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GASTROINTESTINAL, HEPATIC, PULMONARY, AND RENAL

Role of Adenosine A₁ Receptor in the Regulation of Gastrin Release

Department of Physiology, University of British Columbia, Vancouver, British Columbia, Canada

Received February 6, 2004; accepted March 24, 2004.

	Abstract

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Adenosine has been demonstrated to inhibit gastric acid secretion. In the rat stomach, this inhibitory effect may be mediated indirectly by the inhibition of gastrin release. Results show that the A₁ receptor agonist N⁶-cyclopentyladenosine (CPA) suppressed immunoreactive gastrin (IRG) release in a concentration-dependent manner. CPA significantly inhibited IRG release at 0.001 µM and maximally inhibited IRG release at 1 µM. At concentrations of 0.001 to 0.1 µM, the A_2A receptor-selective agonist 2-p-(2-carboxyethyl)phenethylamino-5'-N-ethylcarboxamidoadenosine and A₃ receptor-selective agonist 1-deoxy-1-[6-[[(3-iodophenyl)methyl]amino]-9H-purin-9-yl]-N-methyl--D-ribofuranuronamide, had no effect on IRG release, suggesting the involvement of A₁ receptors. In agreement, the A₁ receptor-selective antagonist 8-cyclopentyl-1,3-dipropylxanthine abolished adenosine-induced inhibition of IRG release. Results of immunohistochemistry experiments reveal the presence of A₁ receptor immunoreactivity on mucosal G-cells and D-cells, and the gastric plexi, but not parietal cells, suggesting that adenosine may act directly on G-cells or indirectly on the gastric plexi to modulate IRG release. The structure of the mucosal A₁ receptor was found to be identical to that in the rat brain. Alternative splicing within the coding region of this receptor did not occur. A real-time reverse transcription-polymerase chain reaction assay was developed to measure gastric A₁ receptor gene expression. The highest level of gastric A₁ receptor mRNA was found in the corporeal muscle. However, this level was significantly lower in comparison with the striatum. In conclusion, this study shows that adenosine may suppress IRG release, at least in part, by activating A₁ receptors localized on G-cells and may consequently result in an inhibition of gastric acid secretion.

Adenosine elicits its effects by acting on G protein-coupled receptors belonging to the purinergic P1 receptor family. These receptors are classified into adenosine A₁, A_2A, A_2B, and A₃ subtypes based on their pharmacological and structural properties (Fredholm et al., 2001). In rats, the adenosine receptor subtype(s) involved in modulating IRG release has not been characterized. Thus, the first objective of the present study was to determine the adenosine receptor subtype involved in the inhibitory action of adenosine on IRG release in the isolated vascularly perfused rat stomach using selective adenosine agonists and antagonists.

A low level of A₁ receptor gene expression has previously been found in the whole rat stomach using reverse-transcription polymerase chain reaction (RT-PCR) (Dixon et al., 1996). However, the expression of this receptor in functionally distinct regions of the stomach is undetermined. Although our preliminary results suggest that adenosine A₁ receptors may be involved in the inhibition of IRG release, the cellular localization of these receptors on specific cells of the stomach, such as the gastrin-secreting G-cells, has also not been examined. Therefore, the second objective of this study was to determine the cellular localization, distribution, and gene expression level of the A₁ receptor in the rat stomach using immunohistochemistry, RT-PCR, and real-time RT-PCR, respectively. The adenosine A₁ receptor has been cloned in tissues of various species, including the human and rat brain (Ralevic and Burnstock, 1998), but structural information regarding the rat gastric A₁ receptor is lacking. Northern blot analysis has demonstrated the presence of two A₁ receptor mRNA transcripts in the rat stomach (Mahan et al., 1991; Reppert et al., 1991). However, it is unclear whether these transcripts differ in their coding regions. Differences in the coding region may result in the production of multiple forms of the receptor. Previous studies have shown that multiple variants of the rat A₃ receptor were generated by alternative splicing in the coding region of the A₃ receptor gene (Sajjadi et al., 1996). Because multiple forms of the A₁ receptor may have important functional implications, the final objective of this study was to clone and sequence the entire coding region of the mucosal A₁ receptor gene.

	Materials and Methods

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Stomach Perfusion
Animals were treated in accordance with the guidelines of the University of British Columbia Committee on Animal Care. Male Wistar rats (250–325 g) were housed in light- and temperature-controlled rooms with free access to food and water. Before stomach perfusion, animals were deprived of food but not water for at least 14 h. 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 been described previously (Pederson et al., 1984; Kwok et al., 1990). The stomach was exposed by an abdominal midline incision. The superior mesenteric artery and vasculatures supplying the left and right adrenal glands and kidneys were occluded or cut between double ligatures. The spleen and pancreas were then dissected along the greater curvature of the stomach. Care was taken to preserve the right gastroepiploic artery. A cannula was secured into the gastroduodenal junction to allow for drainage of gastric contents. The pancreas, spleen, and 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, followed by an injection of 2 ml of saline containing 600 U of heparin (Sigma-Aldrich, St. Louis, MO). Venous effluent was collected via a portal vein cannula. Stomach preparations were equilibrated for 30 min before 5-min samples were collected into ice-cold scintillation vials containing 0.3 ml of Trasylol (aprotinin, 10,000 KIU/ml; Miles Labs, Etobicoke, ON, Canada). Aliquots (1 ml) were immediately transferred into ice-cold test tubes and stored at –20°C until assayed.

The stomach was perfused at a rate of 3 ml/min using a peristaltic pump (Cole-Parmer Instrument Co., Vernon Hills, IL). The perfusate was composed of Krebs' solution (120 mM NaCl, 4.4 mM KCl, 2.5 mM CaCl₂, 1.2 mM MgSO₄·7H₂O, 1.5 mM KH₂PO₄, 25 mM NaHCO₃, and 5.1 mM dextrose) containing 0.2% BSA [radioimmunoassay (RIA) grade; Sigma-Aldrich] and 3% dextran (clinical grade; Sigma-Al-drich). The perfusate was continuously gassed with a mixture of 95% O₂ and 5% CO₂ to maintain a pH of 7.4. Both the perfusate and the preparation were kept at 37°C by thermostatically controlled heating units throughout the experiment. Drugs were introduced 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 hemisulfate salt, N⁶-cyclopentyladenosine (CPA), 2-p-(2-carboxyethyl)phenethylamino-5'N-ethylcarboxamidoadenosine HCl (CGS 21680), 1-deoxy-1-[6-[[(3-iodophenyl)methyl-]amino]-9H-purin-9-yl]-N-methyl--D-ribofuranuronamide (IB-MECA), 8-cyclopentyl-1,3-dipropylxanthine (DPCPX), and 3,7-dimethyl-1-propargylxanthine (DMPX). Adenosine analogs were first dissolved in a small volume of dimethyl sulfoxide (BDH, Toronto, ON, Canada) and subsequently diluted with saline or perfusate to 0.03 or 0.5% before perfusing into the stomach. At these concentrations, dimethyl sulfoxide did not alter basal IRG release.

RIA and Data Analysis
The specific RIA used for the measurement of IRG has been described previously (Jaffe and Walsh, 1978; Fujimiya and Kwok, 1997). The gastrin antibody (PM1) was kindly provided by Dr. R. Pederson (Department of Physiology, University of British Columbia). The drugs used in the present study did not cross-react with this antibody. The inter- and intra-assay variations were shown to be less than 6 and 4%, respectively.

Although the basal rate of IRG release varied among animals, previous experiments have demonstrated that basal IRG release was maintained in the perfused rat stomach (Pederson et al., 1984; Kwok et al., 1990). Therefore, results were expressed as mean ± S.E.M. of IRG release (percentage), which was calculated as follows: [IRG release (picograms per minute) during a 5-min period ÷ IRG release (picograms per minute) during period 1] x 100. To compare the effect of analogs, results were also expressed as percentage of inhibition (IRG release), which was calculated as follows: [mean basal IRG release (periods 1–3) – mean IRG release in the presence of drug (periods 4–7)] picograms per minute ÷ [mean basal IRG release (periods 1–3)] picograms per minute x 100. Percentage of change was calculated as follows: [mean IRG release in the presence of drug (periods 4–7) – mean basal IRG release (periods 1–3)] picograms per minute ÷ [mean basal IRG release (periods 1–3)] picograms per minute x 100. Statistical significance (P < 0.05) was determined using one-way analysis of variance followed by Dunnett's multiple comparison test, and the paired or unpaired Student's t test when appropriate. Statistics and estimation of the EC₅₀ were performed using GraphPad Prism (version 3.0, GraphPad Software Inc., San Diego, CA).

Immunohistochemistry
Gastric corporeal and antral tissues from male Wistar rats were fixed overnight, cryoprotected, and sectioned as described previously (Yip et al., 2003). Free-floating sections (30 µm) were sequentially incubated in 0.1 M PBS containing 50 mM NH₄Cl (30 min), 0.1 M PBS containing 100 mM glycine (10 min), and blocking buffer (0.1 M PBS containing 1% BSA and 0.3% Triton X-100, 1 h). Antibodies were diluted in blocking buffer containing 0.1% sodium azide. Tissue sections were incubated with the rabbit anti-A₁ adenosine receptor antibody (1:500; Sigma-Aldrich) for 72 h at 4°C, washed with 0.1 M PBS (3 x 15 min), and then incubated with donkey anti-rabbit IgG conjugated to cyanine Cy3 (1:2000; Jackson ImmunoResearch Laboratories Inc., West Grove, PA) overnight at 4°C. Sections were again washed and either double stained or mounted onto glass slides. For double staining, tissue sections were incubated with another primary antibody for 72 h at 4°C, washed, and incubated with Alexa Fluor 488-conjugated secondary antibody overnight at 4°C. Sections were then washed, mounted onto glass slides, coverslipped using a mixture of 0.1 M PBS in glycerin (1:9), and sealed with nail polish.

Control Experiments for Immunohistochemistry
The A₁ receptor antibody has been shown to cross-react with rat A₁ receptors, and its specificity has previously been examined in rat tissues (Carruthers et al., 2001). Western blotting and immunoneutralization experiments were also performed in the present study to examine the specificity of this antibody on rat gastric tissues. Additional negative control experiments were also performed to confirm that A₁ receptor-immunoreactivity (A₁R-IR) does not result from nonspecific binding of the secondary antibodies.

Western Blotting. Protein was extracted from the rat fundus, corpus, antrum, and the brain using TRIzol reagent, according to manufacturer's instructions. Protein samples were redissolved in 1% sodium dodecyl sulfate at 50°C, and concentrations were determined using the BCA assay kit (Pierce Chemical, Rockford, IL). Protein samples (10 µg) were denatured in 1x sodium dodecyl sulfate gel loading buffer [50 mM Tris-Cl (pH 6.8), 100 mM dithiothreitol, 2% sodium dodecyl sulfate, 0.1% bromphenol blue, and 10% glycerol] by boiling for 5 min. Samples were loaded and resolved on polyacrylamide gels composed of a 4% stacking and 10% separating gel, and then transferred to polyvinylidene fluoride membranes (Bio-Rad, Hercules, CA). The methods used for electrophoresis and immunoblotting have been described previously (Sambrook et al., 1989). Polyvinylidene fluoride membranes were washed in Tris-buffered saline (TBS) and incubated in blocking buffer (TBS containing 0.1% Tween 20 and 5% BSA) for 1 h at room temperature with shaking. Membranes were then washed and incubated with the anti-A₁ adenosine receptor antibody overnight at 4°C. This primary antibody was diluted to 1:5000 with TBS containing 0.1% Tween 20 (TBST) before use. Membranes were washed in TBS and then incubated in TBST containing anti-rabbit horseradish peroxidase-conjugated secondary antibody (1:20,000; Jackson ImmunoResearch Laboratories Inc.) for 1 h at room temperature with shaking. Membranes were then washed in TBST and enhanced chemiluminescence immunodetection was performed according to manufacturer's instructions (Amersham Biosciences UK, Ltd., Little Chalfont, Buckinghamshire, UK). Results show that two immunoreactive products with apparent molecular mass of 40.5 and 67 kDa were detected (Fig. 1). These findings are consistent with the results of Western blot experiments performed in rat trigeminal ganglion neurons using the same antibody (Carruthers et al., 2001). The larger immunoreactive product likely represents the glycosylated form of the A₁ receptor because deglycosylation can shift the molecular mass of the larger product to 40 kDa, the expected size of the A₁ receptor (Carruthers et al., 2001).

View larger version (26K): [in this window] [in a new window]   Fig. 1. Western blot analysis of adenosine A receptors in rat tissues. Two immunogenic bands at 40.5 and 67 kDa were 1detected in protein extracts of the fundus, corpus, antrum, and brain.

Immunoneutralization of the A₁ Receptor Antibody. Immunoneutralization experiments were also performed in the present study to confirm the specificity of the antibody. Because the blocking peptide for the A₁ receptor antibody is unavailable from the supplier, one was synthesized by the Nucleic Acid Protein Services Unit at University of British Columbia. The amino acid sequence Cys-Gln-Pro-Lys-Pro-Pro-Ile-Asp-Glu-Asp-Leu-Pro-Glu-Glu-Lys-Ala-Glu-Asp of this synthetic peptide was based on the immunogen sequence provided by Sigma-Aldrich. The lack of homology between the sequence of the blocking peptide and other proteins was confirmed using the BLAST software featured at the National Institutes of Health National Center for Biotechnology Information Web site. The only proteins that expressed high homology (>70%) with this peptide are A₁ receptors from various species. The rat A_2A, A_2B, and A₃ receptor sequences exhibit no significant similarity. The blocking peptide was dissolved in 0.1 M PBS (pH 7.4), and the concentration was determined using the BCA assay. The A₁ receptor antibody was neutralized by incubating 1 µl of the stock antibody with 0.1 µgofthe blocking peptide in 0.5 ml of 0.1 M PBS containing 1% BSA, 0.3% Triton X-100, and 0.1% sodium azide for 48 h at 4°C. A positive control lacking the blocking peptide was also prepared. Tissues stained with the neutralized antibody did not demonstrate A₁R-IR, whereas tissues stained with the positive control demonstrated extensive A₁R-IR (Fig. 5).

View larger version (74K): [in this window] [in a new window]   Fig. 5. Confocal images showing A1R-IR in the antrum and corpus of the rat stomach. A, A1R-IR cell in the antral mucosa (arrows). B, boxed area in A, viewed under high magnification. C, A1R-IR cells in the corporeal mucosa (arrows). D, A1R-IR cells in the corporeal mucosa, viewed under high magnification. E, A1R-IR on blood vessels (arrow; BV) and nerve fibers (arrowheads) in the submucosal plexus (SMP) of the corpus. F, A1R-IR on nerve fibers and cell bodies of the myenteric plexus (arrowhead), and on nerve fibers of the circular (arrows; CM) and longitudinal muscle (arrows; LM) of the antrum. G, confocal image showing the lack of A1R-IR in the antral mucosa; section was incubated with the immunoneutralized A1 receptor antibody. H, confocal image showing the lack of A1R-IR in the corporeal muscle; section was incubated with the secondary antibody, donkey anti-rabbit IgG conjugated to cyanine Cy3 (1:2000) alone (z-step, 1.0 µm and scale bar, 25 µm for all images).

Controls for Nonspecific Binding. Additional control experiments were also performed to ensure that nonspecific binding did not occur. Experiments included incubating sections with 1% BSA in place of the primary antibody, incubation without the secondary antibody, or incubation with Alexa 488-conjugated anti-mouse IgG secondary antibody or Cy3-conjugated anti-rabbit IgG secondary antibody alone. No immunoreactivity (IR) was observed after these procedures (Fig. 5).

Double Staining
For double-staining experiments, monoclonal primary antibodies against somatostatin (1:500, Soma 8; MRC Regulatory Peptide Group, University of British Columbia), gastrin (1:30,000, 109-21 provided by the late Dr. John Walsh), protein gene product 9.5 (PGP 9.5) (1:200, ab8189; Abcam Limited, Cambridge, UK), human von Willebrand's factor (VWF) (1:50; Serotec, Oxford, UK), and H⁺K⁺-ATPase (1:2000; Affinity Bioreagents, Golden, CO) were used.

Confocal Microscopy
Stained tissue sections were viewed using the Radiance 2000 confocal scanning laser system (Bio-Rad) mounted on a Nikon Eclipse TE300 inverted microscope. A krypton gas laser with an excitation wavelength of 568 nm and emission filter of 575 to 625 nm (for visualization of Cy3), and an excitation wavelength of 488 and emission filter of 500 to 530 nm (for visualization of Alexa Fluor 488) was used. Bleed-through was not detected for any of the antibodies. Lens magnification of 40x was used with a zoom factor of 1.0 and z-step of 0.5 to 1.0 µm, whereas lens magnification of 60x was used with a zoom factor of 1.0 to 1.6 and a z-step of 0.3 to 0.5 µm. The software program Lasersharp 2000 (version 4.1; Bio-Rad) was used to scan tissue sections sequentially using the red and green collection channels and the Kalman collection filter (n = 2). Images with a resolution of at least 512 x 512 pixels were obtained and then analyzed using NIH Image (National Institutes of Health, Bethesda, MD) and Adobe Photoshop (version 7.0; Adobe Systems, San Jose, CA). To determine whether colocalization occurred, the image collected from the red channel (A₁R-IR) was overlaid on the image collected from the green channel (somatostatin, gastrin, VWF, PGP 9.5, or H⁺K⁺-ATPase -IR) using Adobe Photoshop. Colocalization of A₁R-IR with gastrin-IR and somatostatin-IR was quantified in the antral and/or corporeal mucosa by examining at least three tissue sections from four different animals.

RT-PCR
Primer Design and Synthesis. PCR primers were designed based on previously published rat brain A₁ receptor cDNA sequences (accession no. M64299 [GenBank] ) (Mahan et al., 1991) using the software program PCGene (IntelliGenetics, Mountain View, CA). The forward and reverse primer sequences were 5'GTCTGCTGATGTGCC CAGCT3' (corresponding to position 322–341 bp of the rat brain A₁ receptor cDNA sequence) and 5'ACAGGGTGGGACAGGGAGAA3' (corresponding to position 1388–1407 bp), respectively. The amplicon generated (1086 bp) spans the entire coding region of the A₁ receptor gene. The primers were synthesized by the Nucleic Acid Protein Services Unit at University of British Columbia.

Tissue and Total RNA Extraction. Male Wistar rats (200–250 g) were anesthetized with an i.p. injection (60 mg/kg) of Somnotol. 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 gastric mucosa was obtained by gently scraping the luminal surface of the stomach using a sterile glass slide. Total RNA was extracted immediately from the mucosa and striatum. The latter tissue has been shown to express moderate levels of A₁ receptor mRNA (Dixon et al., 1996) and was used as a positive control. Total RNA was extracted from tissues using TRIzol reagent (Invitrogen, Carlsbad, CA) according to manufacturer's instructions. The total RNA concentration was determined by the following calculation: concentration (micrograms per milliliter) = A₂₆₀ x 40 µg/ml x 100 (dilution factor).

DNase I Treatment, First Strand cDNA Synthesis, and PCR. DNase I treatment was performed at room temperature in 1x first strand buffer [50 mM Tris-HCl (pH 8.3 at 25°C), 75 mM KCl, 3 mM MgCl₂] containing 1 U of DNase I/µg total RNA (Invitrogen), according to manufacturers' instructions. First strand cDNA was synthesized from 5 µg of DNase I-treated total RNA using Superscript II RNase H-Reverse Transcriptase (Invitrogen). A sample containing autoclaved distilled water instead of total RNA was used as the negative control.

The PCR reaction mixture (50 µl) consisted of 2 µl cDNA in 1x PCR buffer [20 mM Tris-HCl (pH 8.4), and 50 mM KCl] containing 0.2 mM dNTP mix, 1.5 mM MgCl₂, 100 ng each of forward and reverse primer, and 1 U of Platinum TaqDNA Polymerase (Invitrogen). A positive control sample containing striatal cDNA and a negative control from the first strand synthesis step were included for all experiments. The PCR was performed using the Robocycler Temperature cycler (Stratagene, La Jolla, CA). Thirty cycles of amplification were performed. Each cycle consisted of a 45-s denaturation period at 94°C, a 1-min annealing period at 58°C, and a 1-min extension period at 72°C. PCR products were electrophoresed through a 2.5% agarose gel (w/v) containing 1x Tris acetate EDTA buffer [40 mM Tris acetate and 1 mM EDTA (pH 8.5)] and ethidium bromide (0.5 µg/ml). Gels were run in 1x Tris acetate EDTA buffer at 100 V for 45 min, and visualized and photographed under UV light using the Stratagene Eagle Eye II system (La Jolla, CA).

Cloning and Sequencing of the Mucosal A₁ Receptor Gene. The A₁ receptor RT-PCR product generated using mucosal tissue as the template was ligated into the pGEM-T vector (Promega, Madison, WI) and transformed into DH-competent Escherichia coli cells (Invitrogen). Cells were grown in LB plates containing ampicillin (100 µg/ml), isopropyl -D-thiogalactopyranoside (0.5 mM), and 5-bromo-4-chloro-3-indolyl--D-galactoside (80 µg/ml). Plasmid DNA was purified using the QIAprep Miniprep kit (QIAGEN, Mississauga, ON, Canada) and sequenced at the Nucleic Acid Protein Services Unit using the T7 primer and SP6 primer. The gene sequence of the mucosal A₁ receptor was aligned with the published sequence in the rat brain using the University of Southern California Sequence Alignment Server (http://www-hto.usc.edu/software/seqaln/seqaln-query.html) and submitted to the GenBank database at the National Center for Biotechnology Information.

Quantification of A₁ Receptor Gene Expression by Real-Time RT-PCR. A two-step real-time RT-PCR assay was performed to quantify A₁ receptor gene expression in various regions of the rat stomach. For comparison, A₁ receptor gene expression was also measured in the rat striatum. Primers and probes were designed using the Primer Express Sequence Design software program (version 1.0; Applied Biosystems, Foster City, CA). The reporter dye 6-carboxyfluorescein and the quencher dye 6-carboxytetramethylrhodamine were linked to the 5' and 3' ends of the A₁ receptor probe, respectively. The sequences of the forward primer, reverse primer, and probe are 5'CGGTGACCCCCAGAAGTACTAC3',5'GGGCAAAGAGGAAGAGGATGA3', and 6-carboxyfluorescein-5'CAGCGACTTGGCGATCTTCAGCTCCT3'-6-carboxytetramethylrhodamine, which correspond to positions 963 to 984, 1039 to 1059, and 989 to 1014 bp of the rat brain A₁ receptor gene (accession no. M64299 [GenBank] ), respectively. The primer and probes were synthesized by the Nucleic Acid Protein Services Unit at University of British Columbia and by Synthegen, LLC (Houston, TX), respectively.

RNA transcripts expressing the entire coding region of the A₁ receptor were used as the standard for quantification during real-time RT-PCR. The standard was synthesized using plasmids generated by the previous cloning experiments by in vitro transcription using the Riboprobe in vitro transcription kit and T7 RNA polymerase (Promega). The A₁ receptor RNA standard was DNase I-treated and purified, and its concentration was determined using the RiboGreen reagent quantitation kit (Molecular Probes, Eugene, OR), the FL600 microplate fluorescence reader (Bio-Tek Instruments, Winooski, VT) and the KC4 Kineticalc Software (version 2.6; Bio-Tek Instruments). RNA standards were then serially diluted to concentrations of 1 x 10³ to 1 x 10¹² copies/µl in RNase-free water, aliquoted, stored at –80°C, and thawed only once before use.

Two-Step Real-Time RT-PCR
The level of A₁ receptor gene expression was measured in the whole fundus, corpus, antrum, the corporeal mucosa and corporeal muscle layers, the whole stomach mucosa, and the striatum (positive control). The methods for tissue extraction were described above. To obtain the corporeal mucosa and muscle tissue, the corporeal mucosa was gently scraped off the luminal surface of the corpus using a sterile glass slide. The remaining corporeal muscle tissue was flash frozen in liquid nitrogen and stored at –80°C until total RNA was isolated. Total RNA was extracted immediately from corporeal and whole stomach mucosa samples. The methods used for the isolation, quantification, and DNase I-treatment of total RNA were described above.

Reverse Transcription. One microgram of DNase I-treated tissue RNA was reverse transcribed in a total volume of 10 µl containing 200 ng of random hexamers, 20 U of RNAguard RNase inhibitor, 1x first strand buffer, 10 mM dithiothreitol, 0.5 mM dNTP mix, and 100 U of Superscript II RNase H-Reverse Transcriptase. At least six concentrations of the A₁ receptor RNA standard, ranging from 1 x 10³ to 1 x 10⁶ copies/µl, and a sample containing DNase I-treated RNase-free water in place of the template, were reverse-transcribed simultaneously. The reverse-transcribed RNA standards were used to construct the standard curve for the real-time RT-PCR assay, and the sample containing sterile water was used as the template for the negative control.

PCR. Each assay consisted of six standard curve samples, a negative control sample, and unknown samples. All reactions were performed in triplicate. The PCR reaction mixture (25 µl) consisted of 1x TaqMan buffer A; 200 µM of each dATP, dCTP, and dGTP; 400 µM dUTP; 0.01 U/µl AmpErase uracil-N-glycosylase; and 0.025 U/µl AmpliTaq Gold DNA polymerase from the TaqMan PCR core kit (Applied Biosystems). The reaction mixture also contained 0.5 µl of tissue cDNA, standard cDNA or negative control, 100 nM probe, 300 nM each of the forward and reverse primers, and 7.5 mM MgCl₂. The reaction was performed using the ABI Prism 7700 Sequence Detector (Applied Biosystems) with the following cycling parameters: 2-min hold at 50°C for uracil-N-glycosylase incubation, 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 1-min anneal/extend period at 60°C.

Data Collection and Analysis. Data were collected during each PCR cycle and analyzed using the Sequence Detection Software (version 1.6.3; Applied Biosystems). An amplification plot showing normalized reporter emissions versus cycle number was generated (Fig. 8B). The threshold cycle (C_T), the cycle where an increase in fluorescence is associated with exponential growth, was determined by the software using the fluorescence emitted during the first 15 cycles. A standard curve of C_T versus Log (initial A₁ receptor standard concentrations) was generated (Fig. 8C). The initial concentration of each unknown sample was determined by interpolation using the C_T value determined by the assay. The correlation coefficient of each standard curve was >0.95, and the C_T of the no template controls exceeded 40 cycles in every assay, indicating the absence of DNA contamination. Results were expressed as copies of mRNA per micrograms of total RNA. Statistical significance was determined using GraphPad Prism and the two-tailed unpaired Student's t test, where P < 0.05 was considered significant.

View larger version (47K): [in this window] [in a new window]   Fig. 8. Distribution and abundance of A1 receptor mRNA in the rat stomach. A, results of RT-PCR performed using primers that span the entire coding region of the A1 receptor gene. A single amplicon was generated using cDNA from the fundus (lane 2), corpus (lane 3), antrum (lane 4), mucosa (lane 5), and striatum (positive control; lane 1) as the template. Ladder, 100-bp ladder. B, amplification plots generated by the A1 receptor real-time RT-PCR assay using various concentrations of A1 receptor RNA as the standard. The amplification plots (from left to right) were generated using 1 x 109, 1 x 108, 1 x 107, 1 x 106, 1 x 105, 1 x 104, and 1 x 103 copies of A1 receptor RNA. C, corresponding standard curve generated by the amplification plots shown in B. Correlation coefficient is 0.99. D, A1 receptor gene expression levels measured in various regions of the rat stomach using real-time RT-PCR. Each bar represents the mean ± S.E.M. of at least four animals. A1 receptor gene expression was also quantified in the striatum and found to contain 850 ± 55 thousand copies of A1 receptor mRNA/µg total RNA.

	Results

Top Abstract Materials and Methods Results Discussion References

 
Effect of Adenosine Agonists on IRG Release. To examine the adenosine receptor(s) involved in the regulation of IRG release, the effect of the A₁-(CPA), A_2A-(CGS 21680), and A₃ (IB-MECA)-selective agonists was examined because specific agonists for individual receptor subtypes are unavailable. Basal IRG release (periods 1–3) was shown to remain relatively constant in experiments examining the effect of 0.1 µM CPA (194 ± 42 to 194 ± 47 pg/min), 0.1 µM CGS 21680 (123 ± 21 to 124 ± 19 pg/min), and 0.1 µM IB-MECA (185 ± 24 to 201 ± 24 pg/min) on IRG release (Fig. 2). Figure 2A demonstrates that the administration of 0.1 µM CPA caused a significant inhibition of IRG release starting at period 5 and continued throughout the drug perfusion period. Upon the cessation of CPA perfusion, IRG release returned to basal levels. The same concentration of CGS 21680 and IB-MECA did not alter basal IRG release (Fig. 2, B and C). The effect of various concentrations of these analogs on IRG release was also examined. The results are expressed as percentage of inhibition and summarized in Fig. 3. CPA was shown to suppress IRG release concentration dependently. CPA caused significant inhibition of IRG release starting at a concentration of 0.001 µM (12 ± 5%), and maximum suppression of IRG release was reached at 1 µM (43 ± 5%). The empirical EC₅₀ of CPA in inhibiting IRG release was estimated to be 0.067 µM, with a 95% confidence interval between 0.014 and 0.325 µM. CGS 21680 (0.001–0.1 µM) did not alter IRG release. However, significant inhibition (14 ± 5%) was observed when 1 µM CGS21680was perfused into the stomach. All concentrations of IB-MECA examined (0.001–1 µM) did not alter IRG release.

View larger version (21K): [in this window] [in a new window]   Fig. 2. Effect of CPA, CGS 21680, and IB-MECA on basal IRG release. Results are expressed as IRG release (percentage) as described under Materials and Methods. Each column represents the mean ± S.E.M.; n 5. *, P < 0.05 compared with period 3 of respective experiments using repeated measures analysis of variance followed by Dunnett's multiple comparison test.

View larger version (11K): [in this window] [in a new window]   Fig. 3. Effect of various concentrations of CPA, CGS 21680, and IB-MECA on IRG release. Results are expressed as percentage of inhibition and calculated as described under Materials and Methods. Each column represents the mean ± S.E.M. of at least five experiments. *, P < 0.05 and ***, P < 0.001 compared with the mean IRG release (picograms per minute) in the presence of drug during periods 4 to 7 with that of periods 1 to 3 (basal release).

Effect of DPCPX and DMPX on IRG Release. We have demonstrated previously that the endogenous compound adenosine also inhibited IRG release concentration dependently (Kwok et al., 1990). To test whether this inhibitory action is mediated by A₁ receptors, the effect of the antagonists DPCPX (A₁-selective) and DMPX (A₂-selective) on basal and adenosine-stimulated IRG release was examined. When DPCPX or DMPX was perfused alone into the stomach for 20 min (periods 4–7), no significant changes in IRG release were apparent; the percentage of change of IRG in the presence of DPCPX or DMPX alone were 7 ± 4% (n = 6) and 0 ± 4% (n = 6), respectively. Figure 4A shows that adenosine (10 µM) significantly inhibited IRG release when it was administered alone for 15 min. To test the effect of DPCPX and DMPX on adenosine-induced IRG release, the antagonist was perfused 5 min before the concomitant perfusion of both adenosine and antagonist for 15 min. In the presence of DPCPX, the inhibitory effect of adenosine was completely abolished (Fig. 4B). Adenosine-induced inhibition of IRG release was not altered by the concomitant perfusion with DMPX (Fig. 4C).

View larger version (20K): [in this window] [in a new window]   Fig. 4. Effect of DPCPX and DMPX on adenosine-induced inhibition of IRG release. A, effect of adenosine (ADO) on basal IRG release. B, effect of DPCPX on ADO-induced inhibition of IRG release. C, effect of DMPX on ADO-induced inhibition of IRG release. Each column represents the mean ± S.E.M.; n 5; *, P < 0.05 and ***, P < 0.001 compared with period 3 of respective experiments.

Cellular Localization and Distribution of A₁R-IR. Because the results of the perfusion studies suggest that the A₁ receptor is involved in the inhibition of IRG release, immunohistochemistry experiments were performed to determine the cellular localization and distribution of A₁ receptors in the rat stomach. The distribution of A₁R-IR was similar in the gastric antrum and corpus (Fig. 5). Abundant A₁R-IR was observed along the basal region of the antral mucosa (Fig. 5A). When viewed under high magnification, A₁R-IR was found to be intense and punctuate (Fig. 5B). A₁R-IR was dispersed uniformly throughout the corporeal mucosa, except at the tips of the mucosa (Fig. 5, C and D). In both the corpus and antrum, A₁R-IR was also observed on cell bodies and nerve fibers of the myenteric plexus, nerve fibers of the circular muscle layer, longitudinal muscle layer, muscularis mucosae and submucosal plexus, and blood vessels (Fig. 5, E and F). No A₁R-IR was observed when tissue sections were incubated with the immunoneutralized A₁ receptor antibody (Fig. 5G) or with the Cy3-conjugated secondary antibody alone (Fig. 5H).

To examine whether A₁R-IR is localized on gastrin-secreting G-cells, double staining experiments were performed. Abundant gastrin-IR was observed along the base of the antral mucosa (Fig. 6B). Double staining revealed that all gastrin-IR cells examined contained A₁R-IR (Fig. 6C). In addition, A₁R-IR (Fig. 6A) seemed most intense in mucosal cells expressing gastrin-IR. Although all gastrin-IR cells expressed A₁R-IR, approximately 28 ± 2% of A₁R-IR cells of the antral mucosa expressed gastrin-IR.

View larger version (34K): [in this window] [in a new window]   Fig. 6. Double staining of A1R-IR with gastrin-IR and somatostatin-IR in the rat stomach. A to C, colocalization of A1R-IR (A) with gastrin-IR (B) in the antral mucosa. D to F, colocalization of A1R-IR (D) and somatostatin-IR (E) in the antral mucosa. G to I, colocalization of A1R-IR (G) and somatostatin-IR (H) in the corporeal mucosa. J to L, high-magnification image showing the colocalization of A1R-IR (J) and somatostatin-IR (K) in a single cell of the corporeal mucosa (L). Cells expressing both A1R-IR (red) and gastric peptide-IR (green) are indicated by arrows and look yellow in the merged panel (C, F, I, and L). A1R-IR cells that do not contain gastrin-IR or somatostatin-IR are indicated by arrowheads (z-step, 1.0 µm and scale bar, 25 µm for all images).

To determine whether A₁R-IR was also localized on somatostatin-secreting D-cells of the antral and corporeal mucosa, double staining experiments were also performed with antibody against somatostatin. Intense and abundant somatostatin-IR was observed in the antral and corporeal mucosa, whereas sparse somatostatin-IR was observed throughout the rest of the tissue. Double staining with A₁R-IR revealed that some somatostatin-IR cells of the antral and corporeal mucosa also expressed A₁R-IR (Fig. 6, D–I). In somatostatin-IR cells of the corpus, A₁R-IR seemed most intense at the end of the cell processes (Fig. 6, G–I and J–L). Quantification of A₁R-IR revealed that all somatostatin-IR cells of the corporeal mucosa expressed A₁R-IR, whereas approximately 24 ± 2% of somatostatin-IR cells of the antrum expressed A₁R-IR. The present study also demonstrates that 8.5 ± 0.6% of A₁R-IR cells of the antral mucosa expressed somatostatin-IR. A₁R-IR was shown not to colocalize with somatostatin-IR in other regions of the antrum and corpus, including the myenteric plexus and muscle layers.

Double staining experiments were also performed to localize A₁R-IR in relation to the immunoreactivity of H⁺K⁺-ATPase (parietal cell marker), VWF (endothelial cell marker), and PGP 9.5 (neuronal marker). Results show that A₁R-IR was not colocalized with H⁺K⁺-ATPase -IR (Fig. 7, A–C). However, A₁R-IR was colocalized with VWF-IR and PGP 9.5-IR throughout the corpus and antrum (Fig. 7, D–I). A₁R-IR was expressed with VWF-IR in the muscle layers, myenteric and submucosal plexi (Fig. 7, D–F), and with PGP 9.5-IR 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. 7, G–I).

View larger version (51K): [in this window] [in a new window]   Fig. 7. Double staining of A1R-IR with H+K+-ATPase , VWF, and PGP 9.5-IR in the rat corpus. A to C, A1R-IR (A) with H+K+-ATPase -IR (B) in the corporeal mucosa. D to F, colocalization of A1R-IR (D) and VWF-IR (E) on a blood vessel in the circular muscle. G to I, colocalization of A1R-IR (G) with PGP 9.5-IR (H) on nerve fibers and cell bodies of the myenteric plexus (arrowhead) and nerve fibers of the circular muscle (arrows). Cells expressing both immunoreactivities look yellow in the merged panels (F and I) (z-step, 1.0 µm and scale bar, 25 µm for all images).

Regional Distribution, Structure, and Abundance of Adenosine A₁ Receptor mRNA. The expression of A₁ receptor mRNA was detected in all gastric regions examined, including the fundus, corpus, antrum, and mucosa. Only one RT-PCR amplicon of the expected size (1086 bp) was generated for all tissues, including the striatum, which was used as the positive control (Fig. 8A). Cloning and sequencing of the mucosal RT-PCR amplicon demonstrated that the coding region of the gastric mucosal A₁ receptor (submitted to GenBank; accession no. AF042079 [GenBank] ) was identical to the published sequence in the rat brain (Mahan et al., 1991) (accession no. M64299 [GenBank] ).

The real-time RT-PCR assay developed to quantify A₁ receptor gene expression in gastric tissues was shown to measure a 7 log range of A₁ receptor RNA concentrations. A representative amplification plot generated by the A₁ receptor real-time RT-PCR assay is shown on Fig. 8B. When a high concentration of A₁ receptor RNA standard (1 x 10⁹ copies/µl) was used as the template, the reaction was affected by limiting reagents by cycle 22 and entered the plateau phase at cycle 26 (Fig. 8B). The standard curve generated by plotting the threshold cycle (C_T) against the Log(A₁ receptor RNA standard concentration) is shown to be linear with an R value of 0.99 (Fig. 8C). Quantification of A₁ receptor gene expression using this assay demonstrated that the A₁ receptor mRNA level was highest in the corporeal muscle and lowest in the corporeal mucosa (Fig. 8D). A₁ receptor mRNA expression did not differ significantly among the corpus, antrum, and whole stomach mucosa. The fundic A₁ mRNA level was lower than those in the corpus, but not different from those in the antrum and mucosa. The striatal A₁ receptor mRNA level was at least 2-fold greater than levels measured in any gastric region examined. The striatum contained approximately 8.5 x 10⁵ copies of A₁ receptor mRNA/µg total RNA, whereas the highest level measured in the stomach (corpus muscle) contained approximately 4.6 x 10⁵ copies of A₁ receptor mRNA/µg total RNA.

	Discussion

Top Abstract Materials and Methods Results Discussion References

 
Adenosine has been shown to regulate gastric acid secretion and to protect the stomach against ulcer formation. In rats, adenosine inhibits gastric acid secretion, but it does not act directly on the parietal cells to elicit this effect. Our laboratory has demonstrated that adenosine may mediate this inhibitory action by suppressing the release of gastrin (Kwok et al., 1990). Results of the present study show that CPA, the A₁ receptor-selective agonist (Williams et al., 1986), significantly inhibited IRG release starting at a concentration of 0.001 µM, and maximally inhibited IRG release at 1 µM. Although the A_2A receptor-selective agonist CGS 21680 did not alter IRG release at lower concentrations, this compound did inhibit IRG release significantly at 1 µM. The percentage of inhibition of IRG release elicited by 1 µM CGS 21680 was similar to that produced by 0.001 µM CPA. Thus, the inhibitory effect of CGS 21680 is considerably weaker than that of CPA. The A₃ receptor-selective agonist IB-MECA did not alter IRG release. The potent inhibitory effect of CPA on IRG release may, therefore, suggest that the A₁ receptor subtype is likely involved in modulating IRG release. This proposal was further supported by the observation that the inhibitory effect of the endogenous compound adenosine on IRG release was completely abolished by the A₁ receptor-selective antagonist DPCPX (Lohse et al., 1987) but not by the A₂ receptor-selective antagonist DMPX (Seale et al., 1988). DPCPX exhibits a 700-fold preference for the A₁ receptor over the A₂ receptor (Bruns et al., 1987), whereas DMPX exhibits a 4-fold preference for the A₂ receptor over the A₁ receptor (Seale et al., 1988). At 1 µM, DPCPX and DMPX have been shown to completely abolish A₁ and A_2A receptor-mediated actions, respectively (Lohse et al., 1987; Seale et al., 1988). The present study also examined the possible site of action of the adenosine A₁ receptor-mediated inhibitory action on IRG release by examining the immunohistochemical localization of the A₁ receptor.

A₁R-IR was expressed on all gastrin-IR G-cells examined, suggesting that adenosine may act directly on G-cells to inhibit IRG release. This proposal agrees well with studies showing that adenosine inhibited the release of gastrin in rat preparations containing G-cells (DeSchryver-Kecskemeti et al., 1981; Harty and Franklin, 1984) and in primary cultures of canine G-cells (Schepp et al., 1990). In these studies, the adenosine-induced inhibition was sensitive to blockade by theophylline and 8-phenyltheophylline (DeSchryver-Kecskemeti et al., 1981; Schepp et al., 1990). Xanthine derivatives such as theophylline, 8-phenyltheophylline, and caffeine are nonselective antagonists of the adenosine receptors and may block adenosine receptor-mediated responses (Ralevic and Burnstock, 1998). The present study further shows that the inhibitory effect of adenosine can be abolished by the A₁ receptor-selective antagonist DPCPX, thus confirming the involvement of extracellular adenosine receptors.

A₁R-IR was colocalized with somatostatin-IR in cells of the antral and corporeal mucosa, suggesting that A₁ receptors may be expressed on D-cells. Somatostatin was shown to inhibit IRG release (Saffouri et al., 1980). Thus, adenosine may inhibit IRG release by modulating the release of somatostatin. Adenosine has been demonstrated to regulate somatostatin-like immunoreactivity (SLI) release in the isolated vascularly perfused rat stomach (Kwok et al., 1990). Activation of adenosine A₁ and A_2A receptors may lead to inhibition and stimulation of SLI release, respectively (Yip and Kwok, 2004). However, it is doubtful that adenosine inhibits IRG release indirectly by modulating SLI release. Activation of A₁ receptors inhibits SLI release, which would lead to an increase, rather than a decrease in IRG release. Activation of A_2A receptors stimulates SLI release, which may subsequently inhibit IRG release. However, the adenosine-induced inhibition of IRG release was completely abolished by DPCPX and not altered by DMPX, suggesting that A_2A receptor-induced SLI release is not involved in suppressing IRG release. Previous studies have shown that adenosine A_2A receptors are expressed on D-cells, indicating that adenosine may act directly on these cells to stimulate somatostatin release (Yip and Kwok, 2004). The colocalization of A₁R-IR with somatostatin-IR observed in the present study suggests that adenosine may also act directly on D-cells to inhibit somatostatin release. Adenosine may, therefore, play a dual role in the regulation of somatostatin release. It may inhibit and stimulate somatostatin release via activation of A₁ and A_2A receptors on D-cells, respectively.

A₁R-IR was not colocalized with H⁺K⁺-ATPase -IR, indicating the absence of A₁ receptors on parietal cells. The lack of A₁R-IR on parietal cells suggests that adenosine does not act directly on these cells to inhibit acid secretion. This finding confirms previous studies demonstrating the inability of N⁶(–)-phenylisopropyladenosine to alter carbachol- and histamine-stimulated acid secretion in rat parietal cell preparations (Puurunen et al., 1987).

A₁R-IR was also expressed on vasculature and nerve fibers throughout the stomach, as indicated by its colocalization with VWF-IR and PGP 9.5-IR, respectively. The vascular localization of these receptors fits well with previously observed A₁ receptor-mediated vascular actions (Tabrizchi and Bedi, 2001). Abundant A₁R-IR was present on nerve fibers of the myenteric and submucosal plexi. These localizations are consistent with studies demonstrating the involvement of A₁ receptors in the inhibition of neurotransmitter release from myenteric (Palmer et al., 1987) and submucosal (Barajas-Lopez et al., 1991) neurons. A₁ receptors have been implicated in the inhibition of acetylcholine and noradrenaline release (Barajas-Lopez et al., 1991; Nitahara et al., 1995; Ribeiro et al., 1996). Because both neurotransmitters can alter IRG release (Koop et al., 1982; Koop et al., 1983), it is conceivable that the effect of adenosine on IRG release is secondary to its effect on adrenergic and cholinergic neurotransmission. Although this possibility was not determined, the localization of A₁R-IR on all gastrin-IR cells examined suggests that adenosine may inhibit IRG release, at least in part, by acting directly on G-cells.

The structure of the gastric A₁ receptor was also examined in the present study. Two distinct A₁ receptor mRNA transcripts have been detected by Northern blot analysis in the rat stomach (Mahan et al., 1991; Reppert et al., 1991). However, when RT-PCR experiments were performed using primers that span the entire coding region of the A₁ receptor gene, only one amplicon was produced. Thus, alternative splicing is unlikely to occur within the coding region of the gastric A₁ receptor gene. In humans, two promoters drive the synthesis of two distinct A₁ receptor transcripts that differ in the 5'-untranslated region (Ren and Stiles, 1995). Thus, the two A₁ receptor transcripts previously detected in the rat stomach may be generated by separate promoters or alternative splicing of the 5'-untranslated region. The present cloning and sequencing experiments reveal that the coding region of the gastric A₁ receptor was identical to the rat brain sequence (Mahan et al., 1991), indicating that only one form of the A₁ receptor, identical to the rat brain receptor, exists in the stomach.

Results also show that A₁ receptor gene expression was expressed in all functionally and morphologically distinct regions of the stomach. Real-time RT-PCR was used to measure the level of A₁ receptor gene expression in various gastric regions. This assay was shown to accurately quantify over a 7 log range of concentrations and less than 500 copies of A₁ receptor RNA/µg total tissue RNA (25 copies/reaction tube). In agreement with previous studies (Dixon et al., 1996), results demonstrate that A₁ receptor gene expression was higher in the striatum than the stomach. The striatal A₁ receptor mRNA level was at least 2-fold higher than any gastric region examined. Among the gastric tissues, the A₁ receptor mRNA level was lowest in the corporeal mucosa and highest in the corporeal muscle. These findings correspond well with results of the immunohistochemistry studies. In the corpus, A₁R-IR was found on some mucosal cells, but the majority was localized on nerve fibers and vasculature of the muscle layers and myenteric plexus. The abundant A₁R-IR observed at the base of the antral mucosa and the moderate A₁R-IR observed throughout the corporeal mucosa is also consistent with the higher level of A₁ receptor mRNA measured in the whole stomach mucosa compared with the corporeal mucosa.

In conclusion, the results of this study suggest that, in the rat stomach, adenosine may mediate its inhibitory action on IRG release, at least in part, by acting on A₁ receptors of G-cells. The inhibition of IRG release may subsequently lead to a decrease in gastric acid secretion. The gastric A₁ receptor was found to be structurally identical to that in the rat brain and was expressed on various cells throughout the stomach, including the G-cells. The gene expression of the A₁ receptor was extremely low in all gastric tissues examined, although it was easily quantified by the sensitive real-time RT-PCR assay developed in the present study.

	Footnotes

 
This work was supported by the Canadian Apoptosis Research Foundation Society, Canada Foundation for Innovation, Wah Sheung Fund, and the former British Columbia Health Research Foundation. L.Y. was supported by the Cordula and Gunter Paetzold Fellowship and the University of British Columbia Graduate Fellowship. A portion of this work was included in Linda Yip's Ph.D. dissertation entitled Adenosine A₁ and A_2A Receptors in the Rat Stomach: Biological Actions, Cellular Localization, Structure, and Gene Expression. Citation of meeting abstracts where part of this work was previously presented: Yip L and Kwok YN (2002) Gastric A₁ and A_2A receptors: cellular localization, gene sequence and gene expression levels. Drug Dev Res 56:551 and Yip L, Leung CH, and Kwok YN (2003) Cellular localization of adenosine A₁ and A_2A receptors in the rat stomach. FASEB J 17:A40.

DOI: 10.1124/jpet.104.066654.

ABBREVIATIONS: IRG, immunoreactive gastrin; RT-PCR, reverse transcription-polymerase chain reaction; BSA, bovine serum albumin; RIA, radioimmunoassay; CPA, N⁶-cyclopentyladenosine; CGS 21680, 2-p-(2-carboxyethyl)phenethylamino-5'N-ethylcarboxamidoadenosine; IB-MECA, 1-deoxy-1-[6-[[(3-iodophenyl)methyl]amino]-9H-purin-9-yl]-N-methyl--D-ribofuranuronamide; DPCPX, 8-cyclopentyl-1,3-dipropylxanthine; DMPX, 3,7-dimethyl-1-propargylxanthine; A₁R-IR, A₁ receptor-immunoreactivity; TBS, Tris-buffered saline; TBST, Tris-buffered saline/Tween 20; PBS, phosphate-buffered saline; IR, immunoreactivity; PGP 9.5, protein gene product 9.5; VWF, von Willebrand's factor; bp, base pair(s); SLI, somatostatin-like immunoreactivity.

Address correspondence to: Dr. Yin Nam Kwok, Department of Physiology, University of British Columbia, 2146 Health Sciences Mall, Vancouver, BC, Canada V6T 1Z3. E-mail: kynkwok{at}interchange.ubc.ca

	References

Top Abstract Materials and Methods Results Discussion References

 

Barajas-Lopez C, Surprenant A, and North RA (1991) Adenosine A1 and A2 receptors mediate presynaptic inhibition and postsynaptic excitation in guinea pig submucosal neurons. J Pharmacol Exp Ther 258: 490–495.[Abstract/Free Full Text]

Bruns RF, Fergus JH, Badger EW, Bristol JA, Santay LA, Hartman JD, Hays SJ, and Huang CC (1987) Binding of the A1-selective adenosine antagonist 8-cyclopentyl-1,3-dipropylxanthine to rat brain membranes. Naunyn-Schmiedeberg's Arch Pharmacol 335: 59–63.[CrossRef][Medline]

Carruthers AM, Sellers LA, Jenkins DW, Jarvie EM, Feniuk W, and Humphrey PP (2001) Adenosine A(1) receptor-mediated inhibition of protein kinase A-induced calcitonin gene-related peptide release from rat trigeminal neurons. Mol Pharmacol 59: 1533–1541.[Abstract/Free Full Text]

DeSchryver-Kecskemeti K, Greider MH, Rieders ER, Komyati SE, and McGuigan JE (1981) In vitro gastrin secretion by rat antrum: effects of neurotransmitter agonists, antagonists and modulators of secretion. Lab Investig 44: 158–163.[Medline]

Dixon AK, Gubitz AK, Sirinathsinghji DJ, Richardson PJ, and Freeman TC (1996) Tissue distribution of adenosine receptor mRNAs in the rat. Br J Pharmacol 118: 1461–1468.[Medline]

Fredholm BB, AP IJ, Jacobson KA, Klotz KN, and Linden J (2001) International Union of Pharmacology. XXV. Nomenclature and classification of adenosine receptors. Pharmacol Rev 53: 527–552.[Abstract/Free Full Text]

Fujimiya M and Kwok YN (1997) Effect of carbachol on vascular and luminal release of immunoreactive gastrin from isolated perfused rat duodenum. Dig Dis Sci 42: 634–639.[CrossRef][Medline]

Geiger JD and Glavin GB (1985) Adenosine receptor activation in brain reduces stress-induced ulcer formation. Eur J Pharmacol 115: 185–190.[CrossRef][Medline]

Gerber JG, Nies AS, and Payne NA (1985) Adenosine receptors on canine parietal cells modulate gastric acid secretion to histamine. J Pharmacol Exp Ther 233: 623–627.[Abstract/Free Full Text]

Gerber JG and Payne NA (1988) Endogenous adenosine modulates gastric acid secretion to histamine in canine parietal cells. J Pharmacol Exp Ther 244: 190–194.[Abstract/Free Full Text]

Glavin GB, Westerberg VS, and Geiger JD (1987) Modulation of gastric acid secretion by adenosine in conscious rats. Can J Physiol Pharmacol 65: 1182–1185.[Medline]

Harty RF and Franklin PA (1984) Effects of exogenous and endogenous adenosine on gastrin release from rat antral mucosa. Gastroenterology 86: 1107.

Heldsinger AA, Vinik AI, and Fox IH (1986) Inhibition of guinea-pig oxyntic cell function by adenosine and prostaglandins. J Pharmacol Exp Ther 237: 351–356.[Abstract/Free Full Text]

Jaffe B and Walsh JH (1978) Gastrin and related peptides, in Methods of Hormone Radioimmunoassay (Jaffe BM and Behrman HR eds) pp 455–477, Academic Press, New York.

Koop H, Behrens I, Bothe E, Koschwitz H, McIntosh CH, Pederson RA, Arnold R, and Creutzfeldt W (1983) Adrenergic control of rat gastric somatostatin and gastrin release. Scand J Gastroenterol 18: 65–71.[Medline]

Koop H, Behrens I, Bothe E, McIntosh CH, Pederson RA, Arnold R, and Creutzfeldt W (1982) Adrenergic and cholinergic interactions in rat gastric somatostatin and gastrin release. Digestion 25: 96–102.[Medline]

Kwok YN, McIntosh C, and Brown J (1990) Augmentation of release of gastric somatostatin-like immunoreactivity by adenosine, adenosine triphosphate and their analogs. J Pharmacol Exp Ther 255: 781–788.[Abstract/Free Full Text]

Lohse MJ, Klotz KN, Lindenborn-Fotinos J, Reddington M, Schwabe U, and Olsson RA (1987) 8-Cyclopentyl-1,3-dipropylxanthine (DPCPX)–a selective high affinity antagonist radioligand for A1 adenosine receptors. Naunyn-Schmiedeberg's Arch Pharmacol 336: 204–210.[CrossRef][Medline]

Mahan LC, McVittie LD, Smyk-Randall EM, Nakata H, Monsma FJ Jr, Gerfen CR, and Sibley DR (1991) Cloning and expression of an A1 adenosine receptor from rat brain. Mol Pharmacol 40: 1–7.[Abstract]

Nitahara K, Kittel A, Liang SD, and Vizi ES (1995) A1-receptor-mediated effect of adenosine on the release of acetylcholine from the myenteric plexus: role and localization of ecto-ATPase and 5'-nucleotidase. Neuroscience 67: 159–168.[CrossRef][Medline]

Palmer JM, Wood JD, and Zafirov DH (1987) Purinergic inhibition in the small intestinal myenteric plexus of the guinea-pig. J Physiol (Lond) 387: 357–369.[Abstract/Free Full Text]

Pederson RA, Kwok YN, Buchan AM, McIntosh CH, and Brown JC (1984) Gastrin release from isolated perfused rat stomach after vagotomy. Am J Physiol 247: G248–G252.[Medline]

Puurunen J, Ruoff HJ, and Schwabe U (1987) Lack of direct effect of adenosine on the parietal cell function in the rat. Pharmacol Toxicol 60: 315–317.[Medline]

Ralevic V and Burnstock G (1998) Receptors for purines and pyrimidines. Pharmacol Rev 50: 413–492.[Abstract/Free Full Text]

Ren H and Stiles GL (1995) Separate promoters in the human A1 adenosine receptor gene direct the synthesis of distinct messenger RNAs that regulate receptor abundance. Mol Pharmacol 48: 975–980.[Abstract]

Reppert SM, Weaver DR, Stehle JH, and Rivkees SA (1991) Molecular cloning and characterization of a rat A1-adenosine receptor that is widely expressed in brain and spinal cord. Mol Endocrinol 5: 1037–1048.[Abstract/Free Full Text]

Ribeiro JA, Cunha RA, Correia-de-Sa P, and Sebastiao AM (1996) Purinergic regulation of acetylcholine release. Prog Brain Res 109: 231–241.[Medline]

Saffouri B, Weir GC, Bitar KN, and Makhlouf GM (1980) Gastrin and somatostatin secretion by perfused rat stomach: functional linkage of antral peptides. Am J Physiol 238: G495–G501.

Sajjadi FG, Boyle DL, Domingo RC, and Firestein GS (1996) cDNA cloning and characterization of A3i, an alternatively spliced rat A3 adenosine receptor variant. FEBS Lett 382: 125–129.[CrossRef][Medline]

Sambrook J, Fritsch EF, and Maniatis T (1989) Detection and analysis of proteins expressed from cloned genes, in Molecular Cloning: A Laboratory Manual (Nolan C ed) pp 18.11–18.88, Cold Spring Harbor Laboratory Press, Plainview, NY.

Schepp W, Soll AH, and Walsh JH (1990) Dual modulation by adenosine of gastrin release from canine G-cells in primary culture. Am J Physiol 259: G556–G563.

Seale TW, Abla KA, Shamim MT, Carney JM, and Daly JW (1988) 3,7-Dimethyl-1-propargylxanthine: a potent and selective in vivo antagonist of adenosine analogs. Life Sci 43: 1671–1684.[CrossRef][Medline]

Tabrizchi R and Bedi S (2001) Pharmacology of adenosine receptors in the vasculature. Pharmacol Ther 91: 133–147.[CrossRef][Medline]

Westerberg VS and Geiger JD (1987) Central effects of adenosine analogs on stress-induced gastric ulcer formation. Life Sci 41: 2201–2205.[CrossRef][Medline]

Westerberg VS and Geiger JD (1989) Adenosine analogs inhibit gastric acid secretion. Eur J Pharmacol 160: 275–281.[CrossRef][Medline]

Williams M, Braunwalder A, and Erickson TJ (1986) Evaluation of the binding of the A-1 selective adenosine radioligand, cyclopentyladenosine (CPA), to rat brain tissue. Naunyn-Schmiedeberg's Arch Pharmacol 332: 179–183.[CrossRef][Medline]

Yip L and Kwok YN (2004) Role of adenosine A2A receptor in the regulation of gastric somatostatin release. J Pharmacol Exp Ther 309: 804–815.[Abstract/Free Full Text]

Yip L, Kwok YN, and Buchan AM (2003) Cellular localization and distribution of neurokinin-1 receptors in the rat stomach. Auton Neurosci 104: 95–108.[CrossRef][Medline]

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