vectors containing the A] receptor competitor, RT-PCR was performed using the AjFL primer set (see Table 3). The RT-PCR products were concentrated using the QIAquick PCR purification kit, and gel purified using the QIAquick gel extraction kit, as described earlier. The Ai competitor was synthesized by deleting a 136 bp fragment from the Ai receptor template to produce a competitor containing 950 bp. To produce this deletion, the Ai template was digested with Bpm I (New England BioLabs, Beverly, MA), as directed by the manufacturer. Three fragments (515 bp, 435 bp and 136 bp) were formed. The sample was electrophoresed through a 2% agarose gel as previously described. The two fragments containing the 5' and 3' primer sites (515 bp and 435 bp) were excised from the gel, purified using the QIAquick gel extraction kit, and religated with the pGEM-T vector using T4 ligase. The A2A competitor was synthesized in a similar manner. RT-PCR was performed using the A2ASL primers (see Table 3). The A2A competitor was synthesized by deleting a 57 bp fragment from the A2A template to produce a 384 bp competitor. The A2A template was digested with Tsp509 I (New England BioLabs), according to manufacturer's instructions, to produce 3 fragments (286 bp, 98 bp, and 57 bp). The two fragments containing the primer sites (286 bp and 98 bp) were gel-purified and religated with the pGEM-T vector. Transformation and in vitro Transcription: Vectors containing the modified Ai, and A2A inserts were transformed into DHa competent cells, as described earlier. Bacterial colonies were screened by PCR to ensure that the colonies that were chosen contain the Cleavage */ sites ^ 5' primer 3' primer 1. Digest with enzyme which cleaves -100 bp apart / 1 \ 5' primer A discard B 2. separate fragments and 3 primer purity by gel electrophoresis 5' primer A B 3' primer 3. Re-ligate A+B 1 4. Clone modified adenosine receptor mRNA 5. Linearize and purify plasmid containing the modified adenosine receptor transcript 6. Synthesize RNA competitors by in vitro transcription, purify, and quantify Fig. 7. Design and synthesis of the adenosine A, and A2A receptor RNA competitors. A) The general protocol used for the design and synthesis of A, and A2A competitors. B) Agarose gels showing PCR amplicons generated using corpus tissue total RNA as the template (lanes 2; A,, 1086 bp; A2A, 441 bp) or A, and A2A RNA competitor as the template (lanes 3; A,, 950 bp; A2A, 384 bp). A 100 kb ladder is shown on lanes 1. The competitor and tissue template bands can be differentiated easily by their size. modified adenosine receptor transcripts. The modified Ai and A2A receptor transcripts generated PCR amplicons which were 950 and 382 bp in length, respectively (see Fig. 7B). Plasmids from the chosen colonies were isolated using the QIAprep Miniprep kit and linearized by Not I (Amersham Pharmacia) digestion, according to manufacturer's instructions. Complete digestion was confirmed by agarose gel electrophoresis. The linearized plasmids were then purified and concentrated as described earlier. In vitro transcription was performed using the Riboprobe in vitro Transcription System - T7 (Promega). To synthesize the competitor RNA, a reaction mixture (20 pi) containing lx Transcription Optimized buffer [40 mM Tris-HCl (pH 7.9), 6 mM MgCl2, 2 mM spermidine, 10 mM NaCl], 10 mM DTT, 20 U RNasin ribonuclease inhibitor, 0.5 mM rNTPs, 0.2 pg linearized template, and 20 U T7 RNA polymerase, was incubated at 37°C for 1 h. The template DNA was digested with 0.2 U of RQ1 RNase-free DNase. The RNA competitors were purified using the RNeasy mini kit (Qiagen). The volume of the sample was increased to 100 pi with DEPC-treated water. Buffer RLT (350 pi) and ethanol (250 pi) was then added. The sample was applied to a RNeasy mini spin column and centrifuged for 15 sec at 8,000 g. The column was washed twice with 500 pi of Buffer RPE, and the RNA competitors were eluted from the spin column with 30 pi of DEPC-treated water. The purity and concentration of the RNA competitors were determined spectrophotometrically, as described earlier. The integrity of the competitors was examined by gel electrophoresis through a formaldehyde gel. RNA competitors were then serially diluted from 0.1 pmol to 0.01 amol, aliquoted, and stored at -80°C. RNA competitors were thawed only once prior to use to avoid damage caused by multiple freeze-thaw cycles. 3.1.2 Competitive RT-PCR assay Total RNA extraction: Tissues were dissected from various regions of the stomach, including the fundus, corpus, antrum and mucosa. As a positive control, the level of Ai and A2A receptor gene expression was also examined in the striatum. This tissue has been shown to express extremely high levels of A2A receptor mRNA and relatively high levels of At receptor mRNA (Dixon et al, 1996). Total RNA was extracted using Trizol® Reagent, and the purity and concentration of the RNA were determined by measuring the absorbance of A260/A280, and A26o, as described earlier. The A260/A280 ratio for all samples exceeded 1.8. First strand cDNA synthesis: First strand cDNA was synthesized using Superscript II as described earlier with the following modification. Various amounts of At or A2A receptor competitor RNA (from 0.01 amol to 1 pmol) and a fixed amount of total RNA (2.5 pg) were used as the template for the reaction. The cDNA that was synthesized was then used as the template for PCR. PCR: PCR was performed using the AiFL and A2ASL primer sets listed in Table 3. The Ai and A2A receptor competitive PCR reactions mixture (50 pi total) contained 0.5 pi and 2.0 pi of cDNA, respectively, in lx PCR buffer [20 mM Tris-HCl (pH 8.4), 50 mM KC1] with 0.2 mM dNTP mix, 1.5-4.5 mM MgCl2 (see Table 3), 100 ng forward primer, 100 ng reverse primer, and 1 U Platinum Taq DNA Polymerase. PCR was performed using the Robocycler Temperature cycler as described previously. However, the cycles of amplification were limited to 30 cycles to avoid reaching the plateau phase. Reaching this phase of the reaction has previously been shown to introduce errors into the RNA measurements (Wang et al, 1989; Freeman et al, 1999). The PCR products, containing the competitor and template amplicons, were separated by electrophoresis through a 2.5% agarose gel containing ethidium bromide, as described earlier. Bands were visualized and photographed under UV light using the Eagle Eye II System. To determine whether altering the initial amount of cDNA in the PCR mixture affects the final A\ and A2A mRNA measurements, an equal volume of undiluted, 2x , 4x, or 8x diluted cDNA was used as the template for these PCR reactions. 3.1.3 Competitive RT-PCR data analysis During the acquisition of gel images, special care was taken not to allow any bands to reach saturation density. The saturation of the bands can be detected by the Eagle Sight Image analysis software program. The images were saved, and band densities were determined by densiometric analysis using the Eagle Sight image analysis software (see Figure 8A). Analysis of competitive RT-PCR images has previously been described (Piatak et al., 1993; Auboeuf and Vidal, 1997). The competitor band densities were corrected for inherent variation in band density resulting from the difference between template and competitor PCR product length. The correction factor is determined by dividing the size of the tissue template amplicon by the size of the competitor amplicon. Competitor band densities, therefore, were multiplied by the following correction factors: 1.14 (1086 bp/950 bp) for the Ai competitor and 1.15 (441 bp/383 bp) for the A2A receptor competitor. Levels of Ai and A2A receptor mRNA in each tissue sample were determined by plotting the Log [density ratio (corrected competitor band/template band)] against the Log (initial competitor concentration). Since the amount of Ai and A2A template mRNA is equal to the initial Template band density -A, Template band " A: Competitor band-Competitor -band density Initial competitor concentration (amol) 25 10 5 2.5 1 42544 65255 69654 61349 71169 111215 62938 27463 19595 5594 2.99 1.10 0.45 0.36 0.09 Corrected density ratio: competitor/template B 0) *•> ro a. E CD o. E o o CO i-'35 c •a o 0.6 0.4 0.2 0 -0.2 -0.4 -0.6 -0.8 -1 -1.2 - y = 1.04x -1 >^ 0 0.25 0.5 0.75/^ m >r Log [tis • 1 1.25 1 sue A1 concentration] R2 = 0.98 Log [initial A1 competitor concentration (amol)] Fig. 8. Analysis of competitive RT-PCR data. A) Results of an A, receptor competitive RT-PCR assay performed using various amounts of Kx competitor and a fixed amount (2.5 pg) of corpus tissue total RNA. Band densities were acquired using the Eagle Sight software. The competitor band density was multiplied by a correction factor to account for the difference in band density resulting from the smaller size of the competitor (see Methods). B) The Log density ratio (competitor band/ template band) was plotted against the Log (initial competitor concentration). The Log density ratio of the (competitor band/template band) is equal to zero when the template and corrected competitor band densities are equivalent. Thus, the At receptor mRNA concentration in the tissue is determined using the x-intercept of this graph. amount of competitor RNA when the template and corrected competitor band are of equal density, the initial amount of tissue Ai and A2A template was determined at the x-intercept (See Figure 8B). Data were expressed in units of amol of Aj or A2A receptor mRNA/pg total RNA. These values were then converted to the number of copies of Ai or A2A receptor mRNA transcript/pg total RNA. Statistical significance was determined using Student's unpaired t-test, and performed using GraphPad Prism; P < 0.05 was considered significant. 3.2 Quantification by Real-Time RT-PCR The gene expression of the Ai and A2A receptor were also quantified using Real-Time RT-PCR when the ABI Prism 7700 Sequence Detection System (Applied Biosystems, Foster City, CA) became available. Absolute quantification of gastric Ai and A2A receptor gene expression was measured using a standard curve generated by known amounts of Ai and A2A receptor mRNA. Standard curve samples were prepared using in vitro transcribed Ai or A2A receptor RNA. A two-step Real-Time PCR assay was performed. This technique is described in Fig. 9, and is based on the 5' nuclease activity of Taq DNA polymerase (Heid et al., 1996; Lockey et al., 1998). A double-labeled fluorogenic probe containing a reporter dye such as 6-carboxyfluorescein (FAM) and a quencher dye such as 6-carboxytetramethylrhodamine (TAMRA) is included in each PCR reaction. The probe hybridizes to the template at a position flanked by the forward and reverse primers. During PCR amplification, the hybridized probe is cleaved by the 5' nuclease activity of Taq DNA polymerase. When the Fig. 9. Cleavage of the fluorogenic probe during Real-Time RT-PCR. During the extension phase of the Real-Time RT-PCR reaction, the primers and probes are bound to the template DNA. Taq polymerase synthesizes the new DNA strand (A), and encounters the probe (B). As Taq polymerase continues to synthesize the new strand, the 5' to 3' exonuclease activity of this enzyme degrades the probe. When the probe is intact, the energy emitted by the reporter dye, FAM is absorbed by the quencher dye, TAMRA, due to their close proximity. After the probe is degraded, FAM and TAMRA separate and the energy emitted by FAM is no longer quenched by TAMRA (C) and is detected by the ABI prism 7700 Sequence Detection System. probe is intact, the quencher dye will suppress reporter fluorescence due to Forster type energy transfer. Upon cleavage of the probe, reporter emissions are no longer quenched and are detected by the ABI Prism Sequence Detector 7700. The methods used to synthesize the Ai and A2A receptor primers and probes are described in detail below. 3.2.1 Design and synthesis of primers and probes Primers and probes were designed using the software program Primer Express (version 1.0, Applied Biosystems) according to the following guidelines. The guanine (G) and cytosine (C) content of the primers and probes were kept between 30-80% and > 4 runs of an identical nucleotide were avoided. The melting temperatures (Tra) of the primers were kept between 58-60°C, and each primer contained < 2 G/C bases at the 3' end. The Tm of the probes was 10°C higher than that of the primers. Probe sequences contained more C than G, and no G at the 5' end. For both probes, the reporter dye, FAM was linked to the 5' end and the quencher dye, TAMRA was linked to the 3' end. The fluorogenic probes were synthesized by Synthegen, LLC (Houston, Texas) and the primers were synthesized by the NAPS Unit at UBC. The sequences of the Ai and A2A primers and probes are listed in Table 4. 3.2.2 Synthesis of A! and A2A receptor RNA standards RNA transcripts expressing the entire coding region of the adenosine Aj and A2A receptors were used as the standards for quantification during Real-Time RT-PCR. To synthesize these standards, plasmids containing the AiFL and A2AFL transcripts were used. Table 4: Ai and A2A receptor Real-Time RT-PCR assay: primers and probes Rat Ai Adenosine Receptor Position Length Tm %GC Sequence Forward 963 22 59 59 5' CGGTGACCCCCAGAAGTACTAC 3' Probe 989 26 70 58 5' CAGCGACTTGGCGATCTTCAGCTCCT 3' Reverse 1039 21 59 52 5' GGGCAAAGAGGAAGAGGATGA3' Amplicon 962-1039 77 84 60 Rat A2A Adenosine Receptor2 Position Length Tm %GC Sequence Forward 910 22 58 55 5' ACCCCTTCATCTACGCCTACAG 3' Probe 936 22 69 68 5' CGGGAGTTCCGCCAGACCTTCC 3' Reverse 978 20 59 55 5' CGTGGGTTCGGATGATCTTC 3' Amplicon 910-978 69 84 59 Based on Ai receptor sequence in the rat brain; Accession number: M64299 Based on A2A receptor sequence in the rat brain; Accession number: S47609 Plasmids were generated by our previous cloning experiments, described in section 2.2. The orientation of the adenosine Ai and A2A receptor insertions in the plasmid was determined to examine whether the 5' or 3' strand of the insertion would be synthesized by T7 polymerase during in vitro transcription. To determine the orientation of the A\ and A2A insertions, a Sac I digestion and Nco I digestion were performed, respectively. These restriction enzymes were chosen for the position of their cleavage sites, which allows the differentiation between the two orientations. Only the plasmids with insertions in Orientation I (see Fig. 10) were selected. In this orientation, T7 RNA polymerase synthesizes the 5' strand of the cloned inserts during in vitro transcription. The selected plasmids were linearized by Not I digestion. This enzyme does not cleave the Ai or A2A insert, but cleaves at a site located after the cloned insert. The linearized plasmids were purified using the Qiagen miniprep kit, as described earlier. Run-off transcripts were synthesized using the Riboprobe in vitro transcription kit and T7 RNA polymerase, as described previously in section III. RNA standards were treated with DNase I, and then purified using the Qiagen RNeasy mini Kit. The quality of the RNA standards was examined by gel electrophoresis, as described earlier. The concentrations of the RNA standards were estimated spectrophotometrically, and then accurately determined using the RiboGreen Reagent Quantitation kit (Molecular Probes, Eugene, OR). Serial dilutions of the Ai and A2A RNA standards were made. RNA concentrations were determined in lx TE buffer [10 mM Tris-HCl, 1 mM EDTA (pH 7.5) in DEPC-treated water] containing RiboGreen RNA quantitation reagent (0.025 pi /ml), a sensitive fluorescent nucleic acid stain. When bound to RNA, RiboGreen reagent has an Sac I (Pos. 63 of A, insert in Orientation I) A, Insertion (1086 bp) Sac I (Pos. 63 of A, insert in Orientation II) Sac I (Pos. 94 of pGEM-T vector) pGEM-T Vector (3003 bp) B. Orientation I Orientation II Strand synthesized by T7 Polymerase 5' strand 3' strand Molecular Weight of Strand synthesized 368.11 kD 373.44 kD Sac I Cleavage Sites 1 &3 2&3 Fragments generated (bp) 3270 & 818 3737 & 351 Nco I (Pos. 73 of AM insert in Orientation I) Au Insertion (1335 bp) Nco I (Pos. 73 of A^ insert in Orientation II) Nco I (Pos. 37 of pGEM-T vector) pGEM-T Vector (3003 bp) Orientation I Orientation II Strand synthesized by T7 Polymerase 5' strand 3' strand Molecular Weight of Strand synthesized 450.16 kD 453.10 kD Nco I Cleavage Sites 1 & 3 2 & 3 Fragments generated (bp) 4291 & 47 3019 & 1319 Fig. 10. Orientation of A, and A2A receptor transcript insertion into pGEM-T vector. Plasmids containing the A, (A) and A2A (B) transcripts were cleaved with restriction enzymes, Sac I and Nco I, respectively. When the transcript is inserted in one orientation (orientation I), plasmids were cleaved at sites 1 and 3, and when inserted in the other orientation (orientation II), plasmids were cleaved at sites 2 and 3. The size of the fragments generated by restriction enzyme digestion are shown in the tables. excitation wavelength of 480 nm and emission wavelength of 520 nm. The A\ and A2A RNA standards were quantified by measuring the fluorescence against a standard curve composed of samples containing known amounts of ribosomal RNA standard (16S and 23S rRNA from E. coli cells; provided by the manufacturer). The fluorescence measurements were performed using the FL600 Microplate Fluorescence reader (Biotek Inc., Winooski, VT), and RNA concentrations (ng/ml) were determined using the KC4 Kineticalc Software (version 2.6, Biotek Inc.). RNA concentrations were converted from ng/ml to copies/ml using the molecular weight of the standards: Ai RNA standard: 368.11 kD; A2A RNA standard: 450.16 kD. RNA standards were serially diluted to 103 - 1012 copies/pl in RNase-free water, aliquoted, and stored at -80°C. Standards were thawed only once prior to use. 3.2.3 Two step Real-Time RT-PCR assay Real-Time RT-PCR was performed to measure the level of adenosine Ai and A2A receptor mRNA in the fundus, corpus, antrum, mucosa and striatum. Total RNA was extracted using Trizol® reagent, as described in section I, and quantified using the RiboGreen RNA Quantitation Kit. Total RNA was treated with DNase I. Two-step Real-Time PCR was performed using Superscript II RNase H-Reverse transcriptase, and the TaqMan PCR Core Reagents Kit (Applied Biosystems, Foster City, CA) as described below. Step 1 - Reverse transcription: According to the manufacturer's instructions, 1 pg of DNase I-treated tissue RNA was reverse transcribed in a total volume of 10 pi containing 200 ng random hexamers, 20 U RNAguard RNase inhibitor, lx first strand buffer, 10 mM DTT, 0.5 mM dNTP mix, and 100 U Superscript II RNase H- Reverse Transcriptase. Various amounts of RNA standard and a negative control sample containing DNase I-treated RNase-free water in place of template, were reverse transcribed simultaneously. The latter was subsequently used as the template for the negative control during Real-Time PCR. Step 2 - PCR: The standard curve for each Real-Time PCR assay was composed of at least 6 concentrations of either the Ai or A2A receptor standard. Each assay also contained a negative control sample containing no template. All reactions were performed in triplicate. The optimization of the Ai and A2A Real-Time RT-PCR assay conditions are described in the following section (3.2.5.). The PCR reaction consisted of a final volume of 25 pi reaction mixture containing lx TaqMan Buffer A, 200 pM of each dATP, dCTP, and dGTP, 400 pM dUTP, 0.01 U/pl AmpErase uracil-N-glycosylase (UNG) and 0.025 U/pl AmpliTaq Gold DNA Polymerase from the TaqMan PCR Core Kit. This reaction mixture also contained 0.5 pi of tissue or standard cDNA, or 0.5 pi of the control sample (which does not contain template), 100 nM probe, various concentrations of forward and reverse primer (Ai receptor assay contains 300 nM of each, A2A receptor assay contains 100 nM of each) and MgCl2 (Ai receptor assay contains 7.5 mM, A2A receptor assay contains 4.5 mM). The reaction was performed using the ABI Prism 7700 Sequence Detector (Applied Biosystems) with the following cycling parameters: a 2 min hold at 50°C for UNG incubation, a 10 min hold at 95°C for AmpliTaq Gold activation, followed by 40 cycles of amplification consisting of a 15 s denaturation step at 95°C and a 1 min anneal/extend period at 60°C. 3.2.4 Real-Time RT-PCR data collection and analysis The data was collected by the Sequence Detection Software (version 1.6.3, Applied Biosystems) during each cycle of PCR amplification. At the end of the PCR, the data was analyzed by the software program, and an amplification plot was generated. The amplification plot shows the normalized reporter emissions (Rn) vs. cycle number. Reporter emissions were normalized with the passive reference dye, ROX, which was present in the TaqMan Buffer A. The threshold cycle (Cr), where the software detects an increase in fluorescence associated with exponential growth, was determined using the first 15 cycles to represent the baseline. The threshold cycle (Cr) of each reaction occurred where the Rn exceeded the baseline Rn by the value of the threshold. The average Rn and standard deviation of Rn in the first 15 cycles were determined. To ensure that the change in Rn is associated with exponential growth of the amplicon during quantification, the threshold was determined by multiplying the standard deviation of the first 15 cycles by a factor of 10. To quantify the levels of adenosine Ai and A2A receptor mRNA, standard curves of CT VS. Log (initial Ai or A2A receptor RNA standard concentrations) were generated. The initial concentration of each unknown sample was determined by interpolation using the CT value determined by the assay. The correlation coefficient of each standard curve was >0.95. The threshold cycle of the control samples containing no template exceeded 40 cycles in every assay, indicating the absence of DNA contamination. For the absolute quantification of adenosine Ai and A2A receptor mRNA levels, measurements were not normalized using the housekeeping genes, and the results were expressed as copies of mRNA per pg total RNA. Statistical significance was determined using GraphPad Prism and the two-tailed unpaired Student's t-test; P < 0.05 was considered significant. 3.2.5 Optimization of Real-Time RT-PCR Real-Time RT-PCR assays were optimized by adjusting the primer, probe, and MgCl2 concentrations. RT-PCR samples containing 100, 200, or 300 nM of probe and either 100, 200, or 300 nM of primer were prepared. Ai or A2A RNA (1x10s copies) was used as the template for these reactions. The conditions resulting in a fairly low CT were selected for Real-Time RT-PCR. Due to the expense of the probe, the lowest acceptable concentration of this component was selected (see Table 5). Various MgCl2 concentrations (3.5, 4.5, 5.5, 6.5, and 7.5 nM) were then subsequently tested using these primer and probe concentrations (see Table 6). The following optimized Ai and A2A Real-Time RT-PCR assay conditions were established. The Ai receptor assay contained 100 nM probe, 300 nM primers, and 7.5 nM MgCl2, while the A2A receptor assay contained 100 nM probe, 100 nM primers, and 4.5 nM MgCl2. III. Regulation of adenosine receptor gene expression and function in altered physiological states of the stomach The effect of fasting and omeprazole treatment on Ai and A2A receptor mRNA levels was examined to determine if changes in the physiological state of the stomach alter adenosine receptor gene expression. The gene expression of gastrin and somatostatin was also measured to determine if changes in adenosine receptor gene expression occurred with changes in gastric peptide expression. The effect of omeprazole on adenosine agonist-induced changes in IRG and SLI release was also examined to further determine if changes in adenosine receptor gene expression may result in changes in adenosine receptor function. Table 5: Optimization of primer and probe concentrations for Real-Time RT-PCR Probe Concentration (nM) Primer Concentration (nM) Ai Assay: Cj A2A Assay: CT' 100 40.0 29.1 100 200 40.0 29.4 300 35.5 30.1 100 40.0 28.9 200 200 35.1 29.1 300 40.0 29.9 100 40.0 29.5 300 200 40.0 29.2 300 40.0 29.1 1 The threshold cycle (Cj) was obtained using 1x10s copies of Ai or A2A RNA standard as the template for Real Time RT-PCR. Real-Time RT-PCR was performed as described in the methods section. Table 6: Optimization of MgCl2 concentrations for Real-Time RT-PCR MgCh Concentration (nM) Ai Assay: Cj1 A2A Assay: Cr1 3.5 40.0 28.6 4.5 40.0 27.6 5.5 40.0 28.2 6.5 40.0 27.6 7.5 30.3 27.6 1 The threshold cycle (Cj) was obtained using 1x10s copies of Aj or A2A RNA standard as the template for Real Time RT-PCR. Real-Time RT-PCR was performed as described in the methods section. 1. Quantification of Ai and A2A receptor, somatostatin and gastrin gene expression 1.1 Animal treatment Fasting: Male Wistar rats (250-300g) housed in light and temperature-controled rooms with free access to water were deprived of food for 24 or 36 h. Control animals were allowed free access to both food and water. Rats were anaesthetized with Somnotol® (60 mg/kg i.p.), and tissues were extracted between 11 a.m. to 12 p.m. for all groups. Omeprazole treatment: Male Wistar rats (250-270g) housed in light and temperature-controlled rooms with free access to food and water were used for this experiment. Omeprazole (AstraZeneca, Molndal, Sweden) was prepared for administration according to the manufacturer's instructions. Omeprazole (50 pmol/ml) was thoroughly dispersed in vehicle using the Tissumizer® tissue homogenizer. The vehicle consisted of 0.25% Methocel® (Dow, Midland, Michigan) containing 2mg/ml NaHC03 (pH 9.0). The drug was aliquoted, frozen, and stored at -20°C. The drug was thawed overnight at 4°C and brought to room temperature prior to administration. A new aliquot of drug was used daily to prevent degradation due to repeated freezing and thawing. Test groups were treated by gavage with omeprazole at a dose of 400 pmol/kg once daily between 9 and 10 a.m. for either 1 or 3 days. This omeprazole concentration and length of treatment were selected since gastric acid secretion is decreased by up to 80% after 1 day of treatment at 400 pmol/kg (Larsson et al., 1988; Lee et al., 1992), and steady state gastric acid inhibition is achieved after 3 days of treatment with the same dose (Carlsson et al., 1986). Control groups were treated similarly with vehicle. Animals were sacrificed 24 h after the last treatment of omeprazole. Rats were anaesthetized with Somnotol®(60 mg/kg, i.p.), and tissue extraction occurred between 10 a.m. to 11 a.m. 1.2 Tissue extraction The corporeal mucosa, corporeal muscle, and antrum, were dissected. These regions were selected because the corporeal mucosa contains somatostatin-secreting D-cells, the corporeal muscle contains somatostatin-releasing nerve fibers, and the antrum contains D-cells, gastrin-secreting G-cells, and somatostatin-releasing nerve fibers. Care was taken to avoid inclusion of tissue at the antro-corpus border. The corpus tissue was rinsed in ice-cold saline, and the mucosa was removed from the corpus with gentle scraping using a sterile glass slide. The mucosa was then rinsed off the slide with Trizol® reagent. Total RNA was extracted immediately from the corporeal mucosa as described in previous section. The corporeal muscle was flash frozen in liquid nitrogen and stored at -80°C until RNA extraction. Since the antrum is a relatively small tissue, total RNA was extracted from the whole antrum to ensure consistency between samples. The antral tissue was rinsed in sterile ice-cold saline, flash frozen in liquid nitrogen, and stored at -80°C until RNA extraction. 1.3 Sample preparation Total RNA was extracted using Trizol Reagent®, and concentrations were determined using the RiboGreen Quantitation Kit, as described earlier. Total RNA samples were then treated with DNase I and reverse transcribed as described in section II (3.2). 1.4 Real-Time RT-PCR The Real-Time RT-PCR procedure established in the present study was employed to measure adenosine Ai and A2A receptor mRNA levels. For these experiments, glyceraldehyde-3'-phosphate dehydrogenase (GAPDH), 18S ribosomal RNA (rRNA), somatostatin and gastrin gene expression levels were also measured. GAPDH and 18S rRNA gene expression: GAPDH mRNA and 18S rRNA levels were used as endogenous controls, and were measured using the Rodent GAPDH Control Reagents Kit (Applied Biosystems) and the TaqMan ribosomal RNA control reagents kit (Applied Biosystems), respectively. GAPDH mRNA and 18S rRNA levels were determined using the protocol described in the PE Applied Biosystems User Bulletin #2. A relative standard curve was constructed'using the Rodent Control RNA Standard (50 ng/pl) provided with the Rodent GAPDH Control Reagents Kit. Rodent Control RNA (100 ng) was reverse transcribed into 10 pi of cDNA using Superscript II RNase H- Reverse Transcriptase. This sample was designated as the 10 ng/pl sample since 10 ng of Rodent control total RNA was used to produce 1 pi of cDNA. Reverse transcription was performed as described in section II (3.2). The 10 ng/pl sample was diluted to 7.5, 5, 2.5, 1, 0.5, and 0.1 ng/pl, and these samples were used to construct the relative standard curve for the Real-Time RT-PCR assay. The 18S and GAPDH Real-Time RT-PCR assays were performed according to the manufacturer's instruction. Data were collected and analyzed as described in section II (3.2.4). Since fasting did not affect 18S rRNA levels, this value was subsequently used to normalize A\ receptor, A2A receptor, somatostatin and gastrin gene expression for possible differences arising from variations in total RNA extraction efficiency, first strand cDNA synthesis efficiency, and total RNA and cDNA degradation. Gastrin and somatostatin gene expression: Relative gastrin and somatostatin gene expression levels were measured using a sample of rat antrum total RNA to produce the standard curve. Antral total RNA (1 pg) was reverse transcribed into 10 pi of cDNA using Superscript II RNase H- Reverse Transcriptase. This sample was designated as the 100 ng/pl sample since 100 ng total RNA was reverse transcribed to produce 1 pi of cDNA. This sample was diluted to various concentrations to produce a relative standard curve. Standard sample concentrations ranged from 0.1 ng/pl to 100 ng/pl The primers and probes used to measure somatostatin and gastrin gene expression were designed using Primer Express and are listed in Table 7. The same guidelines which were used to design the Ai and A2A receptor primers and probes were used here (see Section 3.2.1). The primers and probes were synthesized by the NAPS Unit at UBC, and by Synthegen, respectively. Gastrin and somatostatin mRNA levels were measured by performing a two-step Real-Time RT-PCR assay as described in section II (3.2.3). The primer, probe and MgCl2 concentrations used for these assays were optimized according to the method described in section II (3.2.5). For both assays, the primer and probe concentrations were found to be 100 nM, and the MgCl2 concentration was found to be 7.5 mM. 1.5 Data collection and analysis Data were collected as described in section 3.2.4. To compare gene expression levels between the test and control animals, measurements were first normalized using the endogenous housekeeping-gene (18S rRNA) expression levels. This was performed according to the procedures described in the Applied Biosystems User bulletin #2: The Table 7: Gastrin and somatostatin Real-Time RT-PCR assay: primers and probes Rat Gastrin1 Position Length Tm %GC Sequence Forward 6 22 61 50 5 'TGCCACAAC AGTTAACCGTTCC3' Probe 80 28 69 57 5 TGCTGGCTCTAGCTACCTTCTCGGAAGC3' Reverse 129 20 61 60 5' CTGGGAGCGAGGTTTCCAAG3' Amplicon 6-129 124 82 54 Rat Somatosl tatin2 Position Length Tm %GC Sequence Forward 234 21 59 52 5' TG AGCCC AACC AG AC AGAGAA3' Probe 260 23 69 65 5 'CCCTGGAGCCTGAGGATTTGCCC3' Reverse 308 21 58 57 5' CTC ATCTCGTCCTGCTC AGCT3' Amplicon 234-308 75 84 60 Based on rat gastrin mRNA sequence; Accession number: M38653 2 Based on rat somatostatin mRNA sequence; Accession number: M25890 OS adenosine Ai or A2A receptor, gastrin or somatostatin gene expression levels for each individual sample were divided by the housekeeping-gene expression level to obtain a normalized value of measurement (MN). The mean ± S.E.M. of MN for the fed control and fasted test group were calculated. The mean of each control group was then expressed as 100%, while the gene expression levels of the test groups were expressed as a percentage of controls. Statistical significance was determined using the two-tailed unpaired Student's t-test, where P < 0.05 is considered significant. 2. Effect of omeprazole on A] and A2A receptor-induced changes in SLI and IRG release Male Wistar rats (250-275g) were treated with omeprazole as described in the previous section. Rats were deprived of food for 12 h but had free access to water prior to stomach perfusion. Stomach perfusion was performed 24 h after the last drug treatment. Animals were anaesthetized with Somnotol (60mg/kg), and stomachs were prepared for perfusion as decribed in Section I (1). The Ai receptor agonist CPA and A2A receptor agonist CGS 21680 were administered by side arm infusion at a rate to give a final concentration of 0.1 or 1 pM. These concentrations were selected because CPA has been shown to significantly inhibit and stimulate gastric SLI release when it is administered at a concentration of 0.1 and 1 pM, respectively. Both concentrations of CPA have also been shown to inhibit IRG release. The administration of CGS 21680 has been shown to significantly stimulate gastric SLI release at a concentration of 0.1 pM and to maximally stimulate SLI release at a concentration of 1 pM (see results). The effect of omeprazole treatment on CPA induced changes in SLI and IRG release and CGS-induced changes in SLI release was examined. Samples were collected and stored as described in section I (2). Radioimmunoassay was performed to measure SLI and IRG release as described on section I (3 & 4). The release of SLI and IRG were both expressed as the mean ± SEM for each 5 min sample. SLI and IRG release (%), and the % change in release was performed as described in Section I, 5. Statistical significance was determined using the paired Student's t-test, where P < 0.05 is considered significant. Single-factor ANOVA for repeated measures followed by Dunnett's t-test was also performed using GraphPad Prism. IV. Cellular localization of the gastric Ai and A2A receptors 1. Tissue preparation for immunohistochemistry (IHC) The methods used for IHC have previously been described (Yip et al., 2003). Four male Wistar rats (225-250 g) were anaesthetized with Somnotol (60 mg/kg i.p.). Stomachs were dissected out, cut along the greater curvature, and thoroughly rinsed with ice cold saline. Tissue specimens were immersion fixed in freshly prepared 0.1 M phosphate buffer (pH 7.4) containing 4% paraformaldehyde for 30 min (A2A receptor IHC) or overnight (Ai receptor IHC) at 4°C. This was followed by cryoprotection in 0.1 M phosphate buffer containing 20% sucrose for 12 h. The corpus (excluding the region adjacent to the fundus and antrum) and the antrum were mounted on corks, embedded in O.C.T. compound (Miles, Etobicoke, ON) and frozen in isopentane (-60°C) for 1 min. Cryostat sections of 30 pm thickness were cut at -25°C and placed in 0.1M phosphate buffered saline (PBS) containing 0.1% sodium azide. Random tissue sections were placed in 0.1 M PBS (pH 7.4) and stained as floating sections to enhance the penetration of antisera into these sections. Prior to the application of primary antibody, sections were sequentially incubated in 0.1 M PBS containing 50 mM NH4C1 for 30 min, 0.1 M PBS containing 100 mM glycine for 10 min, and 0.1 M PBS containing 1% BSA and 0.3% Triton X-100 for 1 h. Primary and secondary antibodies were diluted in 0.1 M PBS containing 1% BSA, 0.3% Triton X-100, and 0.1% sodium azide. Sections were incubated in adenosine Ai or A2A receptor primary antibody for 72 h at 4°C, washed in 0.1 M PBS (3x15 min), and incubated with cyanine Cy3-conjugated secondary antibody overnight at 4°C. Sections were again washed in 0.1 M PBS (3x15 min) and either double stained or mounted onto glass slides. For double staining, tissue sections were incubated in one of the other primary antibodies for 72 h at 4°C, washed in 0.1 M PBS (3x15 min) and incubated with Alexa Fluor® 488-conjugated secondary antibody overnight at 4°C. Sections were again washed in 0.1 M PBS (3x15 min) and mounted onto glass slides. Coverslips were placed on sections using a mixture of 0.1 M PBS in glycerin (1:9) and sealed with nail polish. 2. Adenosine receptor antibodies and control peptides The rabbit anti-Ai adenosine receptor antibody was purchased from Sigma-Aldrich and was developed using amino acids 309-326 of the rat Ai adenosine receptor C-domain as the immunogen. This antibody has been shown to cross-react strongly with the rat Ai receptor (Nakata, 1993; Ciruela et al., 1995; Rivkees et al., 1995). The specificity of this antibody in rat tissues has been examined by Western blotting (Carruthers et al., 2001). When samples were deglycosylated, this antibody detected only one band of the expected size (38-40k Da). In addition, pre-absorption of this antibody with the Ai receptor peptide resulted in a lack of immunoreactivity (Caruthers, 2001). The Ai receptor antibody was used at a dilution of 1:500 and immunostaining was detected using Donkey anti-rabbit IgG conjugated to Cy3 (Jackson ImmunoReasearch, West Grove, PA) at a concentration of 1:2000. Since an anti-rat A2A receptor antibody was unavailable, a rabbit anti-canine A2A receptor antibody was used. This antibody was purchased from Alpha Diagnostic International (San Antonio, TX) and was generated using an immunogen corresponding to a 30 amino acid sequence in the 4th intracellular domain of the canine A2A receptor. This region has 43% homology with the rat A2A receptor and has been shown to cross-react with the rat adenosine A2A receptors (Diniz et al, 2003). Its specificity has previously been established (Nie et al, 1999; Christofi et al, 2001). This antibody was used at a dilution of 1:200 and immunostaining was detected using donkey anti-rabbit IgG conjugated to Cy3 at a concentration of 1:2000. Since the blocking peptide for the Ai receptor antibody was not available from Sigma-Aldrich, one was custom made by the NAPS Unit at UBC. This synthetic peptide was based on the following immunogen sequence provided by Sigma-Aldrich: Cys-Gln-Pro-Lys-Pro-Pro-Ile-Asp-Glu-Asp-Leu-Pro-Glu-Glu-Lys-Ala-Glu-Asp. The blocking peptide was re-dissolved in 0.1 M PBS (pH 7.4), and the concentration was determined using the BCA assay (Pierce, Rockford, IL). The blocking peptide for the A2A receptor antibody was obtained from Alpha Diagnostic International. Preabsorption of the A2A receptor primary antibody with this control peptide has been shown to abolish A2A receptor labeling (Christofi et al, 2001). 3. Antibodies used for double-staining To examine whether the adenosine Ai and A2A receptors are localized on specific cell types, double staining was performed to identify somatostatin, gastrin, H+K+-ATPase p, von Willebrand's factor (VWF), and protein gene product (PGP) 9.5. The secondary antibody, donkey anti-mouse IgG conjugated to Alexa Fluor® 488 (1:2000, Molecular Probes, Eugene, OR) was used to detect this staining. Somatostatin-IR was used to identify D-cells of the corporeal and antral mucosa and somatostatin-IR nerves within the muscle layers and enteric plexi. Somatostatin was stained using the somatostatin antibody (SOMA 8; 1:500) provided by the MRC regulatory peptide group (UBC). G-cells of the antral mucosa were identified by gastrin staining using the gastrin antibody (109-21; 1: 30,000) provided by the late Dr. John Walsh. Both these antibodies have been used to identify G-cells (Pederson et al., 1984) and D-cells (Buchan et al., 1985b) in the rat stomach. The parietal cells were identified by staining the P-subunit of the proton pump (H+K+-ATPase). The H+K+-ATPase P antibody was purchased from Affinity Bioreagents Inc. (Golden, CO) and used at a concentration of 1:2000, as suggested by the manufacturer. To identify vascular endothelial cells lining blood vessels in the stomach, VWF was stained. This protein is synthesized by endothelial cells and is stored in the Weibel-Palade bodies of the endothelial cells (Schmugge et al., 2003). The VWF antibody was used at a dilution of 1:100 and was purchased from Serotec (Oxford, England). PGP 9.5 staining was used as a neuronal marker. This protein is expressed in the cytoplasm of central and peripheral neurons (Schofield et al., 1995), and has previously been used to study the innervation of the gastrointestinal tract (Yip et al., 2003). The PGP 9.5 antibody was used at a dilution of 1:200 and was purchased from Abeam Limited (Cambridge, UK). The specificity of these antibodies have previously been characterized (Wall et al, 1980; Buchan et al., 1985b; Vincent et al., 1985; Wilson et al., 1988; Chow and Forte, 1993). 4. Confocal microscopy Ai receptor immunoreactivity (AiR-IR) and A2A receptor immunoreactivity (A2AR-IR) were viewed using the Biorad Radiance 2000 confocal scanning laser system mounted on a Nikon Eclipse TE300 inverted microscope. The system utilizes a krypton gas laser with an excitation wavelength of 568 nm and emission filter 575-625 nm (for visualization of Cy3), and an excitation wavelength of 488 and emission filter 500-530 nm (for visualization of Alexa 488). Lens magnification of x40 was used with a zoom factor of 1.0 and z-step of 0.5 to 1.0 pm, while lens magnification of x60 was used with a zoom factor of 1.0-1.6 and a z-step of 0.3 to 0.5 pm. The software program Lasersharp 2000 (version 4.1, Biorad) was used to scan tissues sequentially using the red and green collection channels and the Kalman collection filter (n=2). The Kalman collection filter averages the images collected to increase the signal to noise ratio. Images of 512x512 pixels were obtained and analyzed using NIH image (National Institutes of Health, Bethesda, MD, USA) and Adobe Photoshop (version 7.0, Adobe Systems Inc.). To determine whether co-localization occurs, a single optical image collected from the red channel (AiR-IR or A2AR-IR) was overlaid on the corresponding single optical image collected from the green channel (somatostatin, gastrin, VWF, PGP, H+K+-ATPase, VIP or CGRP-IR) using Adobe Photoshop. 5. Quantification of co-localization Co-localization of AiR-IR and A2AR-IR with somatostatin-IR and gastrin-IR was quantified in the antral and corporeal mucosa by examining at least 3 tissue sections from 4 different animals. For quantification of AjR-IR with gastrin-IR, at least 12 fields of view were analyzed. AiR-IR was significantly more abundant than A2AR-IR. The corporeal and antral mucosa contained only 1.1±0.2 and 1.3±0.2 A2AR-IR cells per view, respectively. Thus, a total of at least 55 fields of view were analyzed for quantification of A2AR-IR with somatostatin-IR. 6. Control studies Immunoneutralization experiments were performed to confirm antibody specificity. The A]R antibody was neutralized by incubating lpl of the stock antibody (Sigma-Aldrich) with 0.1 pg of the blocking peptide in 0.5 ml 0.1 M PBS containing 1% BSA, 0.3% Triton X-100, and 0.1% sodium azide for 2 nights at 4°C. As suggested by the manufacturer, the A2AR antibody was neutralized by incubating 1 pg of antibody with 40 pg of control peptide in 0.2 ml of 0.1 M PBS containing 1% BSA, 0.3% Triton X-100, and 0.1% sodium azide for 2 nights at 4°C. A positive control lacking the control peptide was also prepared for both antibodies. Tissues stained with the neutralized antibody did not demonstrate AiR-IR or A2AR-IR, while tissues stained with the positive control demonstrated extensive AiR-IR and A2AR-IR. Additional control experiments were also performed to ensure that non-specific binding did not occur. These included sections which were processed as follows: 1) incubation in 1% BSA in place of the primary antibody, 2) incubation in adenosine Ai or A2A receptor primary antibody without secondary antibody, and 3) incubation in adenosine Ai or A2A primary antibody followed by Alexa 488-conjugated anti-mouse IgG secondary antibody. No immunostaining was observed following these procedures. To ensure that bleed-through did not occur, single-stained sections were examined under the microscope using both filters. Bleed-through was not detected for any of the antibodies used. Chapter 3: Results I. Effect of adenosine on gastric IRG and SLI release 1. Effect of adenosine analogs on gastric IRG release The effect of CPA (Ai-selective agonist), CGS 21680 (A2A-selective agonist), and IB-MECA (A3-selective agonist) on the inhibition of IRG release from the isolated vascularly perfused stomach was examined. The basal release of IRG during periods 1-3 remained relatively constant for all experiments. The basal IRG release (periods 1 to 3) ranged from 114±19 to 117±19pg/min (Fig. 11). When 1 pM CPA was administered, gastric IRG release was inhibited immediately upon drug perfusion (Fig. 11). The inhibition was apparent throughout the drug perfusion period, and the release of IRG returned to basal levels within 5 min after drug removal. The effect of various concentrations of CPA on IRG release was also examined. Results are expressed in % inhibition and are summarized in Fig. 12. CPA caused a concentration-dependent inhibition of IRG release. Maximal inhibition was achieved at a concentration of 1 pM. The administration of 10 pM CPA did not further suppress IRG release. The empirical EC50 of CPA in inhibiting IRG release was estimated to be 0.067 pM, with a confidence interval between 0.014 and 0.325 pM. The administration of CGS 21680 (0.1 pM) and IB-MECA (0.1 pM) did not alter basal IRG release. 200 -i o 150H (0 | 100-5 50H CPA (1uM) I X 1 2 3456 7 89 10 5 min periods Fig. 11. Effect of CPA on gastric IRG release. Results are expressed as IRG release (%) as described in the Methods section. Each column represents the mean ± SEM of 5 experiments. *P < 0.05 and ***p < 0.001 when compared with period 3 using repeated measures ANOVA followed by Dunnett's multiple comparison test. 100n 90-80-70-60-50-40-30-20-10-0--10-*** *** 0.001 0.01 0.1 1 10 0.1 CPA CGS 21680 (|iM) (|iM) 0.1 IB-MECA (ixM) Fig. 12. Effect of various concentrations of CPA, and 0.1 uM of CGS 21680 and IB-MECA on IRG release. Results are expressed as the % inhibition of IRG release, and calculated as described in the Methods section. Each column represents the mean ± SEM of at least 5 experiments; *P < 0.05 and ***P < 0.001 when the mean IRG release (pg/min) during drug perfusion periods 4-7) is compared to the mean basal IRG release (periods 1-3). 2. Effect of adenosine receptor antagonists on gastric IRG release Adenosine has been shown to inhibit gastric IRG release (Kwok et al, 1990). To determine the receptor subtype involved in this adenosine-mediated action, the effect of the Ai receptor-selective antagonist, DPCPX, and the A2 receptor-selective antagonists, DMPX, on adenosine-induced inhibition of IRG release were examined. Neither antagonists altered basal IRG release (Fig. 13). Fig. 14 shows the effect of DPCPX and DMPX on adenosine-induced changes in IRG release. After a 15 min basal period, 1 pM of the antagonist was administered alone for 5 min prior to the concomitant perfusion with adenosine (10 pM) for 15 min. In the presence of 1 pM DPCPX, the adenosine-induced inhibition of IRG release was completely abolished. DMPX (1 pM), however, had no effect on adenosine-induced changes in IRG release. 3. Effect of adenosine receptor agonists on gastric SLI release To determine which adenosine receptor subtype is involved in regulating gastric SLI release, the effect of CPA (Apselective agonist), CGS 21680 (A2A-selective agonist), and IB-MECA (A3-selective agonist) were examined. The basal release of SLI during periods 1-3 remained relatively constant for all experiments. In experiments examining the effect of agonists (Fig. 15 and 16), the rates of basal SLI release (period 1-3) were 174±24 tol78±27 pg/min (CPA 0.1 pM), 173±19 to 180±21 pg/min (CPA 1 pM), 92±18 to 95±18 pg/min (CGS 21680) and 191±39 to 195±41 pg/min (IB-MECA). When administered at a concentration of 0.1 pM, CPA immediately inhibited gastric SLI release (Fig. 15A). The inhibition continued throughout the drug perfusion period, and SLI release returned to basal (A 10 | 100-o CC 50-DPCPX(VM) 23456789 10 5 min periods B 200-i 9 15(H CO | 100-0£ O 0£ 50-DMRX(1 nM) 23456789 10 5 min periods Fig. 13. Effect of DPCPX and DMPX on gastric IRG release. Each column represents the mean ± SEM of at least 5 experiments. Statistics were performed using repeated measures ANOVA followed by Dunnett's multiple comparison test. Neither drug altered basal IRG release. (A <0 cc o ft ADO(10nM) 50-*** *** 1 23456789 10 5 min periods B 200: v 15°: tn to | 100-1 K O K DPCFX(1 nM) ADO(10iiM) > 50-0: 1 23456789 10 5 min periods 4) in a a> <u oc O 0£ 200 -150-^ 100-^ 0: DMPX(1 nM) ADO(10(iM) _ 1 1 23456789 10 5 min periods Fig. 14. Effect of DPCPX and DMPX on adenosine-induced changes in IRG release. A) Effect of adenosine on IRG release; B) Effect of DPCPX on adenosine-induced changes in IRG release; C) Effect of DMPX on adenosine-induced changes in IRG release. Results are expressed as IRG release (%) as described in the Methods section. Each column represents the mean ± SEM of at least 5 experiments. *P < 0.05 and ***P < 0.001 when compared with period 3 using repeated measures ANOVA followed by Dunnett's multiple comparison test. ~ 150-(0 0) (0 50-CPA (O.VM) 1 23456789 10 5 min periods B 500-^ 400-58 300-« t£ 200-• W 100-1 i—i 0 CPA (1 [Ml) 1 23456789 10 5 min periods Fig. 15. Effect of 0.1 uM (A) and 1 uM (B) CPA on gastric SLI release. Results are expressed as SLI release (%) as described in the Methods section. Each column represents the mean ± SEM of at least 5 experiments. *P < 0.05 when compared with period 3 using repeated measures ANOVA followed by Dunnett's multiple comparison test. 2000-1 — 1500-0) W | 1000H OS CO 500-CGS 21680 (0.1 (iM) II. 3 4 5 6 7 8 5 min periods 9 10 B 0) _0) 0) 300-1 200-_ 100-_l CO IB-MECA (0.1 pM) 3 4 5 6 7 8 9 5 min periods 10 Fig. 16. Effect of 0.1 uM CGS 21680 (A) and IB-MECA (B) on gastric SLI release. Results are expressed as SLI release (%) as described in the Methods section. Each column represents the mean ± SEM of at least 5 experiments. *P < 0.05 when compared with period 3 using repeated measures ANOVA followed by Dunnett's multiple comparison test. Note the difference in the scale of the ordinate. levels 10 min after the drug was removed. Conversely, when CPA was administered at a concentration of 1 pM, an increase in SLI release was observed 15 min after the start of drug perfusion. SLI release returned to basal levels within 5 min after the cessation of CPA perfusion (Fig. 15B). The administration of 0.1 pM CGS 21680 caused a significant increase in SLI release immediately upon its administration (Fig. 16A). The release of SLI continued to gradually increase throughout the drug perfusion period, and reached a plateau at period 6 (1100=1=135 % of basal SLI release). SLI release returned to basal levels 10 min after the drug was removed. IB-MECA (0.1 pM) did not significantly affect SLI release during the first 15 min of drug administration (Fig. 16B), but did significantly increase gastric SLI release to 144 ±8% after 15 min of perfusion. SLI release returned to basal levels 10 min after the withdrawal of drug administration. To examine the concentration dependent effect of CPA, CGS, and IB-MECA in stimulating SLI release, stomachs were perfused with 1 nM to 10 pM of these drugs. Results are expressed as percentage changes of SLI release and shown in Fig. 17. CGS 21680 was the most potent adenosine agonist in stimulating SLI release. CGS 21680 significantly increased SLI release starting at the lowest concentration examined (10 nM; 238±38%). Maximal augmentation was observed at 1 pM (1237±120%). The effect elicited by 1 pM and 10 pM of CGS 21680 were not significantly different. Both IB-MECA and CPA were less potent than CGS 21680. Although higher concentrations (1 and 10 pM) of CPA and IB-MECA enhanced SLI release, lower concentrations of CPA (10 nM and 0.1 pM) significantly inhibited SLI release (P < 0.05). The % changes of SLI release in the presence of 10 nM and 0.1 pM CPA were -12±3 and -19±5%, respectively. Although IB-MECA may alter SLI release (%) in individual samples (4-7) during drug perfusion (Fig. 16B), it did not Fig. 17. Effect of various concentrations of adenosine agonists on gastric SLI release. A: The effect of CGS 21680 on SLI release. B: The effect of CPA (•) and IB-MECA (•) on SLI release. Results are expressed as percentage (%) changes and calculated as described in the Methods section. Each point represents the mean ± SEM of at least 5 experiments. Note the difference in the scale of the ordinate. significantly alter SLI release (% change) when perfused at concentrations from 1 nM to 0.1 pM. The EC50 of CGS 21680 in stimulating SLI release was estimated to be 0.06 pM, with a 95% confidence interval between 0.02 and 0.17 pM. The empirical EC50 of CPA and IB-MECA were estimated to be 3.5 pM (95% confidence interval: 0.5 pM to 23 pM) and 0.3 pM (95% confidence interval: 0.02 pM and 4 pM), respectively. The rank order of potency of these analogs in augmenting SLI release, therefore, was CGS 21680 > IB-MECA > CPA. 4. Effect of adenosine receptor antagonists on gastric SLI release The effect of DMPX, an A2 receptor-selective antagonist was studied to examine the role of A2 receptors in adenosine and adenosine agonist-induced gastric SLI release. When perfused through the gastric vasculature for 20 min, 1 pM DMPX did not alter basal SLI release, while 10 pM DMPX significantly suppressed SLI release. Experiments similar to those described for Fig. 14 were then performed to test the effect of DMPX on adenosine and adenosine receptor agonist-induced release of SLI; 1 or 10 pM DMPX was administered 5 min prior to the concomitant perfusion with the agonist for 15 min. Results are summarized in Fig. 18. In the presence of 1 pM DMPX, adenosine induced-SLI release was attenuated. DMPX (1 pM) also inhibited the response elicited by CGS 21680 (0.01 and 0.1 pM) and IB-MECA (lpM), and completely abolished the effects of CPA (1 pM) on gastric SLI release. CGS 21680-induced release of SLI was significantly suppressed by DMPX in a concentration-dependent manner. The % change of SLI release induced by 0.01 pM CGS 21680 in the presence of 10 pM DMPX was not significantly different from the basal SLI release. 800- I 600-* 400- X 200- X X o--200-DMPX(nM) 1 10 0 1 0 1 10 0 1 10 0 1 0 1 + ADO + CGS + CGS + CPA + IB-MECA (1 uty) (0.01 uM) (0.1 nM) (1 H-M) (1(xM) Fig. 18. Effect of DMPX on basal, adenosine, and adenosine agonist-stimulated SLI release. The effect of DMPX on CPA-, CGS 21680- (CGS), and IB-MECA-induced SLI release was examined. Results are expressed as % changes. Each column represents the mean ± SEM of at least 5 experiments; tp < 0.05 when the mean SLI release (pg/min) in the presence of DMPX (periods 4-7) is compared to the mean basal SLI release (periods 1-3), *P < 0.05 and **P < 0.01 when the mean release of SLI (periods 4-7) in the presence of agonists was compared with the mean release of SLI in the presence of both DMPX and agonists. To test if the stimulatory effect on SLI release is mediated by the A2A receptor, the effect of the potent selective A2A receptor antagonist, ZM 241385, was examined. When perfused alone, ZM 241385 (1 pM) significantly inhibited basal SLI release during period 5 and 7 (Fig. 19A). However, when the % change of SLI release was calculated, SLI release in the presence of 1 pM ZM 241385 did not differ significantly from basal SLI release (Fig. 20). ZM 241385 abolished the stimulatory effect of adenosine; in the presence of ZM 241385 and adenosine, the release of SLI was inhibited (Fig. 20). When perfused concomitantly with CGS 21680 (0.1 pM), ZM 241385 (1 pM) also abolished the stimulatory effect of CGS 21680, and a significant inhibition of SLI release was apparent (Fig. 19). The effect of 10 pM ZM 241385 on CGS 21680 (0.1 pM) were similar to that of 1 pM ZM 241385. The stimulatory effect of CPA (lpM) and IB-MECA (1 pM) on SLI release was also completely abolished by ZM 241385. In the presence of both the agonist and antagonist, an inhibition of SLI was apparent (Fig. 20A). The effect of the Ai selective antagonist DPCPX was also examined. DPCPX (1 pM) significantly inhibited basal SLI release, but had no effect on CGS 21680- (0.1 pM) stimulated SLI release (Fig. 20). 5. Involvement of endogenous adenosine on SLI release To examine the role of endogenous adenosine in modulating SLI release, experiments were also performed to test the effect of EHNA, an ADA inhibitor, on gastric SLI release using the same experimental protocol described in Fig. 13. EHNA (1-10 pM) caused a concentration-dependent increase in SLI release, while 0.1 pM EHNA had no effect (Fig. 21). The stimulatory effect of adenosine on SLI release was potentiated in the presence of EHNA. gj 200-(0 aj CD DC — 100-_l (/) ZM 241385 (1|xM) X 1 23456789 10 5 min periods B 2000-1 1500-Q) W CO CD 0) 0£ 1000 500-OGS (0.1 nM) I ELTZL 1 23456789 10 5 min periods 300-1 0) (A CO aj — 100-(0 ZM 241385 (1 (iM) 200- CGS (0.1 nM) ***** 1 23456789 10 5 min periods Fig. 19. Effect of ZM 241385 on basal and stimulated SLI release. A: The effect of ZM 241385 on basal SLI release; B: The effect of CGS 21680 (CGS) on basal SLI release; C: The effect of ZM 241385 on CGS-stimulated SLI release. Each column represents the mean ± SEM of at least 4 experiments; *P < 0.05 when compared with period 3 of respective experiments using repeated measures ANOVA followed by Dunnett's multiple comparison test. 800 |) 600 1 u gT 400 CO 200 -200 t 1 10 ZM 241385 (M.M) X I ZM 241385 (nM) + ADO (1 nM) 0 1 10 ZM 241385 (nM) + CGS (0.1 nM) "a 0 1 0 1 ZM241385 (nM) ZM241385((iM) + CPA (1 (iM) + IB-MECA (1 ^M) B 1000-800-v at c 10 600-400-200-0--200J I DPCPX (UM) 0 1 DPCPX (nM) f CGS (0.1 M-M) Fig. 20. Effect of ZM 241385 (A) and DPCPX (B) on basal and adenosine (ADO) and agonist-stimulated SLI release. The effect of ZM 241385 on CPA-, CGS 21680- (CGS), and IB-MECA-induced SLI release was examined. Results are expressed as % changes. Each column represents the mean ± SEM of at least 4 experiments; ^P < 0.05 and ^P < 0.01 when the mean SLI release (pg/min) in the presence of ZM 241385 or DPCPX (periods 4-7) was compared to the mean basal SLI release (periods 1-3), **P < 0.01 when the mean release of SLI (periods 4-7) in the presence of agonists was compared with the mean release of SLI in the presence of both ZM 241385 and agonists. 600-1 0> O) 500-(0 c O 400-T <B 300-<n CO CO « 200-CO 100-tt 0.1 1 10 EHNA (nM): 0 1 EHNA (|xM) + ADO (1|xM) Fig. 21. Effect of EHNA on basal and adenosine (ADO)-induced SLI release. Results are expressed as % changes of SLI release, and each column represents the mean ± SEM of at least 6 experiments; fP < 0.05 and t|P < 001 when comparing the mean SLI release (pg/min) in the presence of EHNA during periods 4-7 with that of periods 1-3 (basal release), **P < 0.01 when compared with the ADO-induced release of SLI. II. Distribution and sequence analysis of adenosine receptor mRNA 1. Distribution of adenosine receptor mRNA in various regions of the stomach The regional distribution of all four gastric adenosine receptors was examined using RT-PCR. Results obtained from these experiments demonstrate that the mRNAs of all four adenosine receptors are present in every gastric region examined, including the fundus, corpus, antrum, and mucosa (Fig. 22). The mRNA of these receptors was also detected in the rat striatum, which was used as a positive control. Each PCR amplicon was of the expected length (Ai: 1086 bp; A2A: 441 bp; A2B 380 bp; A3; 489 bp). Since adenosine Ai and A2A receptors were shown to play a role in the control of gastric regulatory peptide release, these receptors were examined in more detail. RT-PCR was therefore performed using primers which spanned the entire coding region of the rat Ai and A2A receptor gene. Results demonstrate that alternative splicing does not occur in any region of the stomach since only one specific Ai and A2A receptor transcript was detected in all gastric regions studied (Fig. 23). The specificity of the RT-PCR reactions was confirmed by restriction enzyme digestion of the RT-PCR amplicons (Fig. 24-28). The observed fragment sizes shown on these figures are estimates based on the migration distance of the fragment. Small fragments of less than 150 bp were often not visible by gel electrophoresis due to the low intensities of their fluorescence under UV light. 1000 bp* 500 bp-^j 400 bp_^ 300 bp> 1000 bp-W 500 bp .J 400 bp-W 300 bp* Fig. 22. Results of A,, A2A, A2B and A3 RT-PCR performed using rat gastric tissues. RT-PCR was performed using tissues from the rat fundus (A), corpus (B), antrum (C) and mucosa (D) and primers designed to detect the A, (1086 bp), A2A (441 bp), A2B (380 bp), and A3 (489 bp) receptor mRNA (see Table 3). All four adenosine receptor subtypes are present in the gastric tissues examined. 1337 bp Fig. 23. Amplification of the A, (A) and A2A (B) receptor coding region by RT-PCR. RT-PCR was performed using tissues obtained from the striatum (positive control; lane 2), fundus (lane 3), corpus (lane 4), antrum (lane 5) and mucosa (lane 6). The A[FL and A2AFL primer sets (see Table 3) used span the entire coding region of the rat adenosine A, and A2A receptor and generated only one amplicon, of the expected length, for each tissue examined. 1000 bp 800 bp 300 bp • 200 bp • •303 bp 208 bp Uncleaved size & region amplified* Enzyme Recognition site Cleavage site Expected fragments (bp) Observed fragments (bp) 1086 bp (322-1407 bp) Narl GG/CGCC 527 880; 206 907; 208 Sac I GAGCT/C 631 776; 310 791; 303 *Positions based on A, receptor sequence in the rat brain (Accession No. M64299) Fig. 24. Restriction enzyme digestion of the adenosine A, receptor PCR amplicon. RT-PCR products synthesized using the A[FL primer set were concentrated, purified, and cleaved with restriction enzymes, Nar I and Sac I, and separated on a 2% agarose gel. Expected fragment sizes are listed in the table. The size of the observed fragments were estimated using the Stratagene Eagle Sight Image Analysis Software, as described in the Methods. EcoRI Mbo I Pstl Rsa I Uncleaved size & region amplified* Enzyme Recognition site Cleavage site Expected fragments (bp) Observed fragments (bp) 441 bp from 99-449 bp EcoRI G/AATTC 385 286; 155 300;165 Mbo I /GATC 160 380; 61 390 Pstl CTGCA/G 178 362; 79 361 Rsa I GT/AC 399 300; 141 319;160 *Positions based on A2A receptor sequence in the rat brain (Accession No. S47609) Fig. 25. Restriction enzyme digestion of the adenosine A2A receptor PCR amplicon. RT-PCR products synthesized using the A2ASL primer set were concentrated, purified, and cleaved with restriction enzymes, EcoR I, Mbo, I, Pst I, and Rsa I. Fragments were separated on a 2% agarose gel. Expected fragment sizes are listed in the table. The sizes of the observed fragments were estimated using the Stratagene Eagle Sight Image Analysis Software, as described in the Methods. Ava II Hpa II Uncleaved size & region amplified* Enzyme Recognition site Cleavage site Expected fragments (bp) Observed fragments (bp) 380 bp from 223-603 bp Ava II G/G(A or T)CC 479 256;124 278 Hpa II C/CGG 413 190; 190 190 *Positions based on A2B receptor sequence in the rat brain (Accession No. M91466) Fig. 26. Restriction enzyme digestion of the adenosine A2B receptor PCR amplicon. RT-PCR products synthesized using the A2BSL primer set were concentrated, purified, and cleaved with restriction enzymes, Ava II and Hpa II. Fragments were separated on a 2% agarose gel. Expected fragment sizes are listed in the table. The sizes of the observed fragments were estimated using the Stratagene Eagle Sight Image Analysis Software, as described in the Methods. Ava II Hinf I 360 bp Uncleaved size & region amplified* Enzyme Recognition site Cleavage site Expected fragments (bp) Observed fragments (bp) 489 bp from 319-788 bp Ava II G/G(A or T)CC 452; 546 133; 262; 94 288 Hinf I G/A(ACG or T)TC 653 334;155 360 *Positions based on A3 receptor sequence in the rat brain (Accession No. M94152) Fig. 27. Restriction enzyme digestion of the adenosine A3 receptor PCR amplicon. RT-PCR products synthesized using the A3SL primer set were concentrated, purified, and cleaved with restriction enzymes, Ava II and Hinf I. Fragments were separated on a 1% agarose gel. Expected fragment sizes are listed in the table. The sizes of the observed fragments were estimated using the Stratagene Eagle Sight Image Analysis Software, as described in the Methods. EcoR I Hinf I Uncleaved size & region amplified* Enzyme Recognition site Cleavage site Expected fragments (bp) Observed fragments (bp) 1337 bp from 41-1377 bp EcoR I G/AATTC 385 992; 345 1015; 356 Hinf I G/A(ACG or T)TC 719 658; 679 -690 *Positions based on A2A receptor sequence in the rat brain (Accession No. S47609) Fig. 28. Restriction enzyme digestion of the full length adenosine A2A receptor amplicon. RT-PCR products synthesized using the A2AFL primer set were concentrated, purified, and cleaved with restriction enzymes, EcoR I and Hinf I. Fragments were separated on a 2% agarose gel. Expected fragment sizes are listed in the table. The sizes of the observed fragments were estimated using the Stratagene Eagle Sight Image Analysis Software, as described in the Methods. 2. Ai and A2A receptor coding region and A2B and A3 receptor partial cDNA sequence Experiments were performed to determine the sequences of the Ai and A2A receptor coding regions, and the sequences of the partial A2B and A3 receptor cDNAs. The mucosal Ai and A2A receptor sequences cloned using the full length Ai and A2A receptor primers (Table 3) were submitted to Genbank and assigned accession numbers AF042079 and AF228684, respectively. The A2B and A3 receptor sequences cloned using the short length A2B and A3 receptor primers (Table 3) were assigned accession numbers AF084241 and AF102804, respectively. Results of the sequencing experiments demonstrate that the coding region of the gastric mucosal Ai receptor (accession # AF042079) was identical to the published sequence in the rat brain (Mahan et al, 1991). The sequence of the gastric mucosal A2A receptor coding region (accession # AF228684) was also found to be identical to that in the brain (Fink et al, 1992). Fig. 29 and 30 show the alignments of the rat mucosal and rat brain Ai and A2A receptor coding regions. DNA sequence analysis was also performed on the PCR amplicons generated using the A2BSL and A3SL primer sets listed in Table 3. An alignment of these regions with corresponding regions of the rat brain receptors is shown in Fig. 31. The sequence of the mucosal A2B receptor PCR amplicon is identical to that of the rat brain. However, analysis of the A3 receptor PCR amplicon reveals a silent mutation at position 492. At the third position of the codon for alanine, adenine (A) is replaced by thymine (T). The position of this mutation corresponds to amino acid #59 of the A3 receptor protein, which is situated in the second transmembrane domain (Salvatore et al, 1993). 1 tgtctgctgatgtgcccagctcctgcccaccatgccgccctacatctcggccttccaggc 60 541 gtttgagaaggttatcagcatggagtacatggtctacttcaacttcttcgtctgggtgct 600 1 1 1 1 1 1 1 1 1 1 1 1 1 1 11111111111 1 1 1 1 1 1 1 1 II 1 1 1 1 1 1 1 1 1 1 1 1 1 11111111111 1 1 1 1 1 1 1 1 1 1 1 1 II 111111111111111111111111111111111111 1 1 1 1 1 1 1 1 1 1 1 276 tgtctgctgatgtgcccagctcctgcccaccatgccgccctacatctcggccttccaggc 335 816 gtttgagaaggttatcagcatggagtacatggtctacttcaacttcttcgtctgggtgct 875 61 tgcctacattggcatcgaggtgctcattgccttggtctctgtgcccggaaatgtactggt 120 601 gccgccactgctcctcatggtcctcatctacctggaggtcttctacctgatccgtaagca 660 1II111111II111 1 1 111111111 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 11111111 1 1 1 1 1 1 1 1 1 1 II 1 1 1 11111111111111111111111111111111111 1 1 1 1 1 1 1 1 1 1 1 336 tgcctacattggcatcgaggtgctcattgccttggtctctgtgcccggaaatgtactggt 395 876 gccgccactgctcctcatggtcctcatctacctggaggtcttctacctgatccgtaagca 935 121 gatttgggctgtgaaggtgaaccaggcacttcgcgatgccaccttctgcttcatcgtgtc 180 661 gctcaacaaaaaggtgtcagcctcctccggtgacccccagaagtactacgggaaggagct 720 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1  1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 II 1 1 1 1 1 1 1 1 396 gatttgggctgtgaaggtgaaccaggcacttcgcgatgccaccttctgcttcatcgtgtc 455 936 gctcaacaaaaaggtgtcagcctcctccggtgacccccagaagtactacgggaaggagct 995 181 actggcggtagctgatgtggccgttggcgccctggtcatcccactggccatccttatcaa 240 721 gaagatcgccaagtcgctggccctcatcctcttcctctttgccctcagctggctgccgct 780 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 II 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1  1 1 1 1  1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 456 actggcggtagctgatgtggccgttggcgccctggtcatcccactggccatccttatcaa 515 996 gaagatcgccaagtcgctggccctcatcctcttcctctttgccctcagctggctgccgct 1055 241 cattgggccacagacctacttccacacctgcctcatggtggcctgccctgtcctcatcct 300 781 gcatatcttgaactgtatcaccctcttctgccccacctgccagaaacccagcattctgat 840 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1  1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 516 cattgggccacagacctacttccacacctgcctcatggtggcctgccctgtcctcatcct 575 1056 gcatatcttgaactgtatcaccctcttctgccccacctgccagaaacccagcattctgat 1115 301 cacccagagctccattctggctctgctcgccattgctgtggatcgatacctccgagtcaa 360 841 ctacatcgccatcttcctcacacacggcaactccgccatgaaccccatcgtctatgcctt 900 1 11 1 1 1 11 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 11 1 1 11 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 11 1 II1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1  1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 576 cacccagagctccattctggctctgctcgccattgctgtggatcgatacctccgagtcaa 635 1116 ctacatcgccatcttcctcacacacggcaactccgccatgaaccccatcgtctatgcctt 1175 361 gatccctctccggtacaagacagtggtgacccagcggcgggcggcagtggccatagctgg 420 901 ccggatccacaagttccgggtcacctttctgaagatttggaatgaccacttccgatgcca 960 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 11 11 1 111 1 111 1 1 II 1 1 1 11 1 1 11 1 1 1 1 1 1 1 1 1 1 1 636 gatccctctccggtacaagacagtggtgacccagcggcgggcggcagtggccatagctgg 695 1176 ccggatccacaagttccgggtcacctttctgaagatttggaatgaccacttccgatgcca 1235 421 ctgctggattctctcccttgtggtaggcctgacacccatgtttggctggaacaacctgag 480 961 gcctaagcctcccatcgatgaagacctcccagaggagaaagctgaggactagactctgcc 1020 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 696 ctgctggattctctcccttgtggtaggcctgacacccatgtttggctggaacaacctgag 755 1236 gcctaagcctcccatcgatgaagacctcccagaggagaaagctgaggactagactctgcc 1295 481 tgtggtagagcaagactggagagccaacggcagtgttggggagcccgtgatcaagtgtga 540 1021 ttgctccgtctagcccatgcccagcggctctctgttcaactcccacgttctccctgtccc 1080 1 1 1 1 1 1 1 1 1 1 1 1 1 111111 111111 1 1 1 1 1 1 1 1 1 1 1 1 1 1111 11111 II 1 1 1 111 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 756 tgtggtagagcaagactggagagccaacggcagtgttggggagcccgtgatcaagtgtga 815 1296 ttgctccgtctagcccatgcccagcggctctctgttcaactcccacgttctccctgtccc 1355 1081 accctgtc 1088 1 1 1 1 1 1 1 1 1356 accctgtc 1363 Fig. 29. Alignment of the rat mucosal and brain A! receptor coding regions. The rat mucosal A, receptor cDNA sequence (top strand; accession number: AF042079) is aligned with the corresponding region the rat brain receptor (bottom strand; accession number: M64299) o 1 ctgctgagcctgcccaagtgtggctgctcccaccatgggctcctcggtgtacatcacggt 60 661 gcccctgccaggggagcggactcggtccacgctgcagaaggaggtccacgctgccaagtc 720 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 M 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 I 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1  1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 41 ctgctgagcctgcccaagtgtggctgctcccaccatgggctcctcggtgtacatcacggt 100 701 gcccctgccaggggagcggactcggtccacgctgcagaaggaggtccacgctgccaagtc 760 61 ggagctggccatcgctgtgctggccatcctgggcaacgtgctcgtgtgctgggccgtgtg 120 721 cctggccatcatcgtcgggctctttgctctgtgctggttgccgctgcacatcatcaactg 780 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 11111111111111111111II111111111II1111 1 1 1 1 1 1 1 1 1 1 101 ggagctggccatcgctgtgctggccatcctgggcaacgtgctcgtgtgctgggccgtgtg 160 761 cctggccatcatcgtcgggctctttgctctgtgctggttgccgctgcacatcatcaactg 820 121 gatcaacagtaacctgcagaacgtcaccaacttctttgtggtatcgctggcggcggctga 180 781 tttcaccttcttctgctccacgtgccggcacgcccctccgtggctcatgtacctggccat 840 11111111111111II1111111111111111111111II 1 1 M 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 161 gatcaacagtaacctgcagaacgtcaccaacttctttgtggtatcgctggcggcggctga 220 821 tttcaccttcttctgctccacgtgccggcacgcccctccgtggctcatgtacctggccat 880 181 cattgcagtgggtgtgctcgccatccccttcgctatcaccatcagcaccggcttctgcgc 240 841 catcctctcccacagcaactccgtcgtcaaccccttcatctacgcctacaggatccggga 900 11111111 1 1 1 1 1 1 II 1 1 1 1 1 1 1 1 1 1111111111 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1111 1 1 1 1 1 1 1  1 1 1 1 1 1 1 1 1 1 1 1 111111111111111111111111111111111II11111 1 1 1 1 1 221 cattgcagtgggtgtgctcgccatccccttcgctatcaccatcagcaccggcttctgcgc 280 881 catcctctcccacagcaactccgtcgtcaaccccttcatctacgcctacaggatccggga 940 241 cgcctgccacggctgcctcttcttcgcctgttttgtcctggtcctcacgcagagttccat 300 901 gttccgccagaccttccggaagatcatccgaacccacgtcctgaggcggcaggaaccctt 960 1 1 11 111111 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 II11111111 1 1 1 1 1 1111111 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1111 1 1 1 1 1 1 1111 1 1 1 1 1 1 1 111 1 1 1 1 1 1 1 1111 1 II 1 1 1 1 1 111 1 1 1 1 1 1 1111 1 1 281 cgcctgccacggctgcctcttcttcgcctgttttgtcctggtcctcacgcagagttccat 340 941 gttccgccagaccttccggaagatcatccgaacccacgtcctgaggcggcaggaaccctt 1000 301 ctttagcctcttggctatcgccatcgaccgctacatcgccatccgaattccactccggta 360 961 ccaggcagggggttccagtgcctgggccctggcagctcacagcactgagggagagcaggt 1020 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 II 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 11111 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 11111111111111111 1 111111111 1 1 1 1 1 1 1 1 1 II 1 1 341 ctttagcctcttggctatcgccatcgaccgctacatcgccatccgaattccactccggta 400 1001 ccaggcagggggttccagtgcctgggccctggcagctcacagcactgagggagagcaggt 1060 361 caatggcttggtgacaggtgtgagggcgaagggcatcattgcaatttgctgggtgctgtc 420 1021 tagcccccgccttaatggccaccccctgggggtatgggccaacggcagtgccacccattc 1080 11 1 1 1 1 1 1 1 1 1 II 1 II 1 1 1 1 1 1 1 1 1 1 II 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 II 1 1 1 1 1 1 11 11 1 1 11111 1 1 1 1 1 1 1 1 1 1 1 1 11 1 1 1111111111111111 1 1111111111 1 1 1 1 II 1 1 1 1 1 1 401 caatggcttggtgacaggtgtgagggcgaagggcatcattgcaatttgctgggtgctgtc 460 1061 tagcctccgccttaatggccaccccctgggggtatgggccaacggcagtgccacccattc 1120 421 gtttgccattggcctgacccccatgctgggctggaacaactgcagtcagaaagacgggaa 480 1081 cggacggcggcccaatggctacactctggggctggggggtggagggagtgcccaaggctc 1140 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1  II 1 II 1 1 1 II11II1111111II11II111111 1 111111II1II1 1 1 II 1 1 1 II 1 461 gtttgccattggcctgacccccatgctgggctggaacaactgcagtcagaaagacgggaa 520 1121 cggacggcggcccaatggctacactctggggctggggggtggagggagtgcccaaggctc 1180 481 ctccacgaagacctgcggcgagggccgggtgacctgtctgttcgaggacgtggtgcccat 540 1141 tcctcgggatgtggagcttcctacccaggagcgccaggaaggccaagagcaccctggcct 1200 1 II 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1111111111 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 II 1 1 1 11111111111 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 111II1111111111 1 1 1111111II1 1 1 1 1 1 1 1 1 1 1 1 521 ctccacgaagacctgcggcgagggccgggtgacctgtctgttcgaggacgtggtgcccat 580 1181 tcctcgggatgtggagcttcctacccaggagcgccaggaaggccaagagcaccctggcct 1240 541 gaattacatggtttactacaacttctttgcgttcgtgttactgccccttctgctcatgct 600 1201 aaggggtcatctggtccaggctagagtaggagcttcctcatggtcttcagagtttgcccc 1260 1 1 1 1 1 1 1111111111111 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1II1111II1 1 1 1 1 II 1 1 1 1 1 1 1 1 11111 1 II 1 11 111 1 111 1 11 1 II1111111 1 1 111 1 1 1 1 1 II 1 1 1 581 gaattacatggtttactacaacttctttgcgttcgtgttactgccccttctgctcatgct 640 1241 aaggggtcatctggtccaggctagagtaggagcttcctcatggtcttcagagtttgcccc 1300 601 ggccatctacctacggatttttctggcggcccggagacagctgaagcagatggagagcca 660 1261 ttcctgagggaaagacattttaatatttttggttggctggaccaatctcactaagggaag 1320 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 11111111II1111111111 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 II 1 1 1 1 1 641 ggccatctacctacggatttttctggcggcccggagacagctgaagcagatggagagcca 700 1301 ttcctgagggaaagacattttaatatttttggttggctggaccaatctcactaagggaag 1360 1321 agaaacccaatgggc 1335 1 1 1 1 1 1 1 1 1 1 1 1 M 1 1361 I I I I I I I I I I I I I I I agaaacccaatgggc 1375 Fig. 30. Alignment of the rat mucosal and brain A2A receptor coding regions. The rat mucosal A2A receptor cDNA sequence (top strand; accession number: AF228684) is aligned with the corresponding region the rat brain receptor (bottom strand; accession number: S47609) 1 actactttctggtgtccctggcgacggcggacgtggctgtgggactcttcgccatcccct I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 235 actactttctggtgtccctggcgacggcggacgtggctgtgggactcttcgccatcccct 61 ttgccatcaccatcagcctgggcttctgcacggactttcacagctgcctcttcctcgcct I I I I I I I I I I II I I I I I I I I I I i I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 295 ttgccatcaccatcagcctgggcttctgcacggactttcacagctgcctcttcctcgcct 121 gcttcgtgctggtgctcacacagagctccatctttagcctcttggcggtggctgtcgacc I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 355 gcttcgtgctggtgctcacacagagctccatctttagcctcttggcggtggctgtcgacc 181 ggtatctggccattcgcgtcccgctcaggtataaaggtttggtcactggaacacgagcaa I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I II I I I I I I I I I I I I 415 ggtatctggccattcgcgtcccgctcaggtataaaggtttggtcactggaacacgagcaa 241 gagggatcatcgctgtcctctgggtccttgcctttggcattggactgactcctttcctgg I I I I I I I I II I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 475 gagggatcatcgctgtcctctgggtccttgcctttggcattggactgactcctttcctgg 301 gttggaacagtaaagaccgtgccaccagcaactgcacagaacctggggatggcatc 356 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 535 gttggaacagtaaagaccgtgccaccagcaactgcacagaacctggggatggcatc 590 60 294 120 354 180 414 240 474 300 534 B 319 61 379 121 439 181 499 241 559 301 619 361 679 421 739 481 799 aaagccaacaataccacgacgagtgccttgtggttgcaaatcacctacgtcaccatggag I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I aaagccaacaataccacgacgagtgccttgtggttgcaaatcacctacgtcaccatggag gctgccattggtctctgtgctgtagtgggcaacatgctggtcatctgggtggtcaagctg I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I gctgccattggtctctgtgctgtagtgggcaacatgctggtcatctgggtggtcaagctg aaccgcactctgaggaccaccaccttctatttcatcgtctccctagcactggc t jacatt I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I aaccgcactctgaggaccaccaccttctatttcatcgtctccctagcactggc a jacatt gctgttggggtgctggtcatacccttggccattgccgtcagcctggaggtccagatgcac I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I gctgttggggtgctggtcatacccttggccattgccgtcagcctggaggtccagatgcac ttctatgcotgccttttcatgtcctgtgtgcttctggtcttcacccatgcttccatcatg I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I II ttctatgcctgccttttcatgtcctgtgtgcttctggtcttcacccatgcttccatcatg tccttgctggccattgctgtagaccgatacctgcgagtcaagctgacagtcagatataga I I I I I II I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I tccttgctggccattgctgtagaccgatacctgcgagtcaagctgacagtcagatataga acggttaccactcaaagaagaatatggctattcctgggcctctgctggctagtgtccttt I I I II I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I II I I I acggttaccactcaaagaagaatatggctattcctgggcctctgctggctagtgtccttt ctggtgggactgacccccatgtttggctggaatagaaaagtgaccttagagctctctcaa I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I II I II I I I I I I I I I I I I I I I ctggtgggactgacccccatgtttggctggaatagaaaagtgaocttagagctctctcaa aacagctcc 489 I I I I I I II I aacagctcc 807 60 378 120 438 180 498 240 558 300 618 360 678 420 738 480 798 Fig. 31. Alignment of the rat mucosal and brain A2B and A3 cDNA sequences. The partial cDNA sequence of the rat mucosal A2B (A) and A3 (B) receptor was aligned with corresponding regions of the rat brain receptors. A: Top strand: gastric mucosal A2B receptor sequence (accession number: AF084241); Bottom strand: brain A2B receptor sequence (accession number: M91466). B: Top strand: gastric mucosal A3 receptor sequence (accession number: AF102804) ; Bottom strand: brain A3 receptor sequence (accession number: M94152). A silent mutation at position 174 bp of the mucosal A3 receptor cDNA sequence is detected. III. Quantification of gastric adenosine Ai and A2A receptor mRNA expression To study the changes in adenosine A\ and A2A receptor gene expression induced by various states of the stomach, competitive RT-PCR and Real-Time RT-PCR assays were developed. 1. Competitive RT-PCR Fig. 32 shows the results of a representative Ai and A2A receptor competitive RT-PCR assay which was performed to quantify Ai and A2A receptor mRNA levels in the rat striatum. This tissue has been shown to contain a moderate level of Ai receptor mRNA and an extremely high level of A2A receptor mRNA. In agreement with previous studies, the level of striatal A2A receptor mRNA (6.6xl07 copies/pg total RNA) was significantly greater than the level of Ai receptor mRNA (3.9xl06 copies/pg total RNA). See Fig. 32C. When the competitive RT-PCR samples are electrophoresed through an agarose gel, the density of the template band was shown to increase as the initial amount of competitor added was decreased. The Log [density ratio (corrected competitor band/template band)] plotted against the Log (initial competitor concentration) displayed a linear relationship with a R value of > 0.95, and a slope of approximately 1.0. The graphs generated by the Ai and A2A competitive RT-PCR assays were shown to have slopes of 1.0510.04, and 1.07+0.05, respectively, for all assays performed. An ideal slope of 1.0 indicates that the amplification efficiency of the template is equal to that of the competitor (Freeman et al., 1999). The R2 values for all assays performed in these studies were > 0.95. B E j£ 0.40 0 00 q 8 2- -0.20 'ui c: <l> 2. -0.40 "[competitor] -template "competitor y = 0.8516x- 1.0522 R! = 0.98 5 0.7 0.9 1.1/ 1.3 1.5 1.7 1.9 2.1 2 Log [intial competitor concentration (amol)] =0 0.75 -pH Log [intial competitor concentration (amol)] 100 < K o 4 a B Striatum Fig. 32. Striatal A, and A2A receptor gene expression measured by competitive RT-PCR. Results of a representative Aj (A) and A2A (B) receptor competitive RT-PCR assay performed using 2.5 pg of striatum total RNA and various amounts of A, and A2A RNA competitor as the template. See inset for competitor concentrations (amol) used. C: A, (filled bar) and A2A (empty bar) receptor mRNA expression in the striatum (* P < 0.05, n = 4) Similar mRNA measurements were also obtained regardless of the amount of PCR template used. Ai and A2A receptor mRNA measurements were not significantly altered by using equal volumes of undiluted, 2x, 4x, or 8x diluted cDNA as the template for PCR (Fig. 33). The results of representative Ai and A2A competitive RT-PCR assays performed using fundus, corpus, antrum and mucosa total RNA are shown in Fig. 34 and 35. Ai and A2A receptor mRNA levels in all gastric regions were lower than those measured in the striatum (Fig. 32C). The level of Ai receptor mRNA found in the corpus, antrum and mucosa were approximately 3-fold higher than in the fundus (Fig. 36). Among the gastric tissues, the highest level of A2A receptor mRNA was found in the corpus. The fundus, antrum and mucosa contained similar levels of A2A receptor mRNA, which were approximately 2 fold less than in the corpus. In the antrum and mucosa, the level of Ai receptor mRNA was significantly higher than the level of A2A receptor mRNA, but in the fundus and corpus these levels were not significantly different. 2. Real-Time RT-PCR Fig. 37A shows a representative amplification plot generated by a Real-Time RT-PCR assay for measuring adenosine Ai receptor mRNA. The corresponding standard curve is also shown in Fig. 37B. Results show that this assay was able to amplify and measure adenosine Ai receptor mRNA within a 7 log range of Ai receptor RNA standard concentrations. When the highest concentration of standard was used as the template (lxlO9 copies/pl), the reaction was affected by limiting reagents at cycle 22, and the plateau phase of the reaction was reached by cycle 26. However, this does not affect quantification since measurements were made during the exponential phase at the threshold cycle. The standard 0.75-, o n Q. 0.50-E S 0.25 a E o S £ -0.25-1 (A C 0) §> -0.50 u. 0.00--0.75-1 0.75 1.00 Log (initial A1 competitor concentration) Sample A^amol/ng total RNA) undiluted 0.82 2 x diluted 0.88 4 x diluted 0.79 8 x diluted 0.79 0.75-, •2 0.50-5 m a. £ a> ^* I 0.25-a> a E 8, o.oo-"o" £ -0.25 '55 c © w -0.50 o 00 1.25 _o 75J Log (initial A2A competitor concentration) Sample (amol/(ig total RNA) undiluted 0.57 2 x diluted 0.50 4 x diluted 0.50 8 x diluted 0.54 Fig. 33. Effect of template concentrations on quantification of A, and A2A receptor mRNA by competitive RT-PCR. A[ (A) and A2A (B) receptor competitive RT-PCR experiments were performed using undiluted (•), 2x (A), 4x (•), or 8x (•) diluted cDNA as the template. The cDNA samples were synthesized using a fixed amount (2.5 pg) of corpus total RNA and various amounts of A, or A2A receptor RNA competitor. 0.38 amol/|ig total RNA Log [intia! competitor concentration (amol)] B I 0.5 -0.25 2.0 amol/|ig total RNA Log [intial competitor concentration (amol)] 5 0.25 tj 0.00 & -Ol £ -0.50 c M 2, -0.75 00 o -1 -1.00 y= 1.15X-0.56 RJ = 0.99 1.2 amol/ng total RNA 1.6 amol/ng total RNA Log [intial competitor concentration (amol)] Log [intial competitor concentration (amol)] Fig. 34. Quantification of gastric A[ receptor gene expression by competitive RT-PCR. Results of representative competitive RT-PCR assays for the fundus (A), corpus (B), antrum (C) and mucosa (D) are presented. The A, receptor gene expression levels measured in these tissues were 0.38, 2.0, 1.2, and 1.6 amol/pg total RNA, respectively. Fig. 35. Quantification of gastric A2A receptor gene expression by competitive RT-PCR. Results of representative competitive RT-PCR assays for the fundus (A), corpus (B), antrum (C) and mucosa (D) are presented. The A2A gene expression levels measured in these tissues were 0.36, 0.68, 0.59 and 0.29 amol/pg total RNA, respectively. £ oo Fundus Mucosa: Antral/Corporeal 714 ±77* A,A: 288 ± 63 Fig. 36. Competitive RT-PCR quantification of A, and A2A receptor gene expression in the rat stomach. Adenosine receptor mRNA levels (in thousand copies/pg total RNA) are shown for each region. *P < 0.05 when A, receptor mRNA levels are compared to A2A receptor mRNA levels, n > 4. 1.600 0.000 -0.200 0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34 36 38 40 Cycle Number B i i inurn i iiiinn i inmn i 111iiHI i ilium iiiiuiii IIHUIII iiiiimi i mini) 10*2 10*3 10*4 10*5 10*6 10*7 10*8 10*9 10*10 A1 Receptor RNA Standard Concentration Fig. 37. Results of a representative A, receptor Real-Time RT-PCR assay. A: An amplification plot showing changes in normalized reporter emission after each cycle of PCR. Samples contained lxlO9 (•), lxlO8 ( ), lxlO7 ( ), lxlO6 (•), lxlO5 (•), lxlO4 (•), lxlO3 (•) initial copies of At receptor RNA. B: The corresponding standard curve generated by the amplification plot. The threshold cycle (CT) represents the cycle where the change in normalized reporter emission exceed the threshold. curve generated by plotting the threshold cycle against the Log(Ai RNA standard concentration) was found to be linear with a R value of 0.99. Fig. 38 shows the amplification plot and standard curve obtained for a representative A2A receptor Real-Time PCR assay. The standard curve was also linear within a 7 log range of A2A RNA standard concentrations, with a Rvalue of 0.98. Fig. 39 shows the absolute levels of adenosine Ai and A2A receptor mRNA measured in the striatum and in various regions of the stomach, including the fundus, corpus, antrum, mucosa, corporeal muscle and corporeal mucosa. In agreement with previous studies, adenosine Ai and A2A receptor levels were highest in the striatum (Dixon et al, 1996). Striatal Ai and A2A receptor mRNA levels were approximately 2-fold and 70-fold higher than the highest gastric Ai and A2A mRNA levels in the corporeal muscle region, respectively. In all gastric tissues, Ai receptor mRNA levels were significantly higher than A2A receptor mRNA levels. In the fundus, corpus, antrum, mucosa, corporeal muscle and corporeal mucosa, the Ai receptor mRNA levels were 3.4, 7.8, 8.3, 28.4, 14.4, and 14.6 times higher than A2A receptor mRNA levels, respectively. The highest and lowest levels of Ai receptor mRNA were found in the corporeal muscle and corporeal mucosa, respectively. No significant difference in the Ai receptor mRNA levels was observed among the corpus, antrum and the whole stomach mucosa. A2A receptor mRNA levels were not significantly different among the fundus, corpus and antrum, but were lowest in the whole gastric mucosa and in the corporeal mucosa. i i i i i i i i i i 2 4 6 8 I I I I I I I I I I I I I I I 10 12 14 16 18 20 22 24 26 28 30 32 34 36 38 40 Cycle Number B i i ilium i i II nm i MI nm iiiiiim iiiiiim iiniiiii MIIIIUI i in inn iiiimfi 10*2 10*3 10*4 10*5 10*6 10*7 10*8 10*9 10*10 A2A Receptor RNA Standard Concentration Fig. 38. Results of a representative A2A Real-Time RT-PCR assay. A: An amplification plot showing changes in normalized reporter emission after each cycle of PCR. Samples contained lxlO9 (•), lxlO8 ( ), lxlO7 ( ), lxlO6 (•), lxlO5 (•), lxlO4 (•), lxlO3 (•) initial copies of A2A receptor RNA. B: The corresponding standard curve generated by the amplification plot. The threshold cycle (CT) represents the cycle where the change in normalized reporter emission exceed the threshold. < z or 5 o E 3000 2500 2000 1500 D00 Striatum Fundus Corpus Antrum Mucosa Corporeal Corporeal muscle mucosa Striatum Fundus Corpus Antrum Mucosa Corporeal Corporeal muscle mucosa Fig. 39. Real-Time RT-PCR quantification of gastric and striatal adenosine A1 (A) and A2A (B) receptor gene expression. The level of A, and A2A receptor gene expression was highest in the striatum. A! receptor gene expression was also significantly higher than A2A receptor gene expression in all gastric tissues examined (n > 4). Note the difference in the scale of the ordinate. IV. Effect of fasting on adenosine receptor, somatostatin and gastrin gene expression To examine whether the prandial state of the stomach affects adenosine receptor gene expression, rats were fasted for either 24 or 36 h, and gene expression was quantified using Real-Time RT-PCR. Real-Time RT-PCR was utilized for this study since this technique is more sensitive and specific than competitive RT-PCR (Heid et al., 1996). In these experiments, gastrin and somatostatin gene expression were also measured by Real-Time RT-PCR to determine if changes in adenosine receptor gene expression occur with changes in gastric peptide gene expression. 1. Effect of fasting on body weight and Total RNA concentration Rats subjected to 24 and 36 h of fasting were shown to exhibit significant weight loss compared to control animals (see Table 8). Fasting for 36 h, but not 24 h, was also shown to significantly decrease the mass of the antral tissue. Corporeal mucosa and corporeal muscle mass were not affected by either 24 or 36 h of fasting. The total RNA concentration (pg total RNA/mg tissue) was also altered by fasting in some regions (see Table 8). Fasting for 24 and 36 h significantly decreased the total RNA concentration in the antrum, while fasting for 24 h significantly decreased total RNA concentration in the corporeal muscle. 2. Effect of fasting on 18S rRNA and GAPDH gene expression The expression levels of housekeeping genes were measured and used as the endogenous control to standardize the amount of sample added to each reaction. Two Table 8: Effect of fasting and omeprazole treatment on body weight, tissue weight and total RNA concentration Fasting Omeprazole 24 h 36 h 1 day 3 day Change in weight (g) Test -28±2* -35±1* 5±2 17±3 Control 6±1 13±1 6±2 18±2 Tissue weight (mg) Antrum Test 155±10 117±10* 121±10 163=1=17 Control 163±16 182±20 115=1=11 153±26 Corporeal Mucosa Test 55=1=4 69±11 63±4 74±6 Control 63±6 65±10 65±8 83±9 Corporeal Muscle Test 792±23 815±41 689±68 658±53 Control 804±41 804±40 634±17 665±56 Total RNA (M-g/mg tissue) Antrum Test 1.1±0.1* l.OiO.l* 2.2±0.2 1.7±0.3 Control 1.4±0.1 1.7±0.2 2.1±0.2 2.0±0.3 Corporeal Mucosa Test 2.7±0.3 2.4=1=0.5 2.3±0.6 2.7=1=0.3 Control 2.8±0.3 3.9±0.6 2.2±0.2 2.1±0.2 Corporeal Muscle Test 4.7±0.4* 4.7±0.2 5.9=1=0.3 4.1±0.3 Control 5.9±0.3 4.6±0.2 6.6±0.6 4.5=1=0.4 *P < 0.05 when test animals were compared to controls using the unpaired Student's t-test, n > 8 per group. commonly used housekeeping genes were measured, GAPDH and 18S rRNA. Results show that GAPDH mRNA levels were not significantly altered after 24 h of fasting in all gastric tissues examined. However, after 36 h of fasting, significant decreases in GAPDH mRNA levels were observed in the antrum (53% of control), corporeal mucosa (56% of control) and corporeal muscle (70% of control) (Fig. 40). The levels of 18S rRNA were not altered by 24 or 36 h of fasting in all tissues studied (See Fig. 40B). Therefore, 18S rRNA levels were used as the endogenous standard to control for differences in total RNA concentration and RNA integrity in the following studies. 3. Effect of fasting on adenosine Ai and A2A receptor expression Following a 24 h fast, Ai receptor mRNA levels in the corporeal mucosa and antrum were significantly increased to 134 ± 11% and 157 + 19% of control levels, respectively (Fig. 41A). No significant change in Ai receptor gene expression was observed in these tissues when the fasting period was extended to 36 h. In addition, fasting for 24 and 36 h did not alter Ai receptor mRNA expression in the corporeal muscle. The effect of fasting on A2A receptor mRNA expression differed between the corporeal mucosa and the antrum. In the corporeal mucosa, adenosine A2A receptor expression was not altered by 24 h of fasting. However, following 36 h of fasting, A2A receptor expression was significantly reduced to 65 ± 6 % of the control levels (Fig. 41B). In the antrum, the gene expression of adenosine A2A receptor was significantly increased to 173 ±21 % of control levels after a 24 h fast, but unchanged following a 36 h fast. Similar to Ai receptor gene expression, fasting did not alter A2A receptor gene expression in the corporeal muscle. Antrum Corporeal Mucosa Corporeal Muscle 200 c 150 o Antrum Corporeal Mucosa Corporeal Muscle Fig. 40. Effect of fasting on gastric housekeeping gene expression. GAPDH mRNA (A) and 18S rRNA (B) levels were measured in rats fasted for 24 h (grey) or 36 h (black). Results are expressed as a percentage of the control (white) as described in the Methods section. Statistics were performed using the Student's unpaired t-test. * P < 0.05; n > 7. Antrum Corporeal Mucosa Corporeal Muscle T T T T * 1 Antrum Corporeal Mucosa Corporeal Muscle Fig. 41. Effect of fasting on adenosine receptor gene expression. A[ receptor mRNA (A) and A2A receptor mRNA (B) levels were measured in rats fasted for 24 h (grey) or 36 h (black). Results are expressed as a percentage of the control (white) as described in the Methods section. Statistics were performed using the Student's unpaired t-test. * P < 0.05; n > 7. 4. Effect of fasting on gastrin and somatostatin gene expression Antral gastrin gene expression was significantly reduced to 50 ± 9% and 54 + 10% of control levels after 24 and 36 h of fasting, respectively (Fig. 42A). Following fasting, changes in somatostatin gene expression (Fig. 42B) resembled changes in A2A receptor gene expression (Fig. 41B). In the corporeal mucosa, somatostatin mRNA levels were not affected by 24 h of fasting, but significantly decreased (53 ± 5 % of controls) after 36 h of fasting. In the antrum, somatostatin gene expression was significantly augmented (196 ± 27% of controls) after 24 h of fasting, but unaltered after 36 h of fasting. Similar to A2A receptor gene expression, fasting did not affect somatostatin gene expression in the corporeal muscle. When the changes in somatostatin gene expression were compared to changes in adenosine A2A receptor gene expression, a linear relationship with a correlation coefficient of 0.96 was observed (Fig. 43). V. Omeprazole treatment on adenosine receptor and gastric peptide gene expression To examine if the inhibition of gastric acid secretion alters adenosine receptor gene expression, rats were treated with 400 pmol/kg omeprazole daily for 1 or 3 days. Omeprazole inhibits gastric acid secretion by binding irreversibly to the proton pump, H+K+ATPase, on parietal cells (Hirschowitz et al., 1995). Gastrin and somatostatin gene expression were also measured to examine if their expression is also altered with adenosine receptor gene expression. Antrum Corporeal Mucosa Corporeal Muscle Fig. 42. Effect of fasting on gastric peptide gene expression. Gastrin mRNA (A) and somatostatin mRNA (B) levels were measured in rats fasted for 24 h (grey) or 36 h (black). Results are expressed as a percentage of the control (white) as described in the Methods section. Statistics were performed using the Student's unpaired t-test. * P < 0.05; n > 7. Fig. 43. Comparison between fasting-induced changes in somatostatin and adenosine A2A receptor gene expression. Changes in gene expression were measured after 24 or 36 h of fasting in various gastric tissues: corporeal mucosa (36 h; blue), corporeal mucosa (24 h; yellow), corporeal muscle (24 h, green); corporeal muscle (36 h; red), antrum (36 h; purple), antrum (24 h; orange). Each point represents the mean ± SEM of at least 7 animals. 1. Effect of omeprazole treatment on 18S rRNA and GAPDH gene expression Omeprazole treatment for 1 or 3 days did not alter the weight of the rats, the mass of the gastric tissues, or the total RNA concentration of these tissues (see Table 8). Housekeeping gene expression levels were measured and used as the endogenous control. Similar to the effects of fasting, omeprazole treatment also altered GAPDH mRNA levels in some regions. GAPDH gene expression was significantly decreased to 65±5% of control levels in the corporeal mucosa after 3 days of treatment, and significantly increased to 130±9% in the corporeal muscle after 1 day of treatment (Fig. 44A). However, levels of 18S rRNA were not altered by 1 or 3 days of omeprazole treatment (Fig. 44B). Therefore, this housekeeping gene was used as the endogenous control for the Real-Time RT-PCR quantification of Ai and A2A receptor, somatostatin and gastrin mRNA levels. 2. Effect of omeprazole treatment on adenosine Ai and A2A receptor expression Omeprazole significantly decreased Ai receptor gene expression in the antrum to 66±12% and 47±8% of control levels after 1 day and 3 days of treatment, respectively (see Fig. 45A). In the corporeal mucosa, Ai receptor gene expression was significantly decreased to 62±6% of control levels after 1 day of omeprazole treatment, but changes were not apparent after 3 days of treatment. A2A receptor gene expression was significantly decreased to 57±7% of control levels in the antrum after 3 days of treatment (Fig. 45B). In the corporeal mucosa, A2A receptor gene expression was also decreased to 67±14% of control levels after 1 day of omeprazole treatment, but changes were not apparent 3 days after treatment. Both Ai and A2A receptor A 200 -"5 175 -C Antrum Corporeal Mucosa Corporeal Muscle Antrum Corporeal Mucosa Corporeal Muscle Fig. 44.: Effect of omeprazole treatment on gastric housekeeping gene expression. GAPDH mRNA (A) and 18S rRNA (B) levels were measured in rats treated with omeprazole for 1 (grey) or 3 days (black). Results are expressed as a percentage of the control (white), as described in the Methods section. Statistics were performed using the Student's unpaired t-test. * P < 0.05; n>7. Antrum Corporeal Mucosa Corporeal Muscle 200 i B _ 2 175-c o Antrum Corporeal Mucosa Corporeal Muscle Fig. 45. Effect of omeprazole treatment on adenosine A, and A2A receptor gene expression. Aj receptor (A) and A2A (B) receptor mRNA levels were measured in rats treated with omeprazole for 1 (grey) or 3 (black) days. Results are expressed as a percentage of the control (white) as described in the Methods section. Statistics were performed using the Student's unpaired t-test. * P < 0.05, ** P <0.01, ***P < 0.001 ; n > 7. gene expressions were not altered by either 1 or 3 days of omeprazole treatment in the corporeal muscle. 3. Effect of omeprazole treatment on gastrin and somatostatin gene expression Antral gastrin gene expression was significantly increased to 201 ± 18% and 268 ±42% of control levels after 1 and 3 days of omeprazole treatment, respectively (Fig. 46A). In contrast, antral somatostatin gene expression was significantly reduced to 71±6% and 58±4% of controls levels, and corporeal mucosal somatostatin gene expression was reduced to 54±13% and 54±10% of control levels after 1 day and 3 days of treatment, respectively (Fig. 46B). In the corporeal muscle, somatostatin gene expression was not altered by 1 day of treatment. However, after 3 days of treatment, somatostatin gene expression was significantly reduced to 45±6% of control levels. VI. Omeprazole on adenosine agonist-induced changes in IRG and SLI release The effect of omeprazole treatment on adenosine-agonist induced changes in IRG and SLI release was examined. After 1 or 3 days of omeprazole treatment, stomachs were perfused with either CPA (Ai-selective agonist) or CGS 21680 (A2A-selective agonist) at a concentration of 0.1 pM or 1 pM. The lower concentration of 0.1 pM was chosen based on the estimated EC50 value of CPA (0.067 pM) and CGS 21680 (0.06 pM) in inhibiting IRG and stimulating SLI release, respectively. The effect of 1.0 pM concentration was examined since this concentration of CPA and CGS 21680 elicited Antrum 200 Antrum Corporeal Mucosa Corporeal Muscle Fig. 46. Effect of omeprazole treatment on gastric peptide gene expression. Gastrin (A) and somatostatin (B) mRNA levels were measured in rats treated with omeprazole for 1 (grey) or 3 (black) days. Results are expressed as a percentage of the control (white), as described in the Methods section. Statistics were performed using the Student's unpaired t-test. * P < 0.05, ** PO.01, ***p<0.001 ;n>7. maximal inhibition and stimulation of IRG and SLI release, respectively (see Section I of this chapter). The effect of CGS 21680 on IRG release was not examined since this compound did not alter IRG release (see section 1.1). 1. Effect of omeprazole on CPA-induced changes in IRG release Fig. 47 shows the effect of 1 pM CPA on IRG release in animals treated with omeprazole for 3 days. The administration of CPA resulted in a significant inhibition of IRG release in both control and treated animals. However, there was no difference in the release of IRG between these two groups. Results of CPA-induced IRG release in omeprazole-treated and control animals are summarized in Fig. 48. There were no changes in CPA-induced IRG release between treated and control animals. 2. Effect of omeprazole on agonist-induced changes in SLI release Fig. 49 shows the effect of 1 pM CGS 21680 on SLI release in control and 3 day omeprazole-treated animals. In these experiments, SLI release was enhanced by CGS 21680 in both treated and control animals. However, the CGS 21680-induced SLI release was significantly attenuated by 3 day omeprazole treatment. Results of CGS 21680-induced SLI release in control and 1 or 3 day omeprazole-treated animals are summarized in Fig. 50. As described earlier, the administration of 0.1 pM CPA significantly inhibited gastric SLI release, while the administration of 1 pM CPA significantly stimulated SLI release (see Section I). These observations were also apparent in the current experiments (Fig. 51). The inhibition of SLI release induced by 0.1 pM CPA was not altered by either 1 or 3 days of 123456789 10 5 min periods B 200-— 150-01 <A I 100H r-i (9 50-CPA(1nM) 1 23456789 10 5 min periods Fig. 47. Effect of CPA (1 pM) on gastric IRG release from the perfused stomachs of 3 day omeprazole-treated (A) and control (B) animals. Results are expressed as IRG release (%) as described in the Methods section. Each column represents the mean ± SEM of at least 5 experiments. *P < 0.05 and ***P < 0.001 when compared with period 3 using repeated measures ANOVA followed by Dunnett's multiple comparison test. The IRG release during each 5 min period was not significantly different between omeprazole-treated and control animals. 50-45-40-35-30-25-20-15-10-5-X X CPA(uM): 0.1 1 1 day treatment 0.1 1 3 day treatment Fig. 48. Effect of omeprazole treatment on CPA-induced changes in IRG release. The percentage inhibition of IRG release between control (empty bar) and omeprazole-treated (filled bar) animals was compared using the Student's unpaired t-test, and were found not to differ. Each column represents the mean ± SEM of at least 5 experiments. — 1500-0) IA CO £ iooo-0) Of to 500-CGS(1 nM) i*** T J, — t 1 2345 6 789 10 5 min periods B 2000-— 1500-01 (A (0 0) "3 -S 1000-(0 500-CX3S(1nM) X D EL 123456789 10 5 min periods Fig. 49. Effect of CGS 21680 (CGS; 1 uM) on gastric SLI release in 3 day omeprazole-treated (A) and control (B) animals. Results are expressed as SLI release (%) as . described in the Methods section. Each column represents the mean ± SEM of at least 5 experiments. ***P < 0.001 when compared with period 3 using repeated measures ANOVA followed by Dunnett's multiple comparison test. The SLI release during each 5 min period was compared between omeprazole-treated and control rats using the Student's unpaired t-test and found to differ significantly in periods 6, 7, and 8; fP < 0.05. 1400-1200-1000-800-u a c I o 6* 600-• CO 400-I 200-X X X x 1 X CGS (|iM): 0.1 1 1 day treatment 0.1 1 3 day treatment Fig. 50. Effect of omeprazole treatment on CGS 21680-induced changes in SLI release. The percentage change in SLI release between the control (empty bar) and omeprazole-treated (filled bar) animals was compared using the Student's unpaired t-test. Each column represents the mean ± SEM of at least 5 experiments. *P < 0.05. 150-125-§ •g 75-8 5°-Si a DC 25-• 0---25 50_CPA(nM): 0.1 1 1 day treatment 0.1 T X 1 3 day treatment Fig. 51. Effect of omeprazole treatment on CPA-induced changes in SLI release. The percentage change in SLI release between the control (empty bar) and omeprazole-treated (filled bar) animals was compared using the Student's unpaired t-test. Each column represents the mean ± SEM of at least 5 experiments. *P < 0.05. omeprazole treatment. However, 3 days of omeprazole treatment was shown to significantly attenuate the stimulatory effect of 1 pM CPA on SLI release (Fig. 51). VII. Cellular localization of the gastric adenosine Ai and A2A receptors Immunohistochemistry experiments were performed to examine the cellular localization of the A\ and A2A receptors in the rat stomach, and confocal images of various layers of the corpus and antrum were acquired. Fig. 52 shows representative confocal images obtained from the mucosa, muscularis mucosa, submucosal plexus, circular muscle, myenteric plexus, and longitudinal muscle of the rat corpus and antrum. 1. Cellular localization and distribution of AjR-IR Fig. 53 and 54 show that AiR-IR is distributed throughout the corpus and antrum of the stomach. Abundant AiR-IR cells were dispersed throughout the corporeal mucosa, except at the tips of the mucosa (Fig. 53 A and B). In the antrum, AiR-IR cells were observed along the basal region of the antral mucosa (Fig. 54A and B). Distinct AjR-IR was also observed on cell bodies and nerve fibers of the myenteric plexus, and on nerve fibers of the submucosal plexus, circular muscle, and longitudinal muscle, on blood vessels localized throughout the stomach (Fig. 53 and 54: C & D) and on nerve fibers of the muscularis mucosae. Tissues stained with the immunoneutralized Ai receptor antibody did not exhibit any distinct AiR-IR (Fig. 53 and 54: E and F). Tissue sections stained with the secondary antibody, but without the prior incubation with Ai receptor primary antibody also demonstrated a lack of immunoreactivity (Fig. 53G and 54G) Fig. 52. Representative A,R-IR and A2AR-IR in various regions of the rat stomach. A diagram representing a cross-sectional view of the stomach wall is shown with the following layers identified: mucosa (M), muscularis mucosae (MM), submucosal plexus (SMP), circular muscle (CM), myenteric plexus (MYP), longitudinal muscle (LM), and serosa (S). Typical confocal images taken from the regions highlighted by the boxes are shown. A: A,R-IR in the basal region of the antral mucosa; B: A,R-1R in the submucosal plexus of the antrum; C: A2AR-IR in the circular muscle of the corpus; D: A2AR-IR in the myenteric region of the antrum. Fig. 53. Confocal images of A,R-IR in the corpus of the rat stomach. A: A,R-IR cells (arrows) were shown to be distributed throughout the corporeal mucosa. B: Intense A,R-IR was observed on mucosal cells viewed under high magnification (arrows). C: AjR-IR on endothelial cells lining a blood vessel (BV; arrowhead) and on nerve fibers (arrows) of the submucosal plexus (SMP). D: A,R-IR on nerves fibers and cell bodies of the myenteric plexus (arrowhead), and on nerve fibers of the circular (CM) and longitudinal (LM) muscle layers (arrows) of the corpus. Tissue sections stained with the immunoneutralized A, receptor primary antibody in muscle (E) and mucosal (F) layers. Tissue section stained with the secondary antibody, donkey anti-rabbit IgG conjugated to Cy3, but without prior incubation with A( receptor primary antibody (G). (z-step = 1.0 pm, scale bars = 25 pm for all images). CM LM Fig. 54. Confocal images of A,R-IR in the antrum of the rat stomach. A: A,R-IR cells (arrows) were shown to be concentrated along the base of the antral mucosa (MM = muscularis mucosae). A high magnification image of the boxed area is shown in B. Note the punctate pattern of staining. C: A,R-IR on nerve fibers (arrowheads) of the submucosal plexus (SMP), circular muscle (CM) and muscularis mucosae. D: A,R-IR on nerves fibers and cell bodies of the myenteric plexus (arrowheads), and on nerve fibers of the circular and longitudinal (LM) muscle layers (arrows). Tissue sections stained with the immunoneutralized A, receptor primary antibody in muscle (E) and mucosal (F) layers. Tissue section stained with the secondary antibody, donkey anti-mouse IgG conjugated to Alexa Fluor 488, but without the prior incubation with A, receptor primary antibody (G). (z-step = 1.0 pm, scale bars = 25 pm for all images). 2. Double staining with the Ai receptor antibody Double staining experiments were performed to examine if the Ai receptors are localized on gastrin-secreting G-cells. Results show that gastrin-IR was abundant throughout the base of the antral mucosa. Cells were flask-shaped and some were shown to project short processes. Double staining revealed that all gastrin-IR cells examined expressed AiR-IR (Fig. 55). AiR-IR was most intense in antral mucosal cells that express gastrin-IR. However, not all AiR-IR cells contained gastrin-IR. Quantification revealed that approximately 28±2% of AiR-IR cells in the antral mucosa also expressed gastrin-IR. Double staining of AjR-IR with somatostatin-IR was performed to examine whether Ai receptors are localized on D-cells of the corpus and antrum or somatostatin-containing nerves throughout the stomach. Intense and abundant somatostatin-IR was observed in the antral and corporeal mucosa, while sparse somatostatin-IR was observed throughout the rest of the stomach. Double staining revealed that all somatostatin-IR cells of the corporeal mucosa expressed AiR-IR (Fig. 56A-C). AiR-IR staining was especially intense at the processes of the D-cells (Fig. 56D-F). In the antrum, only 24±2% of the somatostatin-IR cells contained AiR-IR (Fig. 56G-I). Although a relatively large proportion of the AjR-IR of the antral mucosa is co-localized with gastrin-IR, only 8.5±0.6 % of the AiR-IR cells in this region was co-localized with somatostatin-IR. Ai receptor-IR was not co-localized with somatostatin-IR in any other region of the stomach, including the myenteric plexus, region, where sparse somatostatin staining was observed (see Fig. 57). To determine whether the Ai receptors are localized on the parietal cells, double staining was performed to detect H+K+ATPase (3, a marker for parietal cells. Dense and Fig. 55. Double staining of A,R-IR with gastrin-IR. Confocal images show co-localization of A,R-IR (A) with gastrin-IR (B) in the antral mucosa. Cells containing both immunoreactivities appear yellow in the merged image (C). All gastrin-IR cells contain A,R-IR (arrows). A,R-IR cells not expressing gastrin-IR are identified by arrowheads (z-step = 0.5 pm; scale bar = 25 pm for all images). Fig. 56. Double staining of A,R-IR with somatostatin-IR in the gastric mucosa. Confocal images show co-localization of A,R-IR (red) with somatostatin-IR (green) in the rat gastric corporeal (A-F) and antral (G-I) mucosa. Cells expressing both A,R-IR and somatostatin-IR are indicated by arrows and appear yellow (C, F, & I). AtR-IR is expressed on all somatostatin-IR cells of the corpus (A-C), and appears concentrated at the processes of the D-cell (D-F). A,R-IR is also co-localized with somatostatin-IR in some cells in the antral mucosa (G-I). Somatostatin-IR cells not expressing A,R-IR are indicated by the arrowheads, (z-step = 0.5 pm; scale bars = 25 pm for all images). A « 1 f * C m LM ^ 1 CM -f n F LM / ^ CM Fig. 57. Double staining of A,R-IR and somatostatin-IR in the gastric muscle. Confocal images show the lack of co-localization of A,R-IR (red; large arrowheads) with somatostatin-IR (green; small arrowheads) in the myenteric plexus of the rat corpus (A-C) and antrum (D-F). LM = longitudinal muscle, CM = circular muscle, (z-step of confocal images = 0.5 pm; scale bars = 25 pm). o intense H+K+ATPase p staining was observed throughout the corporeal mucosa. Double-staining of H+K+ATPase P-IR with AiR-IR, however, demonstrated a lack of co-localization (Fig. 58). To determine with certainty whether AiR-IR are localized on blood vessels, double staining was performed to detect VWF, a marker for the vascular endothelial cells. AiR-IR was found to be co-localized with VWF in both the corpus (Fig. 59A-C) and the antrum (Fig. 59D-F). However, AiR-IR was not co-localized with VWF-IR in the mucosa, except in arterioles at the base of the mucosa (Fig. 59G-I). Careful analysis of single confocal images (z-step 1.0 pm) revealed that some AjR-IR was also observed in close proximity to VWF-IR but was not co-localized with VWF-IR (Fig. 60). This AiR-IR likely represents Ai receptors on vascular smooth muscle cells. Double staining for PGP 9.5 was performed to determine whether AiR-IR is expressed on nerve fibers. PGP 9.5-IR was found to be co-localized with AiR-IR on cell bodies and nerve fibers of the myenteric plexus of the corpus (Fig. 61A-C) and antrum (Fig. 61D-F). PGP-9.5-IR was also co-localized with AiR-IR on nerve fibers throughout the muscle layers of the corpus (Fig. 61G-I) and antrum (Fig. 61J-L). 3. Cellular localization and distribution of A2AR-IR Results show that the distribution of A2AR-IR was similar in the corpus and antrum (Fig. 62 and 63). In both regions, intense A2A receptor staining was observed on mucosal cells (Fig. 62A and 63 A), on nerve fibers of the submucosal plexus (Fig. 62B and 63B), on cell bodies and nerve fibers of the myenteric plexus, and on nerve fibers of the circular muscle layer, longitudinal muscle layer (Fig. 62C and 63C) and muscularis mucosae. Fig. 58. Double-staining of A,R-IR with H+K+-ATPase p-IR. A,R-IR (A) is shown not to co-localize with H+K+-ATPase P-IR (B) i the corporeal mucosa, (z-step of confocal images = 0.5 pm; scale bar = 25 pm). to Fig. 59. Double staining of A,R-IR (red) with VWF-IR (green). Images show the co-localization of A,R-IR with VWF-IR in the circular muscle (CM) of the corpus (A-C) and antrum (D-F). The arrowheads (D-F) identify A,R-IR in the myenteric plexus. Regions expressing both immunoreactivities appear yellow. AtR-IR is not co-localized with VWF-IR in the fine capillaries of the corporeal mucosa (G-I). Images shown in A-C, D-F and G-I represent the immunoreactivity of 1, 20, and 36 stacked images (z-step = 1.0 pm), respectively. Images were stacked to adequately illustrate the immunoreactivity within the entire blood vessel or capillary bed. Scale bars = 25 pm. o c Fig. 60. Localization of A,R-IR (red) relative to VWF-IR (green) in the corporeal muscle. Images shown in A-C and D-F represent the immunoreactivity of 31 stacked sections, and a single section (z-step = 1.0 pm), respectively. A,R-IR is co-localized with VWF-IR, and is also found adjacent to VWF-IR. The A,R-IR indicated by the arrows on panel D may represent A,R localized on on vascular smooth muscle cells. (Scale bars = 50 pm). r Y+4r • -« £ lE \ \ v CM ^ J v ? K v y * L V / Fig. 61. Double staining of A,R-IR (red) with PGP 9.5-IR (green). Confocal images show the co-localization of both immunoreactivities in the myenteric plexus of the corpus (C and I, arrowheads) and antrum (F and L, arrowheads), and in the circular muscle of the corpus (I) and antrum (L). Cells expressing both immunoreactivities appear yellow in the merged images, (z-step = 0.5 pm for all images, scale bars = 25 pm). B LM Fig. 62. Confocal images of A2AR-IR in the corpus of the rat stomach. A: A2AR-1R cells (arrows) were distributed throughout the corporeal mucosa. B: A2AR-IR on nerve fibers in the submucosal plexus (SMP, arrows) and circular muscle (CM, arrowheads). C: A2AR-IR on nerves fibers and cell bodies of the myenteric plexus (arrows), and on nerve fibers of the circular and longitudinal (LM) muscle layers (arrowheads). Tissue sections stained with the immunoneutralized A2A receptor primary antibody in the mucosa (D) and muscle (E) layers. Tissue section stained with the secondary antibody, donkey anti-rabbit IgG conjugated to Cy3, but without prior incubation with A2A receptor primary antibody (F). (z-step = 1.0 pm, scale bars = 25 pm for all images). Fig. 63. Confocal images of A2AR-IR in the antrum of the rat stomach. A: A2AR-IR cells (arrows) were distributed throughout the antral mucosa, MM = muscularis mucosae. B: A2AR-IR on blood vessels (large arrowheads) and nerve fibers (small arrowheads) of the submucosal plexus (SMP), and on nerve fibers of the circular muscle (CM; arrows). C: A2AR-IR on nerves fibers and cell bodies of the myenteric plexus (arrows), on nerve fibers of the circular (CM) and longitudinal (LM) muscle layers (small arrowheads) and on blood vessels of the circular muscle (large arrowheads). Tissue sections stained with the immunoneutralized A2A receptor primary antibody in the mucosal (D) and muscle (E) layers. Tissue section stained with the secondary antibody, donkey anti-mouse IgG conjugated to Alexa Fluor 488, but without prior incubation with A2A receptor primary antibody (F). (z-step = 1.0 pm, scale bars = 25 pm for all images). A2AR-IR was also observed on blood vessels of both the corpus and antrum. The specificity of the A2A receptor antibody was confirmed by immunoneutralization. Tissues stained with the immunoneutralized A2A receptor antibody were shown not to exhibit A2AR-IR (Fig- 62 and 63: D and E). Tissue sections stained with the secondary antibody, but without the prior incubation with A2A receptor primary antibody also demonstrated a lack of A2AR-IR immunoreactivity (Fig. 62F and 63F). 4. Double staining with the A2A receptor antibody Double staining was performed to examine if the A2A receptors are localized on the D-cells, G-cells, parietal cells, neuronal cells and vascular endothelial cells. Somatostatin-IR was abundant in the mucosa, but sparse in other layers of the stomach. Co-localization of A2AR-IR with somatostatin-IR was frequently observed on both corporeal (Fig. 64A-C) and antral mucosal cells (Fig. 64D-F). Quantification of this co-localization demonstrated that 33±4% and 32±8% of A2AR-IR cells expressed somatostatin-IR in the corporeal and antral mucosa, respectively. On rare occasions, A2AR-IR was also shown to co-localize with somatostatin-IR in the myenteric plexus and muscle layers of the corpus (Fig. 65) and antrum (Fig. 66). The co-localization of A2AR-IR with gastrin-IR was not observed in any region of the antral mucosa (Fig. 67). A2AR-IR was also not co-localized with H+K+ATPase P-IR in the corporeal mucosa (Fig. 68). However, extensive co-localization of A2AR-IR with VWF-IR was observed throughout the corporeal and antral muscle layers, and the myenteric and submucosal plexi (Fig. 69). A2AR-IR was not co-localized with VWF-IR in the gastric mucosa. Analysis of single confocal images (z-step 1.0 pm) also reveals that A2AR-IR is localized adjacent to VWF-IR with a pattern similar to that of VWF-IR (Fig. 70). This staining may represent A2AR-IR on vascular smooth muscle cells. Double staining of A2AR-IR with PGP 9.5-IR was also observed in cell bodies and nerve fibers of the myenteric plexus, nerve fibers of the submucosal plexus, circular and longitudinal muscle layers, and muscularis mucosae (Fig. 71). A B c > y V - * . v Fig. 64. Double staining of A2AR-IR with somatostatin-IR in the gastric mucosa. Confocal images showing the co-localization of A2AR-IR (red) with somatostatin-IR (green) in the rat corporeal (A-C) and antral (D-F) mucosa. Cells expressing both A2AR-IR and somatostatin-IR are indicated by arrows and appear yellow (C & F). Somatostatin-IR cells not expressing A2AR-IR are indicated by the arrowheads, (z-step = 0.5 pm, scale bars = 25 pm for all images). o A \ B Si C \ • %r P D E V F fk. - • •" •• •- A If" * * CM " i Fig. 65. Double staining of A2AR-IR with somatostatin-IR in the gastric muscle of the corpus. Confocal images show co-localization of A2AR-IR (red) with somatostatin-IR (green) in the myenteric plexus (A-C) and circular muscle (D-F) of the rat corpus. Co-localization does occur (yellow; arrows); however, this was only observed occasionally. CM = circular muscle, (z-step = 0.5 pm, scale bars = 25 pm). A ^ ( u B c LM ^ D E \ F \ CM Fig. 66. Double staining of A2AR-IR with somatostatin-IR in the gastric muscle of the antrum. Confocal images show co-localization of A2AR-IR (red) with somatostatin-IR (green) in a cell body of the myenteric plexus (A-C) and nerve fibers in the circular muscle (D-F) of the rat antrum. Co-localization was shown to occur (yellow), however this was not frequent. LM = longitudinal muscle, CM = circular muscle, (z-step = 0.5 pm, scale bars = 25 pm). to A 1 \ V * 1 c I / i r * 1 Fig. 67. Double staining of A2AR-IR with gastrin-IR. Confocal images show the lack of co-localization of A2AR-IR (A) with gastrin-IR (B) in the antrum. Cells containing A2AR-IR (arrows) do not express gastrin-IR (arrowheads), (z-step = 0.5 pm; scale bars = 25 pm). Fig. 68. Double-staining of A2AR-IR (red) with H+K+-ATPase pMR (green). A2AR-IR (A; arrow) is shown not to co-localize with H+K+-ATPase (3-IR (B; arrowhead) in the corporeal mucosa, (z-step = 0.5 pm; scale bar = 25 pm). Fig. 69. Double staining of A2AR-IR (red) with VWF-IR (green). Confocal images show the co-localization of A2AR-IR with VWF-IR in the myenteric plexus region of the corpus (A-C; arrows) and in the circular muscle of the antrum (D-F; arrows). Regions expressing both immunoreactivities appear yellow, (z-step = 1 pm; scale bars = 25 pm). Fig. 70. Localization of A2AR-IR relative to VWF-IR in the corporeal muscle. Images shown in A-C represent the immunoreactivity of 16 stacked sections. Images shown on D-F and G-I represent the immunoreactivity taken from a single section (z-step =1.0 pm). A2AR-IR (red) is localized with VWF-IR (green), and is also found to be expressed in close proximity to VWF-IR. The A2AR-IR indicated by the arrows on panel D and G may represent A2A receptors localized on on vascular smooth muscle cells, (scale bar s= 50 pm). A \ °1 % C . , LM CM \V, Y , § $ is E > | LM > v j CM Fig. 71. Double staining of A2AR-IR (red) with PGP 9.5-IR (green). Confocal images show the co-localization of A2AR-IR with PGP 9.5-IR in nerve fibers of the myenteric plexus and circular muscle of the corpus (A-C) and antrum (D-F) (arrows). Cells expressing both immunoreactivities appear yellow (C and F). (CM = circular muscle; LM= longitudinal muscle; z-step = 0.5 pm; scale bars = 25 pm.). or. -J Chapter 4: Discussion This study is the first to establish the significant roles of adenosine Ai and A2A receptors in modulating gastric somatostatin and gastrin release. Biological and immunohistochemical results suggest that adenosine may inhibit IRG release, at least in part, by activating Aj receptors on the G-cells and may alter SLI release by activating both Ai and A2A receptors on the D-cells. Results of the molecular biological experiments further demonstrate that the adenosine Ai and A2A receptors are structurally identical to those in the brain, and are expressed in all functionally and morphologically distinct regions of the stomach. The expression and function of these receptors can be altered by different physiological states of the stomach, suggesting that adenosine receptor expression may be changed under different gastric secretory states. This discussion is organized into the following sections: Section I, the adenosine receptor subtypes involved in regulating gastric IRG and SLI release; Section II, the distribution of adenosine receptor mRNA in functionally distinct regions of the stomach and the structure of the gastric adenosine receptors; Section III, the quantitative RT-PCR techniques developed to measure gastric adenosine receptor gene expression; Section TV, the changes in Ai and A2A receptor gene expression and function by alterations in the physiological states of the stomach; Section V, the significant implications of the cellular localization and distribution of gastric Ai and A2A receptor, as illustrated by immunohistochemistry experiments. I. Alteration of IRG and SLI release by selective adenosine analogs While the effect of adenosine on inhibiting IRG release (DeSchryver-Kecskemeti et al, 1981; Harty and Franklin, 1984; Kwok et al., 1990) and stimulating SLI release (Kwok et al., 1990) has been described, the adenosine receptor subtypes involved have not been identified with certainty. The present study, therefore, was performed to elucidate the adenosine receptor(s) involved in these actions. Using adenosine receptor subtype-selective analogs, the present study demonstrates that the inhibition of IRG release was mediated by the activation of adenosine Ai receptors, while the release of SLI was inhibited and stimulated by the activation of the Ai and A2A receptors, respectively. 1. Effect of adenosine receptor-selective analogs on IRG release To determine the adenosine receptor subtype involved in the inhibition of IRG release, the effect of various adenosine analogs was examined in the isolated vascularly perfused rat stomach preparation. This preparation has been used in our earlier investigation of the effect of adenosine on IRG release (Kwok et al., 1990). Adenosine was found to suppress IRG release concentration-dependently. In the present study, only the administration of CPA significantly inhibited IRG release. CPA is a potent Ai receptor agonist that exhibits a greater binding affinity for the A\ receptor than the A2 and A3 receptors (Klotz et al., 1998; Ji and Jacobson, 1999). The administration of CGS 21680 and IB-MECA did not significantly alter basal IRG release. CGS21680 is a potent A2A receptor agonist, which exhibits a 140-fold selectivity for A2A receptors over Ai receptors (Hutchison et al., 1989), and IB-MECA is a potent A3 receptor agonist (Gallo-Rodriguez et al., 1994) that exhibits a greater affinity for A3 receptors than Ai and A2 receptors. CPA significantly inhibits basal IRG release starting at a concentration of 1 nM. However, CGS 21680 and IB-MECA were unable to elicit an effect even at a concentration of 100 nM. At this concentration, both analogs have been shown to potently elicit various A2A and A3 receptor-mediated actions in the rat (Hutchison et al., 1989; Zucchi et al., 2001). The potent inhibitory effect of CPA and the lack of effect of CGS 21680 and IB-MECA suggest that the Ai receptor is likely the subtype involved in modulating IRG release. To further test this hypothesis, the effect of selective Ai and A2 receptor antagonists on adenosine-induced inhibition of IRG release was examined. DPCPX, a selective Ai receptor antagonist that exhibits a 700-fold preference for the Ai receptor over the A2 receptor (Bruns et al., 1987), completely abolished the inhibition of IRG release induced by adenosine. DMPX, an A2 receptor-selective antagonist (Seale et al., 1988), however, did not alter either basal or adenosine-induced IRG release. The complete blockade of the inhibitory effect of adenosine by DPCPX and the lack of an effect by DMPX further support the suggestion that the Ai receptor subtype is responsible for the inhibition of IRG release. Results of these experiments do not identify the site at which adenosine acts to produce its inhibitory effect, but several sites are suggested. Using immunohistochemistry, the present study demonstrates that Ai receptors were localized on nerve fibers throughout the enteric plexi and on gastrin-containing G-cells. In the enteric plexi, Christofi and co workers have suggested that these receptors reside in presynaptic nerve terminals and function to modulate neurotransmitter release (Christofi et al., 1992). In addition, adenosine has been shown to inhibit the release of acetylcholine and noradrenaline in the guinea pig enteric plexi (Barajas-Lopez et al., 1991; Christofi et al., 1992), most likely through activation of Ai receptors at the presynaptic nerve terminal (Nitahara et al., 1995). In the isolated vascularly perfused rat stomach, IRG release is regulated by cholinergic and adrenergic agents (Koop et al., 1982, 1983). Thus, adenosine may regulate IRG release indirectly by modulating neurotransmitter release in the enteric plexi. Evidence also suggests that adenosine may regulate IRG release by acting directly on the G-cells. In rat antral mucosal fragments, adenosine inhibits basal and carbachol-stimulated gastrin release (Harty and Franklin, 1984). Adenosine has also been shown to inhibit forskolin-stimulated gastrin release in isolated canine G-cells (Schepp et al., 1990). Furthermore, the use of subtype-selective analogs specifically implicated the Ai receptors in mediating this inhibitory effect (Schepp et al., 1990). Additional support for a direct action on G-cells is provided by the present immunohistochemistry experiments which demonstrate intense Ai receptor staining on all antral G-cells of the rat stomach. The presence of adenosine A2A receptors on G-cells was not detected. This observation lends support to the biological experiments, which demonstrated the lack of effect of CGS 21680 on IRG release. In summary, these results clearly show that the adenosine Ai receptor subtype is involved in suppressing IRG release in the rat. In addition, I propose that adenosine may regulate IRG release directly by acting on the G-cells, and/or indirectly by modulating neurotransmitter release in the enteric plexi. 2. Effect of adenosine receptor-selective analogs on gastric SLI release The adenosine receptor subtype(s) involved in regulating SLI release was also examined by studying the effect of various adenosine analogs. In the isolated vascularly perfused rat stomach, our laboratory has demonstrated that adenosine may inhibit or augment gastric SLI release depending on the concentration used (Kwok et al., 1990). Results of the present study suggest that the stimulatory effect of adenosine is most likely mediated by activation of the A2A receptor. The rank order of potency of adenosine analogs in augmenting SLI release was shown to be: CGS 21680 > IB-MECA > CPA. At 1 \M, CGS 21680, an A2A receptor-selective agonist, was shown to elicit an increase in SLI release (% change) that was approximately 26-fold and 7-fold greater than the increase elicited by the same concentration of CPA and IB-MECA, respectively. CGS 21680 potently stimulated gastric SLI release with an EC5o of 0.06 \xM. Although CGS 21680 is selective for the A2A receptor, it can act on both A2A and A2B receptors. The stimulatory effect of CGS 21680 on SLI release, however, is not likely due to the activation of A2B receptors since this drug has a much lower affinity for the A2B receptors (Brackett and Daly, 1994). In the present study, the EC50 of CGS 21680 in augmenting SLI release was found to be in the sub-micromolar range (0.06 uM), as suggested for A2A receptor-mediated actions (Brackett and Daly, 1994). In PC 12 membranes expressing A2A receptors, the stimulation of adenylate cyclase activity by CGS 21680 was characterized by an EC50 of 0.07 pM (Brackett and Daly, 1994). In NIH 3T3 membranes expressing A2B receptors, 100 \xM and 3 \xM CGS 21680 was shown have little effect on adenylate cyclase activity and cAMP production, respectively (Brackett and Daly, 1994; Klotz et al., 1998). Thus, actions mediated by the A2A and A2B receptors can be differentiated by the potency of CGS 21680 in eliciting those effects. The sub-micromolar EC50 value of CGS 21680 in augmenting SLI release suggests the involvement of A2A receptors, but not A2B receptors. The involvement of A2A receptors is also supported by the effect of various selective adenosine antagonists on adenosine and adenosine agonist-induced SLI release. When administered at 1 \xM, DMPX, an A2 receptor antagonist, completely abolished the stimulatory effect elicited by CPA (1 \xM), and significantly attenuated IB-MECA- (lpM) and CGS 21680- (0.01 and 0.1 uM) stimulated SLI release. The inhibitory effect of DMPX on CGS 21680- (0.01 and 0.1 pM) induced SLI release was concentration-dependent. To ascertain whether the A2A receptors are involved in the augmentation of SLI release, the effect of the potent non-xanthine A2A receptor antagonist ZM 241385 (Poucher et al., 1995) on SLI release was tested. This A2A receptor-selective antagonist was able to completely abolish the stimulatory effect of CGS 21680 and adenosine on SLI release when administered at a concentration of 1 uM. This compound is 6,700-fold more selective for A2A receptors than Ai receptors (Poucher et al., 1995), while DMPX is only 4-fold more selective for A2 receptors than Ai receptors (Seale et al., 1988). These results support the proposal that A2A receptors are involved in stimulating SLI release. The Ai receptors are not likely involved in the augmentation of SLI release since DMPX and ZM 241385 completely abolished CPA-induced SLI release. Furthermore, the Ai receptor-selective antagonist, DPCPX, did not alter the effect of CGS 21680 on SLI release. The involvement of the A3 receptor in augmenting SLI release is also unlikely since the stimulatory effect induced by IB-MECA was sensitive to blockade by DMPX and was completely abolished by ZM 241385. In addition, studies have shown that adenosine-induced SLI release can be abolished by the administration of the non-selective antagonist 8-PT (Kwok et al., 1990). Since the rat A3 receptors were shown to be resistant to blockade by xanthine derivatives (Zhou et al., 1992) and A3 receptor-mediated effects in the rat were resistant to blockade by 8-PT (van Galen et al., 1994; Peachey et al., 1996), the involvement of A3 receptors in augmenting SLI release is unlikely. The stimulatory action of CPA and IB-MECA on SLI release, thus, may be due to their non-specific effect on A2A receptors when administered at high concentrations. Potent A2B receptor-selective antagonists were not available, so it is not possible to thoroughly examine the role of the A2B receptor in augmenting SLI release. However, the EC50 of CGS 21680 in eliciting this effect strongly suggests that the A2B receptors are not involved. Thus, the augmentation of gastric SLI release by adenosine is most likely mediated solely by the A2A receptor subtype. In addition to the stimulatory effect of adenosine on SLI release, studies have shown that at low concentrations (10 nM), adenosine can also significantly inhibit SLI release (Kwok et al., 1990). Similar results were obtained in the present experiments, in which a low concentration of CPA (0.1 pM) was shown to significantly inhibit SLI release. It is possible that CPA acts specifically on A] receptors to inhibit SLI release at low concentrations, while this compound acts non-specifically on A2A receptors to stimulate SLI release at higher concentrations. This proposal agrees well with studies demonstrating that, in tissues containing several adenosine receptor subtypes, Ai receptors are preferentially activated by lower levels of adenosine. In the rat, studies have shown that adenosine elicits Aj-, A2A-, A2B-, and A3- mediated effects with EC50 values of 73, 150, 5100, and 6500 nM, respectively (Daly and Padgett, 1992; Zhou et al., 1992; Peakman and Hill, 1994). The present proposal that the inhibition of SLI release is mediated by Ai receptors also explains the observation that basal SLI release was lower in the presence of ZM 241385 and adenosine than in the presence of ZM 241385 alone. When administered alone, ZM 241385 may inhibit SLI release by blocking the effect of endogenous adenosine on the A2A receptors, thereby unmasking the Ai receptor-mediated inhibition of SLI release. When ZM 241385 is administered together with adenosine, adenosine will only act on Ai receptors to inhibit SLI release. Consequently, the inhibitory effect of adenosine is enhanced in the presence of ZM 241385. These results demonstrate that adenosine may play a dual role in the regulation of SLI release in the rat. Activation of Ai and A2A receptors may result in inhibition and stimulation of SLI release, respectively. Results of the perfusion experiments do not clearly identify where adenosine acts to regulate SLI release, but results from the immunohistochemistry experiments suggest several possibilities. These studies demonstrate that Ai and A2A receptors were expressed in the gastric vasculature, the enteric plexi, and D-cells. Previous studies performed in our laboratory have shown that the stimulatory effect of adenosine on SLI release is not likely due to its vasodilatory action in the stomach. Like CGS 21680, nitroprusside, a vasodilator, was found to decrease perfusion pressure in the isolated vascularly perfused rat stomach. However, nitroprusside was shown not to alter SLI release (Yip and Kwok, 2004). The presence of A2A receptors on enteric nerves suggests that adenosine may alter SLI release by regulating neurotransmitter release since cholinergic and p-adrenergic agents are capable of modulating gastric SLI release (Saffouri et al., 1980; Koop et al., 1983). Earlier work performed in our laboratory has demonstrated that the cholinergic blockers, atropine and hexamethonium, and the P-adrenergic blocker, propranolol, did not alter adenosine-induced changes in SLI release (Kwok et al., 1990). Thus, it is unlikely that adenosine stimulates SLI release through A2A receptor-mediated changes in acetylcholine and noradrenaline release. The possibility that adenosine may alter gastric SLI release through non-cholinergic and non-adrenergic neurotransmission cannot be eliminated since both Ai and A2A receptors were localized on nerve fibers throughout the enteric plexi and muscle layers. The enteric peptides CGRP (Ren et al., 1993; Manela et al., 1995) and VIP (Chiba et al., 1980) can stimulate the release of gastric SLI. Thus, the possibility that adenosine modulates the release of these peptides to subsequently enhance SLI release remains to be examined. However, the localization of both Ai and A2A receptors on the corporeal and antral somatostatin-secreting D-cells strongly suggests that adenosine can regulate SLI release, at least in part, by acting directly on the gastric D-cells. The presence of both receptors on the D-cells further lends support to our proposal that adenosine may inhibit and stimulate SLI release by acting on the Ai and A2A receptors, respectively. 3. Regulation of gastric SLI and IRG release by adenosine Results from the present study suggest that endogenous adenosine may be involved in regulating basal SLI release. The administration of DMPX and ZM 241385 significantly inhibited basal SLI release, suggesting that basal SLI release may be maintained by activation of A2A receptors. When EHNA, an ADA inhibitor (Mendelson et al., 1983), was administered, both basal and adenosine-stimulated SLI release were enhanced. ADA is expressed in the gastric epithelia (Witte et al., 1991) and is responsible for metabolizing adenosine to inosine (Geiger et al., 1997). The enhancement of basal and adenosine-stimulated SLI release by EHNA suggests that ADA may play a role in regulating endogenous adenosine levels in the rat stomach. The significant regulatory role of ADA in maintaining adenosine concentrations has been demonstrated in ADA-deficient mice, in which a lack of ADA activity has been shown to significantly increase the endogenous adenosine level in the stomach (Xu and Kellems, 2000). Previous studies performed in our laboratory have also demonstrated that the adenosine uptake inhibitor dipyridamole can enhance basal and adenosine-induced SLI release (Kwok et al., 1990), indicating that endogenous adenosine is also regulated by the cellular reuptake of this nucleoside through equilibrative adenosine transporters. The extracellular adenosine level in the isolated vascularly perfused rat stomach may, therefore, be regulated by the cellular reuptake and metabolic degradation of adenosine. Since EHNA and dipyridamole can alter basal and adenosine-induced SLI release, endogenously released adenosine may be involved in maintaining the release of SLI and IRG under normal physiological states of the stomach. Although the origin of endogenous adenosine in the present study has not been determined, extracellular adenosine in the stomach may originate from several sources. In the perfused stomach preparation, endogenous adenosine may be derived from the metabolism of ATP released into the stomach (see Chapter 1, Section I, 1.3). ATP may be released by enteric nerves or smooth muscle cells of the gastrointestinal tract (Burnstock, 1975; Nitahara et al., 1995). In the stomach, purinergic nerves of the myenteric plexus project to the circular and longitudinal muscles (Burnstock, 1975). ATP was demonstrated to be co-released with other neurotransmitters such as nitric oxide (Selemidis et al., 1997), VIP (Furness et al., 1995) and pituitary adenylate cyclase-activating polypeptide (Furness et al., 1995; Imoto et al., 1998), and may be released by nerve depolarization (Su et al., 1971). In guinea pigs, ATP was also been shown to be released from smooth muscle cells of the ileum (Nitahara et al., 1995). Thus, ATP released from gastric neurons and smooth muscle cells may be an important source of endogenous adenosine in the rat stomach. Adenosine derived from the metabolism of ATP, thus, may depend on neural activity of the enteric plexi. The endogenous adenosine level can change under various conditions. During normoxic conditions, endogenous adenosine concentrations are in the nanomolar range. In the whole blood and plasma of rats, adenosine concentrations have been found to be 79 nM (Phillis et al., 1992) and 320 nM (Conlay et al., 1997), respectively, while in the jejunum, the venous adenosine concentration was found to be 62 nM (Sawmiller and Chou, 1990). Significant elevations in adenosine concentrations can occur during periods of stress, such as during hypoxia, ischemia, and hypoglycemia. The production of ATP is reduced when low glucose or oxygen levels produce an elevated level of AMP (Milusheva et al., 1996), which stimulates 5' nucleotidase activity to promote adenosine formation (Worku and Newby, 1983). The level of adenosine, thus, may increase to micromolar concentrations during ischemia (Latini et al., 1999). For example, ischemia has been shown to increase the adenosine concentration in the striatum from a basal level of 41- 210 nM (Ballarin et al., 1991; Pazzagli et al., 1995; Melani et al., 1999) to an ischemic level of 3 \xM (Melani et al., 1999). The physiological state of the stomach may also alter the endogenous level of adenosine. In humans, the acidity of the stomach may alter adenosine concentrations by altering ADA and 5' nucleotidase activity. In particular, in the fundic mucosa, ADA activity and gastric acid output have been observed to change in parallel (Namiot et al., 1990). Hypersecretors of acid exhibited increased ADA activity, while patients suffering from achlorhydria exhibited lowered ADA activity (Namiot et al., 1990). The activity of 5' nucleotidase was found to be reduced in the gastric juice of patients suffering from gastric cancer, gastric ulcers, and gastritis (Durak et al., 1994). The activity of ADA and 5' nucleotidase, therefore, may be regulated by the acidity of the stomach. This observation is consistent with studies, in the rat skeletal muscle, showing that increased acidity can stimulate 5' nucleotidase activity (Cheng et al., 2000). The prandial state of the stomach may also alter the adenosine level. In the canine jejunum, the venous adenosine level was shown to be increased by the presence of food in the intestinal lumen (Sawmiller and Chou, 1990). Changes to the prandial state or acidity of the stomach may, therefore, alter gastric adenosine concentrations and subsequently influence gastric actions mediated by adenosine, such as the regulation of SLI and IRG release. As discussed earlier, activation of Ai receptors has been shown to inhibit LRG and SLI release, while activation of A2A receptors has been shown to stimulate SLI release. Ai receptors are preferentially activated by lower concentrations of adenosine, while A2A receptors are preferentially activated by higher concentrations of adenosine. Therefore, during conditions that lead to an increase in the endogenous adenosine concentration, A2A receptors may be preferentially activated to stimulate SLI release, while during conditions that would lead to a decrease in the endogenous adenosine concentration, Ai receptors may be preferentially activated to inhibit SLI and IRG release. II. Distribution and structure of gastric adenosine receptor mRNA The mRNAs of all four adenosine receptors are expressed in extremely low levels within the whole rat stomach (Dixon et al., 1996). The localization of adenosine receptors on functionally and morphologically distinct regions of the stomach is unknown. However, functional studies suggest that these receptors may be localized on specific regions of the stomach. For example, studies demonstrating the protective effect of adenosine on gastric mucosal damage (Geiger and Glavin, 1985; Westerberg and Geiger, 1987; Cho and Ogle, 1990; Cho et al., 1991; Bozkurt et al., 1998) and studies showing the inhibitory and stimulatory effect of adenosine on gastrin and somatostatin release, respectively (Kwok et al., 1990), suggest that the adenosine receptors may be expressed on the gastric mucosa. The current study is the first to demonstrate the localization of all four adenosine receptors in the gastric corpus, antrum, and mucosa of the rat using RT-PCR. The specificity of each RT-PCR reaction was confirmed by restriction enzyme digestion using multiple restriction enzymes. Small fragments generated by restriction enzyme digestion (< 150 bp) were often not detected by agarose gel electrophoresis due to their low intensity fluorescence. The larger fragments (>150 bp) were easily detected, and their sizes compared well with the expected sizes, differing by < 9%. The Ai, A2A, A2B and A3 receptor RT-PCR reactions established for this study, therefore, are highly specific. The presence of adenosine receptors in the corpus, antrum and mucosa may have important functional implications since these regions contain cells which secrete gastric acid, somatostatin and/or gastrin. The parietal cells, G-cells and D-cells reside in the gastric mucosa of the stomach (Dockray et al., 1996; Sachs et al., 1997). The corpus contains parietal cells and D-cells, while the antrum contains G-cells and a population of D-cells that are distinct from those in the corpus. In addition to their presence in these tissues, all four adenosine receptors were also expressed in the gastric fundus, the non-glandular region of the stomach that is responsible for the accommodation of food. The presence of Ai and A2A receptor mRNA throughout the stomach suggests the possibility that adenosine receptors may be expressed on various sites of the stomach, including the parietal cells, G-cells and D-cells, nerve fibers and vasculature. Indeed, the localization and distribution of adenosine receptors were confirmed on some of these structures by the present immunohistochemistry experiments. The structure of the Ai and A2A receptors was also examined in the present study by RT-PCR and by cloning and sequencing of the RT-PCR amplicons. Only one Ai and A2A receptor amplicon is produced when RT-PCR was performed using primers that span the entire coding region of the receptor. Northern blot analysis has previously demonstrated the existence of two Ai receptor transcripts in various tissues of the rat, including the brain and stomach (Mahan et al., 1991; Reppert et al., 1991). Two distinct A2A receptor transcripts have also been demonstrated in rat PC12 cells (Chu et al., 1996). The Ai and A2A receptor genes both contain 2 exons, which are separated by one intron (Ralevic and Burnstock, 1998). The synthesis of only one RT-PCR amplicon in the present study demonstrates that the intron is not alternatively spliced to generate variant forms of the Ai and A2A receptor in the rat stomach. Cloning and sequencing of the mucosal Ai and A2A receptor amplicons further demonstrate that the coding region of these receptors is identical to published sequences in the rat brain (Mahan et al., 1991; Fink et al., 1992). Thus, only one form of the gastric Ai and A2A receptors exists in the stomach. The multiple Ai and A2A receptor mRNA transcripts, detected by previous Northern blot analysis studies (Mahan et al., 1991; Reppert et al, 1991; Chu et al., 1996) may arise from differences in the non-coding regions. In rat PC 12 cells, the A2A receptor gene was shown to express two different promoters (Chu et al., 1996). These two promoters drive the synthesis of two distinct A2A receptor transcripts that differ in the 5'UTR. Similarly, in humans, two separate promoters were found in the Ai receptor gene (Ren and Stiles, 1995). These promoters drive the synthesis of two distinct Ai receptor transcripts which were also shown to differ in the 5'UTR. It is unclear whether this also occurs in the rat stomach since the promoter regions of the rat Ai receptor gene have not yet been identified. Unlike the Ai and A2A receptors, two forms of the A3 receptor exist in various rat tissues (Zhou et al., 1992; Sajjadi et al., 1996). The predominant form was cloned by Zhou and coworkers (1992), and was shown to contain 320 amino acids. The alternate form, the A3i receptor, was cloned by Sajjadi and coworkers (1996) and was found to be expressed in the rat brain and spleen (Zhou et al., 1992; Sajjadi et al., 1996). This form contains a 17 amino acid insertion which is produced by alternative splicing of the intron. The RT-PCR primers used in the present study were able to detect both the A3 and A3i receptor transcripts. Results of our study demonstrate that alternative splicing was not shown to occur in any region of the rat stomach since only one A3 receptor transcript was detected. Cloning and sequencing of the PCR amplicon revealed that the A3 receptor, but not the A3i receptor, is expressed in the rat stomach. A silent mutation was also shown to occur at position 492 (second transmembrane domain). This mutation has previously been observed in the A3i receptor transcript of the rat brain (Sajjadi et al., 1996). Other point mutations reported in the A3i receptor, however, were not observed in this region of the rat mucosal A3 receptor gene. Results of these studies demonstrate that all four adenosine receptors are expressed in functionally distinct regions of the rat stomach, and that each subtype exists in only one structural form. Furthermore, the gastric Ai and A2A receptors are structurally identical to the Ai and A2A receptors of the brain in the rat. III. Quantification of Ai and A2A receptor gene expression 1. Competitive RT-PCR and Real-Time RT-PCR assay development In the rat stomach, the level of adenosine Ai and A2A receptor mRNA is extremely low (Dixon et al., 1996). Quantification of gastric adenosine receptor expression using conventional techniques such as Northern blot and slot blot is not possible since moderate levels of mRNA expression and relatively large sample sizes are required. In the present study, competitive RT-PCR and Real-Time RT-PCR techniques were developed to measure absolute Ai and A2A receptor gene expression in different regions of the rat stomach. Both methods are widely accepted techniques for quantifying low levels of gene expression in small tissue samples, such as the gastric mucosa (Raeymaekers, 1995; Heid et al., 1996). Quantitative RT-PCR has been used to measure Ai and A2A receptor gene expression in the rat brain (Rose'Meyer et al., 2003). The relative changes in Ai receptor mRNA expression have been determined in the rat brain using competitive RT-PCR (Othman et al., 2002) and in the rat heart using Real-Time PCR (Ashton et al., 2003). The present study is the first to quantify the absolute level of Ai and A2A receptor gene expression in the rat stomach, where the expression of these receptors are extremely low (Dixon et al., 1996). Both competitive RT-PCR and Real-Time RT-PCR assays were developed to measure this gene expression. Competitive RT-PCR is based on the competitive co-amplification of a fixed quantity of target template with known amounts of internal standard or RNA competitor. The Ai and A2A receptor competitive RT-PCR assays developed in this study were able to measure gene expression ranging from 105 tolO7 copies/pg total RNA. Each reaction tube contained a fixed amount of tissue RNA; 0.125 pg and 0.5 pg were used in the A, and A2A receptor assays, respectively. The amount of template used was optimized so that the tissue template and competitor band were both easily visualized, and so that both the tissue template and competitor cDNA amplified with equal efficiency during PCR. Using the competitive RT-PCR assay, striatal Ai and A2A receptor mRNA levels were found to be at least 4.7 times and 116 times greater than in any region of the stomach, respectively. This is consistent with studies demonstrating moderate Ai and abundant A2A receptor gene expression in the striatum and extremely low Aj and A2A receptor gene expression in the stomach (Dixon et al., 1996). Ai receptor gene expression was shown to be significantly higher than A2A receptor gene expression in the antrum and mucosa, but not in the fundus and the corpus. These results do not agree well with results from the Real-Time RT-PCR experiments. Real-Time RT-PCR results demonstrate that Ai receptor gene expression is significantly higher than A2A receptor gene expression in all gastric regions examined. The discrepancy between the competitive RT-PCR and Real-Time RT-PCR results is addressed later in this section. Real-Time RT-PCR is based on the cleavage of a fluorogenic probe that hybridizes to the template at a position located between the forward and reverse primers (See Fig. 9). The Real-Time RT-PCR assays used to measure Ai and A2A receptor gene expression were shown to be considerably more sensitive than the corresponding competitive RT-PCR assay. The Real-Time RT-PCR assays were able to measure a range of concentrations exceeding a 7 log scale, and as few as 500 copies of Ai or A2A receptor RNA/u.g total tissue RNA. The latter is equivalent to 25 copies of Ai or A2A receptor transcript per reaction since each reaction mixture only contained 0.05 [ig of total RNA as the template. Using Real-Time RT-PCR, Ai receptor gene expression was lowest in the corporeal mucosa, while A2A receptor gene expression was lowest in the whole stomach and corporeal mucosa. These results correspond well with the results of the immunohistochemistry studies. AiR-IR and A2AR-IR were found on some mucosal cells, but the majority of AiR-IR and A2AR-IR was localized on the nerve fibers and vasculature of the muscle and myenteric plexus of the corpus and antrum. In the antral mucosa, AiR-IR was localized on all G-cells and some D-cells, while A2AR-IR was only localized on some D-cells. Thus, the higher level of Ai compared to A2A receptor mRNA in the antrum and whole stomach mucosa may reflect the higher number of G-cells compared to D-cells in these regions (Lehy et al., 1979). Striatal Ai and A2A receptor mRNA levels measured by Real-Time RT-PCR were shown to be at least 2- and 70-fold higher, respectively, than levels measured in any region of the stomach. The mRNA levels of the Ai and A2A receptor measured using this technique were consistently lower than those measured using competitive RT-PCR; Ai and A2A receptor mRNA levels were found to be 2.4±0.2 and 21±3 times lower, respectively. The absolute level of A\ receptor gene expression has not previously been quantified in the rat stomach or striatum. Only the relative level of Ai receptor gene expression in various tissues have been determined using RT-PCR (Dixon et al., 1996). The striatal A2A receptor mRNA levels measured by Real-Time RT-PCR (2.2 x 106 copies/pg total RNA), however, compared well with levels measured in a previous study (Lee et al., 1999). The striatal A2A receptor gene expression measured by Lee and co-workers (1999) was approximately 1.4 x 106 copies/pg total RNA, using a similar experimental protocol. The sequence of the primers and fluorogenic probe, and the total RNA extraction protocol used, however, were different, and may account for the small difference between their striatal A2A mRNA level and that found in the present study. The striatal A2A receptor level determined by competitive RT-PCR (6.6x 107 copies/pg total RNA) was not in the same range as that measured by Real-Time RT-PCR. Real-Time RT-PCR is likely the more accurate and reliable technique. The added component of the fluorogenic probe significantly increases the specificity of the Real-Time RT-PCR assay. This technique is more sensitive since the fluorogenic signal emitted by the reporter dye is detected by a highly sensitive CCD camera. Studies have shown that Real-Time RT-PCR can reliably detect a single specific DNA molecule in a high background of non-specific DNA (Lockey et al., 1998). Our assays were able to consistently measure as few as 25 copies of Ai and A2A receptor RNA per reaction. Changes in reporter fluorescence are also monitored in a Real-Time manner. Thus, quantification is made at the start of the exponential phase and the errors that may be introduced by reaching the plateau phase are avoided (Raeymaekers, 1995; Schmittgen, 2001). The difference between the Ai and A2A receptor mRNA levels measured by competitive RT-PCR and Real-Time RT-PCR is unclear. Several factors can introduce errors to competitive RT-PCR measurements, but in the present study, care was taken to avoid these. For example, during competitive RT-PCR, the number of cycles of amplification was limited to 30 to avoid reaching the plateau phase. As the reaction approaches the plateau phase, there is an increased likelihood of heteroduplex formation. These complexes are formed when the competitor and tissue template PCR products anneal. These formations can significantly decrease the sensitivity and accuracy of competitive RT-PCR quantification (Henley et al., 1996). These complexes would appear as an additional band above the competitor and tissue template bands. However, in the present study, heteroduplex formations were not observed for either the Ai or A2A receptor competitive RT-PCR assays. Accurate quantification with competitive RT-PCR also requires that the template and the competitor behave in a similar manner during the reverse transcription and PCR step (Schmittgen, 2001). In this study, RNA competitors were used instead of DNA competitors to control for variations in the reverse transcription step (Orlando et al., 1998). In addition, homologous competitors, which have a sequence similar to that of the tissue template, were also used to minimize differences in the amplification efficiency during PCR. An equal amplification efficiency of the template and competitor is required for reproducible and accurate quantification (Connolly et al., 1995). The amplification efficiencies were determined in this study by plotting the log [density ratio (competitor/template) band] against the log (initial competitor concentration) for each assay. When the slope of the competitive RT-PCR relationship is equal to 1, both the template and competitor cDNA should amplify with equal efficiency (Connolly et al, 1995). The slopes obtained for the Ai and A2A receptor competitive RT-PCR assays were found to be 1.05±0.04 and 1.07±0.05, respectively. Thus, a significant difference in amplification efficiency between the tissue template and competitor cDNA was not apparent. Some studies have shown that, even when these slopes are ideal (= 1.0), competitive RT-PCR can yield results that differ significantly from actual levels. For example, Hayward et al. have shown that when a known quantity of Na+K+-ATPase mRNA was measured by competitive RT-PCR, the value reported was almost 4-fold higher although the slope of the competitive RT-PCR relationship remained at 1.01 (Hayward et al., 1998). These investigators further demonstrated, using mathematical modeling, that small differences in the amplification efficiency between the competitor and unknown template can drastically alter competitive RT-PCR measurements, while still producing a relationship with a slope of 1.0. A 20% difference in amplification efficiency between the competitor and unknown template was found to result in a 9 to 20-fold difference between the actual mRNA level and the value estimated by competitive RT-PCR (Hayward et al., 1998). In our studies, Ai and A2A receptor mRNA levels measured by competitive RT-PCR were found to be 2.4 and 21-fold higher, respectively, than levels measured by Real-Time RT-PCR. Thus, a difference in the amplification efficiency between the competitor and unknown template may exist in our assays, despite the precautions taken. The Ai and A2A competitors contain a 136 and 57 bp deletion. Decreasing the size of the deletion has been shown to increase the accuracy of competitive RT-PCR (Hayward et al., 1998). However, the resulting similarity between the size of the competitor and unknown template make it difficult to distinguish between the two bands during gel electrophoresis. In view of the limitations of competitive RT-PCR, Real-Time RT-PCR was used in our subsequent experiments to measure changes in Ai and A2A receptor gene expression after various treatments, including food deprivation and omeprazole treatment. This technique is highly specific and is not subject to the limitations of competitive RT-PCR. In addition, this technique was chosen over competitive RT-PCR since striatal A2A receptor mRNA levels measured by Real-Time RT-PCR compared well with previously published results (Lee et al, 1999). The extremely high sensitivity of this method also allows for quantification of Ai and A2A receptor gene expression in small gastric tissue samples such as the mucosa, where the expression levels of these genes are extremely low. 2. Changes of gastric Ai and A2A receptor gene expression and function by altered states of the stomach The present study shows that the Ai and A2A receptors participate in the regulation of IRG and SLI release in the rat stomach. However, it is unclear if the expression of these receptors is altered by different secretory states of the stomach. Changes to adenosine receptor gene expression may subsequently alter the modulatory effect of adenosine on IRG and SLI release. Previous studies have shown that oxidative stress (Nie et al., 1998; Kobayashi and Millhorn, 1999) or stress induced by acute renal failure (Gould et al., 1997) and sleep deprivation (Basheer et al., 2000) can alter adenosine receptor gene and/or protein expression. The present study demonstrates that stress induced by fasting and omeprazole treatment may also alter adenosine receptor gene expression. Real-Time RT-PCR was performed to measure changes in A] and A2A receptor, gastrin and somatostatin gene expression. All measurements were standardized with an endogenous control to account for differences resulting from variations in total RNA extraction efficiency, RNA integrity and RNA concentration. Two common house-keeping genes, GAPDH and 18S rRNA, were considered for use as the endogenous control. These genes can only be used as the endogenous control if their expression is not altered by the experimental treatment. In the current study, GAPDH gene expression was significantly decreased in the corpus mucosa after 24 and 36 h of fasting, and in the antrum after 36 h of fasting. Similar results have been reported in animals which were subjected to 48 h of fasting (Yamada et al, 1997a). GAPDH mRNA levels were also significantly altered in the corporeal muscle and mucosa after 1 and 3 days of omeprazole treatment, respectively. These findings are consistent with another study, which demonstrated that a single dose of omeprazole was capable of increasing GAPDH mRNA levels in the corpus (Sandvik et al., 1995). Furthermore, several studies have cautioned against the use of GAPDH mRNA as an endogenous control since its expression can be altered by a variety of conditions and agents such as hypoxia, oxidative stress, heat shock, glucose and insulin (Schmittgen and Zakrajsek, 2000; Suzuki et al., 2000). In the current study, 18S rRNA levels were not affected by fasting or omeprazole treatment. Thus, this gene was used as the endogenous control for the measurement of all gene expression. Previous experiments have also demonstrated that 18S rRNA expression was not altered by 48 h of fasting (Yamada et al., 1997a). Using 18S rRNA expression as the endogenous control, both fasting and omeprazole treatment were shown to alter adenosine A, and A2A receptor gene expression in the corporeal mucosa and antrum, but not in the corporeal muscle layer. In the fasting experiments, an increase in Ai receptor gene expression was observed in both the corporeal mucosa and antrum after 24 h, but not after 36 h, of fasting. Similarly, fasting was shown to increase A2A receptor gene expression in the antrum after 24, but not after 36 h, of fasting. Although adenosine receptor protein expression may change in parallel with its mRNA level, the return of adenosine receptor gene expression to control levels after 36 h of fasting does not necessarily imply that adenosine receptor protein expression also returns to control levels. It is possible that the elevated adenosine receptor mRNA levels observed after 24 h of fasting resulted in increased adenosine receptor expression, which is maintained even after 36 h of fasting. The effect of fasting on A2A receptor gene expression differed in the antrum and corporeal mucosa. In the antrum, A2A receptor gene expression was shown to increase after 24 h of fasting. In the corporeal mucosa, its gene expression was shown to gradually decrease and become significant after 36 h of fasting. The different effect of fasting on A2A receptor gene expression in the antrum and corporeal mucosa suggests that the gene expression of this receptor may be differentially regulated in these two gastric regions. The lack of change in Ai and A2A receptor gene expression in the corporeal muscle suggests that neither gene is altered by fasting in this tissue. It is currently unclear how adenosine receptor gene expression is altered by fasting. The absence of food in the stomach has been shown to increase gastric acidity (Matsumoto et al., 1989; Jiang et al., 2002). In the rat, Jiang and colleagues (2002) demonstrated that fasting for 24 h significantly decreased the gastric intraluminal pH to 2.0. Similarly, Matsumoto and co-workers (1989) demonstrated that fasting significantly decreased gastric intraluminal pH from a non-fasting level of 3.5 to 2.5, 2.0, and 1.6 after 1, 2, and 3 days of fasting, respectively, and that the intraluminal pH did not return to control levels until after 8 days of fasting. These studies suggest that changes in gastric acidity during fasting may alter gastric adenosine receptor gene expression. In agreement with this proposal, omeprazole-induced achlorhydria was also shown to alter Ai and A2A receptor gene expression. Omeprazole (400 fj,mol/kg) was found to completely inhibit gastric acid secretion within 2 h of administration (Lee et al., 1992). Gastric acid secretion was decreased by up to 80% 24 h after the first treatment (Larsson et al., 1988; Lee et al., 1992), and continuous inhibition of acid secretion was achieved after 3 days of treatment (Carlsson et al., 1986). Results show that omeprazole treatment inhibited both Ai and A2A receptor gene expression in the antrum, which is contrary to what occurs during fasting. Since fasting increases gastric acidity and omeprazole decreases gastric acidity, intraluminal acidity may play a role in altering adenosine receptor gene expression. In support of this proposal, the gene expression of gastric Ai receptors was observed to be increased after fasting but decreased after omeprazole treatment. Results of our immunohistochemistry experiments demonstrate that Ai receptors were expressed on D-cells and G-cells and A2A receptors were expressed on D-cells of the gastric mucosa. In the antrum, both of these cell types are open to the luminal environment and, thus, can respond to changes in gastric acidity, while the D-cells of the corpus are closed and do not readily respond to changes in gastric acidity (Sachs et al., 1997). Therefore, it is possible that gene expression of adenosine receptors in the antrum is readily altered by changes in gastric acidity, while adenosine receptor gene expression in D-cells of the corpus mucosa may not be directly affected by gastric acidity. This suggestion may explain why A2A receptor gene expression in the corporeal mucosa did not alter according to the changes in gastric acidity. In particular, A2A receptor gene expression was decreased by both fasting and omeprazole treatment. The possible mechanisms involved in the regulation of adenosine receptor gene expression by gastric acidity have not been examined in the present study. However, there is evidence to suggest that alterations in the activities of various mitogen-activated protein kinases (MAPK) may be involved. Malek and colleagues (1999) have shown that activation of ERK1, ERK2 and INK can decrease adenosine receptor A2A mRNA levels in rat PC 12 cells. The activities of various MAPK respond to changes in both intracellular and extracellular pH (Feifel et al., 2002). Exposure of human epidermoid carcinoma cell line A431 to low extracellular pH was shown to increase the activity of MAPK (ERK2, JNK, and p38 MAP kinase), and exposure of mouse Swiss 3T3 cells to low extracellular pH was shown to stimulate the activity of MAPK (p42, p44 MAP kinase, ERKs, p38 MAP kinase and JNK) (Xue and Lucocq, 1997). In the Barrett's esophagus adenocarcinoma cell line (SEG-1), increased exposure to acidic media increased ERK, p38 MAP kinase, and JNK activities (Souza et al., 2002). The activity of p38 MAP kinase was also shown to be increased in esophageal mucosa cells of patients who suffer from Barrett's esophagus (Souza et al., 2002). Barrett's esophagus is associated with gastroesophageal reflux disease. Thus, the increased p38 MAP kinase activity in these cells is likely due to the increased exposure to acid. The studies mentioned above have demonstrated that changes in acidity may alter MAPK activity. Increases in MAPK activity may subsequently alter adenosine receptor gene expression. The effect of specific MAPK on Ai receptor gene expression has not been determined. However, in the rat pituitary-derived cell line, Ai receptor transcription is stimulated by epidermal growth factor (EGF) (Navarro et al., 1999). EGF has also been shown to activate MAPK pathways (Yamada et al., 1997b). Although MAPK activity may alter adenosine receptor gene expression in some cell lines (Malek et al., 1999), this has not been examined in the stomach. Adenosine receptor gene expression may also be regulated by changes in intracellular pH. During periods of oxidative stress, such as hypoxia and ischemia, intracellular pH is decreased (Kintner et al., 2000). Previous studies have shown that oxidative stress can induce Ai receptor gene expression in DDT1-MF2 cell line (Nie et al., 1998), and that hypoxia can stimulate A2A receptor gene expression (Kobayashi and Millhorn, 1999). During fasting, it is possible that increased intraluminal acidity leads to subsequent increases in intracellular acidity in the gastric epithelium. In the guinea pig gastric gland, decreasing the luminal pH from 3.0 to 1.9 was shown to decrease intracellular pH in cells of the gastric gland (Schreiber et al., 2000). Similar results have also been found in the isolated Necturus antral mucosa, where acidification of the luminal pH to 2.0 was shown to significantly decrease intracellular pH of the antral mucosal cells (Kivilaakso and Kiviluoto, 1988; Kiviluoto et al., 1990). Since the intraluminal pH level has been shown to decrease from 3.5 to 2.0 after 24 h and 48 h of fasting (Matsumoto et al, 1989; Jiang et al., 2002), it is possible that the increase in Ai and A2A receptor mRNA levels observed in the antrum after 24 h of fasting result from intracellular acidification. Although fasting and omeprazole treatment may regulate adenosine Ai and A2A receptor gene expression by altering the intraluminal acidity, it is unclear whether adenosine receptor gene expression may also respond to other factors. For example, during fasting, adenosine receptor gene expression may be altered by the absence of specific nutritional components of food in the stomach. The presence of peptones, phenylalanine and olive oil, in the gastric lumen has been shown to regulate the gene expression of gastrin and somatostatin (Wu et al., 1990b; Wu et al., 1991). Omeprazole treatment has also been shown to increase plasma levels of GRP (Takehara et al., 1996), which has been shown to regulate the activity of D-cells and G-cells. D-cells of the antrum and corpus can be regulated directly (Schaffer et al., 1997) and indirectly (Schubert et al., 1991) by GRP, respectively, while G-cells can be directly modulated by GRP (Campos et al., 1990). The G-cells and D-cells were shown to express adenosine receptors. Thus, fasting and omeprazole-treatment may also induce changes in adenosine receptor gene expression by preventing the exposure of the gastric lumen to nutritional components of food and by enhanced GRP release, respectively. Since activation of Ai and A2A receptors inhibited gastrin and stimulated somatostatin release, respectively, changes in A\ and A2A receptor expression may also regulate gastrin and somatostatin synthesis and release. Therefore, the effect of fasting and omeprazole treatment on gastrin and somatostatin gene expression was also examined. Results show that gastrin gene expression was significantly inhibited by 24 and 36 h of fasting. These findings are consistent with previous studies demonstrating a significant reduction in gastrin gene expression after 12, 24, 48 and 72 h of fasting (Dimaline et al., 1991; Wuet al., 1991; Larsson and Hougaard, 1993; Sandvik et al., 1993; Yamada et al., 1997a). These results are also consistent with the decrease in antral gastrin concentration, G-cell number and serum gastrin content observed after fasting (Lichtenberger et al., 1975; Track et al., 1978; Shulkes et al., 1983; Dimaline et al., 1991; Larsson and Hougaard, 1993). Omeprazole treatment, on the other hand, was shown to increase gastrin mRNA levels after 1 and 3 days of treatment. Increased gastrin gene expression has also been demonstrated after 4 and 14 days of omeprazole treatment (Wu et al., 1990a; Dockray et al., 1991). The increased gastrin mRNA levels are most likely responsible for the enhanced levels of antral gastrin content and plasma gastrin concentrations (Brand and Stone, 1988; Inauen et al., 1988; Larsson et al., 1988; Dockray et al., 1991), which have been observed following short term (5 days or less) omeprazole treatment. Both fasting and omeprazole treatment were shown to induce reciprocal changes in Ai receptor and gastrin gene expression. Our immunohistochemistry results demonstrate that Ai receptors are expressed on G-cells, which suggests that increased expression and activation of Ai receptors may directly affect gastrin gene expression. Adenosine has been shown to modulate the expression of various genes in the murine heart (Asakura et al., 2002), and may modulate gene expression by activating Ai receptors to inhibit cAMP production. The transcription of certain genes is cAMP-dependent (Dragon et al., 2002). In particular, gastrin gene expression was regulated by cAMP levels (Shiotani and Merchant, 1995). Since activation of Ai receptors inhibits adenylate cyclase activity and cAMP formation (Ralevic and Burnstock, 1998), it is possible that activation of Ai receptor on G-cells may inhibit gastrin gene expression by inhibiting cAMP accumulation. In addition, somatostatin has been shown to inhibit gastrin synthesis by suppressing adenylate cyclase activity and subsequent cAMP production (DelValle and Yamada, 1990). Somatostatin gene expression was also altered by fasting and omeprazole treatment. The antral somatostatin mRNA level was shown to be increased following 24 h of fasting, but returned to control levels after 36 h of fasting. Previous experiments have demonstrated a significant increase in antral somatostatin gene expression following both 24 h (Wu et al., 1991) and 36 h of starvation (Sandvik et al., 1995). The cause of this discrepancy is unclear. However, it may be due to the use of a different housekeeping gene as the endogenous control. Sandvik and co-workers utilized GAPDH mRNA as the endogenous control. Sandvik reported no significant change in GAPDH mRNA expression after 36 h of fasting (Sandvik et al., 1995), contrary to this study and another report (Yamada et al., 1997a). In the corporeal mucosa, the somatostatin mRNA level was reduced following 36 h of fasting. This decrease is consistent with previous studies demonstrating a significant decrease in the somatostatin mRNA level in the oxyntic region following a 48 h fasting period (Sandvik et al., 1993). Results of the present experiments show that antral somatostatin gene expression was significantly decreased after 1 and 3 days of omeprazole treatment (400pmol/kg). These findings are similar to those reported in previous studies (Brand and Stone, 1988; Wu et al., 1990a; Wu et al., 1990b; Sandvik et al., 1995). The lowered antral somatostatin mRNA levels are consistent with the decreased plasma concentration of somatostatin, antral somatostatin concentration, and antral D-cell density that have been shown to occur in the rat stomach during achlorhydria (Allen et al., 1986; Lee et al., 1992; Pawlikowski et al., 1992). In the corpus, the present study shows that somatostatin mRNA levels were significantly decreased after omeprazole treatment. These observations are in agreement with several studies demonstrating reduced corporeal somatostatin mRNA expression during achlorhydria (Tari et al., 1991; Sandvik et al., 1995). Similar to our results, Tari and co-workers reported that the somatostatin mRNA level in the corpus gradually decreased over a period of 3 days during drug treatment (Tari et al., 1991). Other studies have shown that omeprazole can also cause an increase, rather than a decrease, in corporeal somatostatin gene expression (Kapuscinski and Shulkes, 1995; Bolkent et al., 2001). However, these observations were made 8 h after a single treatment of omeprazole (400 pmol/kg) (Kapuscinski and Shulkes, 1995) or after long-term (2 months) omeprazole treatment (Bolkent et al., 2001). The present study demonstrates that somatostatin gene expression was inhibited by achlorhydria in the corporeal muscle. Our immunohistochemistry experiments demonstrate that, in this region, somatostatin was expressed on very few nerve fibers of the circular muscle and myenteric plexus. Achlorhydria has previously been shown to alter neuropeptide mRNA expression in the enteric plexi (Dimaline et al., 1992). In particular, Dimaline and colleagues have shown that omeprazole treatment can increase the antral GRP mRNA level. The increase in somatostatin gene expression in this tissue may be secondary to changes in GRP levels, since GRP has been shown to stimulate somatostatin release (Martindale et al., 1982). Both fasting and omeprazole treatment induced similar changes in A2A receptor and somatostatin gene expression. Results of perfusion experiments performed in the present study demonstrate that activation of A2A receptors stimulated somatostatin release. Furthermore, results of the immunohistochemistry study demonstrate the expression of A2A receptors on D-cells. Thus, it is possible that changes in A2A receptor expression regulate somatostatin synthesis and release. Previous studies have shown that the rat somatostatin gene contains a cAMP response element (Montminy et al., 1986) and that its expression is regulated by cAMP levels (Montminy et al., 1996b). The A2A receptor couples to the Gs protein and can stimulate adenylate cyclase activity to increase cAMP accumulation (Dalziel and Westfall, 1994; Fredholm et al., 2001). A number of studies have demonstrated that activation of A2A receptors induces the expression of genes regulated by the cAMP response element (CRE). Asher et al. have demonstrated that activation of A2 receptors can induce cAMP-dependent gene expression through the phosphorylation of CRE binding protein (CREB) (Asher et al., 2002). In vascular smooth muscle cells, activation of the A2 receptor was suggested to upregulate lysyl oxidase gene expression (Ravid et al., 1999). In addition, CGS 21680 (A2A-selective agonist) has been shown to induce tyrosine hydroxylase gene expression in rat PC12 cells (Chae and Kim, 1997). Like the somatostatin gene, both the lysyl oxidase and tyrosine hydroxylase genes also contain a CRE in the 5' upstream promoter region. Therefore, it is possible that activation of adenosine A2A receptors also increases somatostatin gene expression through activation of the CRE. Although the direct modulation of the gastrin and somatostatin gene expression by activation of the Ai and A2A receptors remains a possibility, specific experiments have not been performed in the present study to confirm this proposal. The regulation of gastrin and somatostatin mRNA expression is complex and may also depend on a number of other factors. To examine whether changes in adenosine receptor mRNA levels may result in actual changes in adenosine receptor function, the effect of omeprazole treatment on adenosine agonist-induced changes in IRG and SLI release was also examined. The level of gene expression often changes in parallel with protein synthesis and receptor function. However, the results of the present study suggest that changes in gastric adenosine receptor gene expression do not always result in changes in receptor function. CPA, an Ai receptor agonist, was demonstrated to be a potent inhibitor of IRG release in the isolated vascularly perfused rat stomach. When administered at low (<0.1 uM) and high (>1 uM) concentrations, CPA suppressed and stimulated SLI release, respectively. A similar concentration-dependent effect has been demonstrated using adenosine (Kwok et al., 1990). After treatment with omeprazole, Ai receptor gene expression was reduced in the corporeal mucosa and antrum. Following 1 or 3 days of omeprazole treatment, CPA-induced inhibition of IRG and SLI release was not altered. However, the stimulatory effect on SLI release induced by the higher concentration (1 uM) of CPA was significantly decreased following 3 day treatment with omeprazole. As discussed earlier, the inhibition of IRG and SLI release induced by CPA is likely mediated by activation of Ai receptors, while the augmentation of SLI release induced by high concentrations of CPA is likely mediated by the non-selective activation of A2A receptors. The attenuation of CPA-stimulated SLI release after omeprazole treatment, therefore suggests that A2A receptor expression is reduced after 3 days of omeprazole treatment. In agreement with this proposal, the stimulatory action of the A2A receptor agonist CGS 21680 on SLI release was also reduced after 3 days of omeprazole treatment. Both 0.1 and 1 pM CGS 21680 potently stimulated the release of SLI in earlier experiments. Although the stimulatory effect of 0.1 pM CGS 21680 on SLI release was suppressed after 3 days of treatment, this effect was not statistically significant. The release of SLI induced by 1 pM CGS 21680, however, was significantly attenuated after 3 day omeprazole treatment. This attenuation of A2A receptor function is consistent with the reduced level of antral A2A receptor gene expression observed after 3 days of omeprazole treatment. Taken together, results from the gene expression and perfusion experiments suggest that omeprazole treatment decreases the number of functional A2A receptors, but does not always produce parallel changes in adenosine receptor expression. In the present study, A2A receptor gene expression was also reduced in the corporeal mucosa after 1 day of omeprazole treatment, but significant changes in CGS 21680-induced SLI release were not observed at this time. Thus, alterations in A2A receptor mRNA levels may not necessarily produce changes in A2A receptor function. It is possible that, although A2A receptor mRNA levels may be altered after 24 h of achlorhydria, actual changes in A2A receptor expression may require more than 24 h to occur. The protein synthesis and expression of the adenosine receptors may also be controlled by other factors. The synthesis of both the A\ and A2A receptors can be regulated at the translational level (Ren and Stiles, 1994b; Chu et al., 1996; Lee et al., 1999). The translation of the human Ai receptor was shown to be inhibited by two upstream AUG codons in exon 4 (Ren and Stiles, 1994b), while translation of the human A2A receptor is inhibited by the presence of several upstream open-reading frames and regions of high GC content (Chu et al., 1996). The expression of the rat A2A receptor is suppressed by the presence of an upstream out-of-frame AUG codon in the 5' untranslated region (Lee et al., 1999). Thus, in addition to the regulation of A\ and A2A receptor expression by its mRNA levels, receptor expression can also be controlled by translational mechanisms. It is uncertain whether fasting and omeprazole treatment also alter the translation of the Ai and A2A receptor genes. Since Ai and A2A receptor gene expression was examined in whole tissue extracts, it is difficult to assess whether changes in A( and A2A receptor mRNA expression only occur in anatomical structures that are involved in the direct control of IRG and SLI release, such as the G-cells and D-cells. Our immunohistochemistry experiments have demonstrated that, in the stomach, the adenosine Ai and A2A receptors are not expressed solely on these cells. The significant functional implications of the widespread localization of Ai and A2A receptors are discussed in the following section. IV. Cellular localization of the adenosine Ai and A2A receptor From the discussion above, it is clear that both adenosine Ai and A2A receptors play a role in regulating gastric secretions. However, studies examining the cellular localization of gastric adenosine receptors are lacking. Autoradiography studies have not been performed in this tissue and in situ hybridization experiments are not sensitive enough to detect the low levels of adenosine receptor mRNA in the stomach (Dixon et al., 1996). The present study uses the technique of immunohistochemistry to determine the cellular localization of adenosine Ai and A2A receptors in the rat stomach. This study is the first to provide the anatomical basis for various gastric actions of adenosine. In the present study, AiR-IR and A2AR-ER- were found to be widely distributed throughout the stomach. Both were present on cells of the mucosa, cell bodies of the myenteric plexus, nerve fibers of the muscle layers and enteric plexi, and the gastric vasculature. The regional localization and relative density of AjR-IR and A2AR-IR in the gastric tissues agree well with results of the Real-Time RT-PCR experiments. Overall, AiR-IR appeared to be more abundant than A2AR-IR. Although some mucosal cells were shown to contain intense adenosine receptor staining, the majority of AiR-IR and A2AR-IR was localized in the enteric plexi. The localization of the gastric Ai and A2A receptors in various regions of the stomach also agrees well with their distribution in other tissues of the gastrointestinal tract. Although the anatomical distribution of adenosine receptors in the rat intestines has not been determined, immunohistochemical localization of the Ai and A2A receptors has been demonstrated in the human intestines (Christofi et al., 2001). Some anatomical structures, which were labeled in the present study, were also labeled in the human intestines. AiR-IR was demonstrated in both the myenteric and submucosal plexi of the human jejunum and colon, respectively. Neurons of the circular muscle of these tissues contained AiR-IR. Using the same A2A receptor antibody as the present study, Christofi and co-workers (2001) also demonstrated the expression of A2AR-IR in myenteric and submucosal neurons of the human jejunum and colon. In the jejunum, A2AR-IR was found to be localized in epithelial cells, while, in the colon, staining of this receptor was observed in myenteric neurons. A2AR-IR was also detected on some nerve fibers in the intestines, in particular, in the longitudinal muscle of the jejunum and circular muscle of the colon. The neuronal localization of these receptors in the stomach and intestine agrees with the role of adenosine in the regulation of motility. The cellular localization of these receptors in the mucosa and vasculature provide the anatomical basis for the known effect of adenosine on somatostatin and gastrin release and vascular actions in the stomach, respectively. 1. Adenosine Ai and A2A receptors of the gastric mucosa Results show that cells containing distinct AiR-IR and A2AR-IR were distributed throughout the corporeal and antral mucosa. These findings support the role of adenosine in regulating various gastric secretions. Tissue sections were double stained to determine whether these receptors are localized on specific cells of the mucosa, in particular, the acid-secreting parietal cells, gastrin-secreting G-cells, and somatostatin-secreting D-cells. Although ECL cells are known to be present in the mucosa, the localization of adenosine receptors on these cells was not examined. ECL cells are difficult to identify by immunohistochemistry since the preservation of histamine and histidine decarboxylase, ECL cell markers, require special fixation techniques (Bordi et al., 2000). ECL cells are commonly identified by chromogranin A immunostaining. However, this product is not exclusive to the ECL cells, and is also found in other endocrine cells such as G-cells. Results of our studies suggest that the AjR and A2AR are unlikely to be expressed on ECL cells of the rat. ECL cells make up a large proportion of the endocrine cell population, and reside at the base of the oxyntic mucosa (Dockray et al., 1996; Bordi et al., 2000). In the corporeal mucosa, the majority of AiR-IR was localized on D-cells. The Ai receptors that were not on D-cells were scattered throughout the mucosa and were not confined to the basal region. A2AR-IR was not abundant in the corporeal mucosa, and was also scattered throughout the oxyntic mucosa. Since the ECL cells make up a large proportion of the endocrine cell population and are localized only at the base of the oxyntic mucosa, any appreciable Ai or A2A receptor expression on these cells should be apparent. The effect of adenosine on histamine release has not been studied in rats. However, previous studies have shown that adenosine does not alter histamine release in either dog or rabbit (Ainz et al., 1993; Payne and Gerber, 1997). These studies, in combination with our immunohistochemistry results, suggest that adenosine is unlikely to alter histamine release in the rat. Adenosine has been shown to stimulate or inhibit gastric acid secretion in the rat stomach, depending on the route of drug administration. When administered orally or intragastrically, adenosine and its analogs have been shown to decrease basal gastric acid secretion (see Table 2). These routes of administration expose adenosine to the gastric mucosa. Thus, adenosine may mediate its gastric acid inhibitory effect by acting on adenosine receptors in mucosal cells. Although radioligand binding studies have shown extremely low specific binding of H-PIA, a selective Ai receptor agonist, on isolated rat parietal cell preparations (Puurunen et al., 1987), results of the present experiments rule out the possible existence of Ai receptors on parietal cells. Results suggest that adenosine does not act directly on the parietal cells since neither Ai nor A2A receptors were found on these cells. In the present study, parietal cells were identified by the staining of the H+K+-ATPase p subunit of the proton pump. Abundant H+K+-ATPase (3-IR was localized throughout the isthmus, neck, and base of the oxyntic glands, as previously shown (Jiang et al., 2002). Double-staining experiments demonstrate that H+K+-ATPase P-IR was not co-localized with either AiR-IR or A2AR-IR- The absence of adenosine receptors on parietal cells also agrees with previous studies showing that R-PIA (Ai agonist) and NECA (A2 agonist) do not alter carbachol- and histamine-stimulated 14C-aminopyrine accumulation in isolated rat parietal cells (Puurunen et al., 1987). Adenosine analogs also did not alter basal or histamine-stimulated adenylate cyclase activity or carbachol-stimulated inositol phosphate formation in these cells (Puurunen et al., 1987). 1.1 Adenosine Ai receptors on gastrin-secreting G-cells Adenosine has been shown to inhibit gastrin release from the rat stomach (Kwok et al., 1990). The present experiments further demonstrate that this action involves Ai receptors, but not A2 or A3 receptors. In agreement with these observations, our immunohistochemistry experiments demonstrate that AiR-IR was present on all mucosal cells expressing gastrin-IR, while A2AR-IR was not observed on any of these cells. The presence of Ai receptors on G-cells suggests that adenosine may act directly on the G-cells to regulate gastrin release. This direct action may also occur in dogs since adenosine has been shown to inhibit forskolin-stimulated gastrin release through the activation of Ai receptors in primary cultures of canine G-cells (Schepp et al., 1990). Several possible mechanisms may be involved in the Ai receptor-mediated inhibition of gastrin release. The release of gastrin is stimulated by the activation of multiple second messenger pathways. In isolated G-cells, activation of adenylyl cyclase, PLC and PKC are involved in stimulating gastrin release (Campos et al., 1990; Buchan and Meloche, 1994; Seensalu et al., 1997). Activation of Ai receptors has been shown to inhibit adenylate cyclase activity, the most common signaling pathway recognized for the Ai receptor (Fredholm et al., 2001). In G-cells, this second messenger pathway may be activated to inhibit basal gastrin release. Activation of the Ai receptor may also oppose the actions of agents, such as forskolin, that act by stimulating adenylate cyclase. Activation of Ai receptors has also been shown to alter PLC and PKC activity in some tissues (see Table 1). In the stomach, these pathways may also be involved in the inhibitory action of adenosine on gastrin release. The release of gastrin is induced by the influx of extracellular calcium (Buchan and Meloche, 1994). High local concentrations of calcium are required for the exocytosis of gastrin (Oomori et al., 1997), and previous studies showed that gastrin secretion from G-cells requires the activation of L-type Ca2+ channels (Ray et al., 1997). Activation of A\ receptors inhibits L-type channel activity in atrial myocytes (Belardinelli et al., 1995). Thus, these receptors may also inhibit gastrin secretion through the inactivation of these channels. 1.2 Adenosine A] and A2A receptors on somatostatin secreting D-cells The present study demonstrates that both Ai and A2A receptors were involved in modulating somatostatin release. Results of double staining immunohistochemistry experiments further demonstrates that both receptor subtypes were expressed on somatostatin-secreting D-cells of the corpus and antrum, suggesting that adenosine may act directly on D-cells to regulate somatostatin release. In isolated rat oxyntic D-cells, somatostatin release has been shown to involve both phospholipid-dependent and cAMP-dependent signaling pathways (Schaffer et al., 1997). In human oxyntic D-cells, somatostatin release was also stimulated by the activation of adenylate cyclase and PKC pathways (Buchan et al., 1990, 1993). The adenosine A2A receptor is coupled to the Gs protein and has been shown to stimulate adenylate cyclase activity (see Table 1). In striatal neurons, stimulation of this receptor has also been shown to activate the PKC pathway. Thus, A2A receptors may stimulate somatostatin release by activating adenylate cyclase and PKC pathways. Conversely, Ai receptors are coupled to the pertussis toxin-sensitive G protein, G;, and activation of this receptor may inhibit somatostatin release through the inhibition of adenylate cyclase activity. The activation of a pertussis-toxin sensitive G protein has previously been implicated in mediating the inhibitory effect of carbachol on pentagastrin-stimulated somatostatin release in canine oxyntic D-cells (Chiba et al., 1989). Somatostatin is released from D-cell processes by an exocytotic process requiring high local concentrations of Ca2+. In isolated oxyntic D-cells, CCK-stimulated somatostatin release was shown to involve the influx of extracellular Ca through the L-type Ca channels (DelValle et al., 1996). The release of somatostatin from D-cells is also stimulated by membrane depolarization (Patel, 1999). In the present study, confocal microscopy performed at high magnification reveals that most AiR-IR appeared to be concentrated at the end of the D-cell processes. These receptors have previously been localized on presynaptic nerve terminals, and have been shown to inhibit the exocytosis of synaptic vesicles through the inactivation of N-type and L-type Ca2+ channels (Ralevic and Burnstock, 1998). Activation of Ai receptors has also been shown to hyperpolarize neurons by increasing K+ conductance (Belardinelli et al., 1995). Thus, in D-cells, Ai receptors may inhibit somatostatin release by regulating exocytosis or by modulating the membrane potential. 2. Adenosine Ai and A2A receptors of the enteric plexi The localization of gastric Ai and A2A receptors in nerve fibers of the myenteric and submucosal plexi is not surprising since studies have demonstrated adenosine-mediated actions in the intestinal enteric plexi (Burnstock, 2001; Christofi, 2001). Adenosine receptors in this region are likely involved in regulating gastric motility by modulating neurotransmitter release. Numerous studies examining the purinergic control of gastrointestinal motility have been performed in guinea pigs (Christofi, 2001). In general, activation of Ai and A2A receptors has been shown to inhibit and stimulate gastrointestinal motility, respectively. In the rat jejunum, activation of the Ai receptors has been shown to inhibit peristalsis (Hancock and Coupar, 1995), and in the ileum, these receptor have been shown to enhance descending relaxation (Storr et al., 2002). In the rat stomach, activation of Ai receptors in the enteric plexi has been shown to mediate gastric relaxation (Matharu and Hollingsworth, 1992). Conversely, activation of A2 receptor has been suggested to potentiate vagally-stimulated gastric contractions (Cho et al., 1993). Thus, adenosine Ai and A2A receptors are involved in the inhibitory (relaxation) and stimulatory (contraction) control of gastrointestinal motility, respectively. The effect of adenosine on gastrointestinal motility is likely mediated by its action on presynaptic and postsynaptic neurotransmission in the enteric plexi. Most studies that examine the neuromodulatory role of adenosine in the intestines have been performed in the guinea pig ileum, in particular, the guinea pig myenteric plexus-longitudinal muscle preparations. In the submucosal plexus of the guinea pig ileum, electrophysiological studies have shown that adenosine inhibits enteric neurotransmission by suppressing the excitability of myenteric neurons (Zafirov et al., 1985; Palmer et al., 1987). This inhibitory effect was mediated by presynaptic Ai receptors on myenteric neurons (Christofi and Wood, 1993). Similarly, activation of presynaptic Ai receptors were shown to inhibit neurotransmitter release from submucosal neurons (Barajas-Lopez et al., 1991). In the guinea pig ileum, adenosine inhibits acetylcholine release from enteric nerves (Sawynok and Jhamandas, 1976; Hayashi et al., 1978; Gustafsson et al., 1985). This inhibition was presynaptic in nature (Gustafsson et al., 1985; Christofi and Cook, 1986). Ai receptor-mediated presynaptic inhibition of noradrenaline (Barajas-Lopez et al., 1991), neurokinin A (Broad et al., 1992) and substance P (Broad et al., 1993) release from enteric nerves endings has also been demonstrated. In addition to its inhibitory effect on neurotransmitter release, adenosine may stimulate neurotransmitter release by acting directly on enteric neurons. This postsynaptic excitatory effect was shown to be mediated by A2A receptors (Barajas-Lopez et al, 1991). Although the effect of adenosine on the intestines has been studied extensively, the effect of this nucleoside is not well-studied in the stomach. In the gastric antrum of the guinea pig, adenosine and its analogs have been shown to inhibit nicotinic transmission in the myenteric plexus by acting on presynaptic adenosine receptors (Christofi et al., 1992). Furthermore, adenosine and its analogs have been shown to inhibit fast EPSPs in approximately 60% of gastric neurons (Christofi et al., 1992). Thus, the relatively high level of AiR-IR and A2AR-IR staining observed in the myenteric plexus of the rat stomach in the present study fits well with the known modulatory effects of adenosine on neurotransmission. In the rat stomach, adenosine has been demonstrated to protect against ethanol-induced gastric lesion formation (see Chapter 1, Section 3), but the mechanism involved has not been determined. Recent studies have shown that inhibition of gastric motility by anti-muscarinic drugs can protect the stomach against lesion formation induced by ethanol (Schicho et al., 2003). Thus, the Ai receptor-mediated inhibition of acetylcholine release may be involved in protecting the stomach against ethanol injury. Adenosine has also been shown to regulate gastric IRG and SLI release. Similar to its action on motility, the effect of adenosine on the release of these gastric peptides may be secondary to its effect on adrenergic and cholinergic neurotransmission. Adrenergic and cholinergic agents have been shown to alter the release of IRG and SLI (Saffouri et al, 1980; Koop et al., 1982, 1983; Schubert and Makhlouf, 1992). Previous experiments performed in our laboratory have shown that the adenosine-induced release of SLI was not altered in the presence of the cholinergic blockers, atropine and hexamethonium, or the P-adrenergic blocker, propranolol (Kwok et al., 1990). Thus, adenosine is unlikely to modulate SLI release by acting on adrenergic or cholinergic nerves. The present study shows that adenosine receptors were expressed in the enteric plexi. Since various non-adrenergic, non-cholinergic nerve fibers are present in the plexi, it is possible that adenosine stimulates SLI release indirectly by modulating non-adrenergic, non-cholinergic neurotransmission. CGRP and VIP have been demonstrated to stimulate gastric somatostatin secretion (Chiba et al., 1980; Manela et al, 1995; Rasmussen et al., 2001). While the effect of adenosine on CGRP and VIP release has not been examined in the stomach, endogenous adenosine has been shown to tonically control CGRP and VIP release in the CNS (Sebastiao and Ribeiro, 2000). The co-localization of A2AR-IR with somatostatin-IR in some nerves of the myenteric plexus and circular muscle observed in the present study suggests that activation of A2A receptors may elicit somatostatin release from nerve fibers. However, most of the somatostatin-IR was shown to reside in the D-cells, as previously reported (Vinik et al., 1981; Ekblad et al., 1985; Keast et al., 1985). Only sparse somatostatin staining was observed in the myenteric plexus and on nerve fibers. These somatostatin-IR fibers do not appear to project to the mucosa, and most of these fibers do not express A2AR-IR. Thus, adenosine is more likely to act directly on mucosal D-cells to elicit SLI release. The possibility that adenosine mediates its inhibitory effect on IRG release through adrenergic or cholinergic mechanisms has not been examined. It is also possible that adenosine mediates its inhibitory actions on IRG release by modulating the release of CGRP and VIP. These neurotransmitters have been shown to inhibit IRG release (Chiba et al., 1980; Manela et al., 1995). However, the localization of AiR on all gastrin-secreting G-cells of the mucosa demonstrated by the present study suggests that adenosine may inhibit gastrin secretion, at least in part, by acting on Ai receptors on the G-cells. This proposal is supported further by the ability of adenosine to inhibit gastrin release from canine G-cells (Schepp et al., 1990) and rat antral mucosal cells in culture (DeSchryver-Kecskemeti et al., 1981). 3. Adenosine Ai and A2A receptors of the gastric vasculature The vascular localization of the gastric adenosine receptors has been suggested by a number of studies demonstrating the action of adenosine on gastric mucosal blood flow (Cho and Ogle, 1990; Cho et al., 1991; Cho et al., 1995; Nagata et al., 1996). In the pulmonary artery (Biaggioni et al., 1989) and the renal vasculature (Holz and Steinhausen, 1987), activation of Ai receptors has been shown to mediate vasoconstriction. Conversely, activation of A2A receptors has been shown to mediate relaxation in coronary and pulmonary arteries, renal vasculature and the aorta (Tabrizchi and Bedi, 2001). In the rat stomach, subcutaneous administration of adenosine has been shown to increase gastric mucosal blood flow (Cho et al., 1991; Cho et al., 1995). Adenosine has been shown to induce vasodilation in submucosal microvessels of the stomach by acting on A2 receptors (Gerber and Guth, 1989; Nagata et al., 1996). This vasodilatory action may be involved in protecting the stomach against ethanol-induced gastric injury since the administration of ethanol has previously been shown to vasoconstrict submucosal blood vessels (Yonei and Guth, 1991). The perfusion of CGS 21680 into the isolated vascularly perfused rat stomach has also been demonstrated to decrease the gastric perfusion pressure (Yip and Kwok, 2004), suggesting that the A2A receptors are involved in mediating gastric vasodilation. Adenosine has been proposed to act on A2A receptors localized on vascular smooth muscle cells and endothelial cells (Tabrizchi and Bedi, 2001). In the rat mesenteric arterial bed, adenosine-induced vasodilation was shown to involve A2A receptors localized on both of these cell types (Hiley et al., 1995). The immunohistochemical localization of the Ai and A2A receptors has previously been demonstrated in the vascular smooth muscle cells and endothelial cells of the human skeletal muscle (Lynge and Hellsten, 2000). Results of the current study clearly demonstrate that, in the stomach, both Ai and A2A receptors are localized on vascular endothelial cells and are also likely expressed on vascular smooth muscle cells. v The vasodilatory actions of adenosine in the stomach are mainly attributed to activation of A2A receptors. An Ai receptor-mediated action in the gastric vasculature has not yet been demonstrated. In the renal vasculature, the effect of adenosine was concentration-dependent. At low micromolar concentrations, adenosine elicited vasoconstriction via activation of Ai receptors, while at higher concentrations (>10 |JM), adenosine elicited vasodilation via activation of A2 receptors (Tabrizchi and Bedi, 2001). In the rat gastric submucosal vessels, adenosine-induced vasodilation was demonstrated using >10 uM adenosine (Nagata et al., 1996). The effect of lower concentrations of adenosine has not been examined. The regulation of vascular tone by the Ai receptor remains to be confirmed in the stomach. In summary, the results of the immunohistochemistry study provide several new and significant insights into the distribution and localization of adenosine receptors. They confirm the widespread distribution of Ai and A2A receptor expression in the enteric plexi and vasculature, and further support the suggestion that adenosine acts on these anatomical structures. The localization of Ai and A2A receptors on the G-cells and D-cells support the proposal that adenosine may act directly on these cells to modulate gastrin and somatostatin release, respectively. The lack of adenosine receptors on the parietal cells provides further evidence that adenosine does not regulate gastric acid secretion by acting directly on the parietal cells. V. Conclusion This study provides fundamental information on the functional role of Ai and A2A receptors in the rat stomach. Results suggest that adenosine may be an important regulator of gastric acid secretion. Unlike in other species, adenosine does not alter gastric acid secretion by acting directly on parietal cells in the rat. This lack of effect is confirmed in the present study by demonstrating the absence of AiR-IR and A2AR-IR on rat parietal cells. Instead, this study suggests that adenosine likely regulates gastric acid secretion by acting on Ai receptors to inhibit gastrin and A2A receptors to stimulate somatostatin release. The localization of A\ and A2A receptors on the G-cells and D-cells suggests that adenosine may act directly on these cells to modulate gastrin and somatostatin release. Since these receptors were also present on nerve fibers throughout the enteric plexi, the possibility that adenosine may modulate gastrin and somatostatin indirectly by altering neurotransmission cannot be ruled out. The localization of A] and A2A receptors in the vasculature and enteric plexi also agrees with the known actions of adenosine in regulating gastric mucosal blood flow and motility. The mucosal adenosine Ai and A2A receptors were shown to be structurally identical to those in the brain, and expressed in extremely low levels, using Real-Time RT-PCR and competitive RT-PCR. Real-Time RT-PCR was also used to examine the relationship between altered physiological states of the stomach and gastric Ai and A2A receptor gene expression. Like gastrin and somatostatin, the gene expression of Ai and A2A receptors is regulated by the gastric luminal environment. Fasting and omeprazole treatment were shown to alter adenosine receptor gene expression, most likely by changing the intraluminal acidity. Since these receptors are localized on G-cells and D-cells, changes in their expression may alter gastrin and somatostatin synthesis and secretion. Changes in adenosine receptor gene expression can be accompanied by corresponding changes in receptor function, suggesting that adenosine receptor expression may be determined, in part, by its gene expression. The present experiments provide fundamental information regarding the function, distribution, and cellular localization of the gastric Ai and A2A receptors. Furthermore, this study suggests that under different physiological states of the stomach adenosine receptor expression may be altered to modulate gastrin and somatostatin synthesis and release, and, therefore, gastric acid secretion. 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Appendix 1: Table of abbreviations 2-CA 2-chloroadenosine 8-PT 8-phenyltheophylline 8-SPT 8-(p-sulfophenyl)theophyline A,R-IR A, receptor immunoreactivity A2AR-IR A2A receptor immunoreactivity ADA adenosine deaminase ADP adenosine diphosphate AK adenosine kinase AMP adenosine monophosphate ATP adenosine triphosphate BSA bovine serum albumin cAMP cyclic AMP CCK cholecystokinin CGRP calcitonin gene related peptide CGS 15943 9-chloro-2(2-furyl)[l,2, 4]triazolo [ 1,5-c]quinazolin-5-amine CGS 21680 2-p-(2-carboxyethyl)phenethylamino-5 'N-ethylcarboxamidoadenosine CHA 5'-N -cyclohexyladenosine CPA N6-cyclopentyladenosine CRE cAMP response element CREB CRE binding protein CT threshold cycle CV1808 2-phenylaminoadenosine DEPC diethlypyrocarbonate DMNV dorsomotor nucleus of the vagus DMPX 3,7-dimethyl-1 -propargylxanthine DMSO dimethylsulfoxide DPCPX 8-cyclopentyl-l,3-dipropylxanthine ECL enterochro maffin-like EGF epidermal growth factor EHNA erythro-9-(2-hydroxy-3 -nonyl)adenine hydrochloride E-NPP ecto-nucleotide pyrophosphatase E-NTPase ecto-nucleoside triphosphate diphoshohydrolase EPSP excitatory postsynaptic potential FAM 6-carboxyfluorescein FL full length GAPDH glyceraldehyde-3 '-phosphate dehydrogenase GRP gatrin releasing peptide i.c.v. intracerebroventricular i.d. intraduodenal i.p. intraperitoneal i.v. intravenous IB-MECA l-deoxy-l-[6-[[(3-iodophenyl)methyl] amino] -9H-purin-9-yl]-N-methyl-B-D-ribofuranuronamide IHC immunohistochemistry IR immunoreactivity IRG immunoreactive gastrin IUPHAR International Union of Pharmacology LB Luria-Bertani MAPK mitogen activated protein kinases NAD+ nicotinamide adenine dinucleotide NECA N-ethylcarboxamidoadenosine NFKB nuclear factor KB NSB non-specific binding NTS nucleus tractus solitarius PCR polymerase chain reaction PGP protein gene product PIA Nb-(2-phenylisopropyl)adenosine PKC protein kinase C PLC phospholipase C RIA radioimmunoassay Rn reporter emissions R-PIA R(-)-N6-(2-phenylisopropyl)adenosine RT-PCR reverse-transcriptase PCR s.c. subcutaneous SAH S-adenosyl homocysteine SAHH S-adenosyl homocysteine hydrolase S-adenosyl methionine SL short length SLI somatostatin-like immunoreactivity S-PIA (S)-N6-(2-phenylisopropyl) adenosine TAMRA 6-carboxytetramethylrhodamine TC total count Tm melting temperature UTR untranslated region VIP vasoactive intestinal peptide VWF von Willebrand's factor ZM 241385 4-(2-[7-Amino-2 (2furyl)[l,2,4]triazolo [2,3-a][l,3,5]triazin-5ylamino]ethyl)phenol 

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