Demonstration and immunolocalization of ATP diphosphohydrolase in the pig digestive system

Département de Biologie, Faculté des Sciences, Université de Sherbrooke, Sherbrooke, Québec, Canada J1K 2R1

	ABSTRACT

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Two isoforms of ATP diphosphohydrolase (ATPDase; EC 3.6.1.5) have been previously characterized, purified, and identified. This enzyme is an ectonucleotidase that catalyzes the sequential release of - and -phosphate groups of triphospho- and diphosphonucleosides. One of its putative roles is to modulate the extracellular concentrations of purines in different physiological systems. The purpose of this study was to define, identify, and localize these two isoforms of ATPDase in the pig digestive system. ATPDase activity was measured in pig stomach, duodenum, pancreas, and parotid gland. Enzyme assays, electrophoretograms, and Western blots with a polyclonal antibody that recognizes both isoforms demonstrate the presence of ATPDase in these organs. Immunolocalization showed intense reactions with gastric glands (parietal and chief cells), intestine (columnar epithelial cells), parotid gland, and pancreas. Smooth muscle cells all along the digestive tract were also highly reactive. Considering the variety of purinoceptors associated with the digestive system, the ATPDase is strategically positioned to modulate purine-mediated actions such as electrolyte secretion, glandular secretion, smooth muscle contraction, and blood flow.

apyrase; ecto-ATPase; stomach; intestine; parotid; pancreas

	INTRODUCTION

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IN THE PAST DECADE, evidence has grown in support of the concept that extracellular ATP and its dephosphorylated metabolites are involved in cell signaling in the different physiological systems of vertebrates (20). ATP is released from cells by exocytotic and nonexocytotic mechanisms; the latter are still undefined. Extracellular concentrations of ATP and its metabolites appear to be modulated primarily by ectonucleotidases, namely, an ATP diphosphohydrolase (ATPDase), an ecto-ATPase, and the 5'-nucleotidase. The 5'-nucleotidase that converts 5'-nucleotides to nucleosides has been well described (38); however, this is not the case for the ectoenzymes responsible for the hydrolysis of ATP to ADP and ADP to AMP. Indeed, until recently, it was widely believed that distinct enzymes were involved in the sequential hydrolysis of the - and -phosphate residues of ATP, i.e., an ecto-ATPase and an ecto-ADPase. This notion was revised after the finding of an ecto-ATPDase in many tissues. The latter enzyme, originally found in pig pancreas (19), has been purified, characterized, and identified (8, 32-33). Its encoding gene has been sequenced and corresponds to the gene of CD39, a marker of activated lymphocytes (14). In addition, other ectonucleotidases (7, 16, 17) that convert ATP to ADP have been described (see Refs. 4, 18, and 28 for review). Several lines of evidence have indicated that extracellular purines play primordial roles in the gastrointestinal tract and some of its associated organs, such as the salivary glands and the pancreas. These actions are mediated by purinergic receptors and modulated by still undefined nucleotidase activities. Extracellular nucleotides and their dephosphorylated derivatives exert major physiological effects on the digestive system, and ATPDase could potentially modulate the concentrations of these nucleotides. It therefore appeared important to determine the localization of ATPDase in the gastrointestinal tract. In light of current literature describing the localization and properties of P₁ and P₂ purinoceptors, our analysis provides new insights into the physiological role of this enzyme. More specifically, ecto-ATPDase reveals high levels of both immunoreactivity and enzyme activity in the stomach, intestine, pancreas, and parotid gland.

	MATERIALS AND METHODS

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Materials

ATP, nitro blue tetrazolium (NBT), 5-bromo-4-chloro-3-indolyl phosphate (BCIP), Tris, tetramisole, imidazole, ammonium molybdate, glycerin, EDTA, sucrose, sodium deoxycholate, phenylmethylsulfonyl fluoride (PMSF), malachite green, sodium azide (NaN₃), and mouse monoclonal antibodies to rabbit IgG conjugated to alkaline phosphatase were obtained from Sigma Chemical (St. Louis, MO). ADP, AMP, and Triton X-100 were purchased from Boehringer-Mannheim (Laval, PQ). CaCl₂, MgCl₂, and Tween 20 were obtained from Fisher (Montréal, PQ). Transfer membrane Immobilon-P was obtained from Millipore (Bedford, MA); Bradford reagent, BSA fraction V, SDS, molecular mass standards, and polyacrylamide were obtained from Bio-Rad (Mississauga, ON). All other reagents were of analytical grade.

Isolation of Particulate Fractions

Organs from three young pigs, anesthetized by an intraperitoneal injection of 33% chloral hydrate in saline (1 ml/kg), were frozen in liquid nitrogen. Particulate fractions were prepared as previously described (32). Briefly, tissues were homogenized with a Polytron in 10 vol of 95 mM NaCl, 0.1 mM PMSF, and 45 mM Tris, pH 7.6, at 4°C. The homogenate was filtered through cheesecloth and centrifuged for 15 min at 700 g. The supernatant was centrifuged for 1 h at 100,000 g in an SW41 Beckman rotor. The latter pellet was suspended in a solution of 0.1 mM PMSF and 1 mM NaHCO₃, pH 10.0, with a potter Elvejehm homogenizer, loaded on a 40% (wt/vol) sucrose cushion, and centrifuged for 90 min at 100,000 g in an SW41 Beckman rotor. The fluffy layer was recovered on the cushion, washed twice with 1 mM NaHCO₃ and 0.1 mM PMSF, and suspended in 7.5% glycerin and 5 mM Tris, pH 8.0, at a concentration of 2-5 mg protein/ml. Enzyme activity was measured the same day.

ATPase and ADPase assays

Enzyme assays were carried out at 37°C in 1 ml of solution containing (in mM) 8 CaCl₂, 5 tetramisole, 50 Tris, and 50 imidazole, pH 7.5. The reaction was started by adding 0.2 mM of the substrate (ATP or ADP) and stopped with 0.25 ml of the malachite green reagent. P_i was estimated according to the method of Baykov et al. (2). Controls were run with the protein sample added after the malachite green reagent. Enzyme activity was expressed as nanomoles of P_i released per minute per milligram of protein, which corresponds to milliunits (19). Protein was estimated by the technique of Bradford (5), using BSA as a standard of reference.

Electrophoresis and Immunoblotting Procedures

PAGE was carried out under nondenaturing conditions (32). Nucleotidase activity was detected by incubating the gel for 3 h at 37°C in 10 mM CaCl₂, 100 mM Tris-imidazole, pH 7.5, and 4 mM ATP, ADP, or AMP. Released phosphate forms a white Ca²⁺ precipitate at the reaction site in the gel. Where indicated, proteins were separated by SDS-PAGE in a polyacrylamide gradient (8-13.5%), as described by Sévigny et al. (32). Immunoblotting procedures were carried out as previously reported, using a rabbit antiserum raised against a synthetic polypeptide corresponding to the NH₂-terminal 16-amino acid sequence of the pig pancreas type I ATPDase, at a dilution of 1:10,000 (33). This antibody specifically recognizes both isoforms (type I and II ATPDase) of the enzyme. Preparation and antibody specificity have been previously described (32-33). The secondary antibody was a mouse monoclonal anti-rabbit IgG (1:6,000) conjugated to alkaline phosphatase, detected with NBT/BCIP.

Immunohistochemistry

Freshly dissected tissues were fixed overnight in 2% paraformaldehyde, 0.17% glutaraldehyde, and 4% sucrose, buffered at pH 7.4 with 0.1 M sodium cacodylate buffer. Tissues were dehydrated in graded ethanol solutions and embedded in paraffin. Paraffin sections were cut at 4-µm thickness and mounted on polyionic slides (Superfrost Plus; Fisher). Paraffin was removed with xylene, and sections were rehydrated through graded concentrations of ethanol to water and rinsed in Tris-buffered saline (TBS; 150 mM NaCl, 0.1 M Tris, pH 7.5). Slides were incubated for 10 min in TBS containing 0.1 M glycine and subjected to a pressure cooker heat-induced epitope retrieval procedure by incubating in 1 mM EDTA and 10 mM Tris, pH 8.0, for 9 min (23). After a 10-min wash in TBS at room temperature, nonspecific binding sites were blocked with 1% BSA and 1% fat-free skim milk in TBS for 30 min at room temperature. Sections were incubated overnight at 4°C with the ATPDase antiserum or the preimmune serum (1:100), washed in TBS several times, and incubated with mouse monoclonal anti-rabbit IgG conjugated to alkaline phosphatase at a dilution of 1:100 for 2 h at room temperature. After several washings in TBS, visualization was obtained by the alkaline phosphatase reaction with NBT/BCIP. Sections were mounted in 5% gelatin, 27% glycerin, and 0.1% sodium azide, preheated at 45°C. Photographs were taken under bright-field illumination, with the use of a Zeiss photomicroscope on Kodak T-Max 100 film.

	RESULTS

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Biochemical Data

As shown in Table 1, high levels of ATPase and ADPase activities were found in the homogenates of stomach, duodenum, pancreas, and parotid gland. ATPase and ADPase activities were enriched by about five times in particulate fractions isolated from these organs, with the highest levels in the stomach. Both ATPase and ADPase activities from these organs were sensitive to sodium azide, a well-known inhibitor of ATPDase. From previous studies we know that the only azide-sensitive ADPase activities are attributable to ATPDases (4). The presence of the latter enzyme in these organs was also assessed by looking at electrophoretograms of the particulate fractions separated by PAGE under nondenaturing conditions. As shown in Fig. 1A, electrophoretograms obtained with ATP and ADP as substrates correspond. Migration distances in the polyacrylamide gel were comparable for ATPDases from stomach, duodenum, and parotid gland but differed for the liver enzyme, which was included for comparison. With the pancreas particulate fraction, a barely detectable precipitate could be seen. In contrast, a strong signal was observed with the enriched ATPDase (isoform I) from the zymogen granule membrane fraction, which had a specific activity of 3 U/mg protein. With all of these preparations, there was no detectable reaction product with AMP as the substrate (data not shown). ATPDase identity was confirmed in these different organs by Western immunoblots, using a previously described polyclonal antibody directed against a specific 16-amino acid sequence of the pig enzyme. Good signals were found in stomach and intestine, whereas immunoreactions were much weaker in the parotid gland and pancreas (Fig. 1B).

View this table: [in this window] [in a new window]   Table 1.   Distribution of ATPase and ADPase activities in the digestive system

View larger version (69K): [in this window] [in a new window]   View larger version (38K): [in this window] [in a new window]   Fig. 1.   A: electrophoretograms of nucleotidase activity after PAGE under nondenaturing conditions. A sample of 75 µg of protein of particulate fractions from each tissue was loaded on a 4-7.5% polyacrylamide gradient [13 µg of protein in the case of zymogen granule membrane (ZGM) of pancreas]. ATPase and ADPase activities were localized by incubating the gel for 3 h at 37°C in 10 mM CaCl2, 100 mM Tris-imidazole, pH 7.5, and 4 mM of either ATP or ADP. Liberated Pi forms a white precipitate with Ca2+ at reaction sites. Notice similar migration patterns of ATPase and ADPase activities. B: Western blots of particulate fractions. Samples of 100 µg of protein of particulate fractions from each tissue were separated by SDS-PAGE and immunoblotted as described in MATERIALS AND METHODS. Positive control consisted of a particulate fraction from bovine spleen (30 µg). The sample of zymogen granules, purified as described in Ref. 32, contained 15 µg of protein.

Immunological Localization

Stomach. Immunological localization shows that ATPDase is associated with gastric glands. As shown in Fig. 2B, at high magnification one finds staining of both parietal and chief cell membranes all along the crypts in cardia and fundus regions. Notice, however, that there is much less reactivity at the crypt extremities (Fig. 2A). In chief cells the reaction product was localized on the plasma membrane, whereas in parietal cells the entire cell appeared to be stained (Fig. 2B). Since the latter cells possess a network of canaliculi, which are invaginations of the plasma membrane, the cytoplasmic staining probably reflects the labeling of this network (Fig. 3, A and B). Strong reactions were observed in stomach smooth muscle cells (muscularis externa) (Fig. 3, C and D). In these cells, ATPDase was mainly found on the plasma membrane, with some variability in signal intensity, suggesting a heterogeneity in the distribution of this enzyme among these cells (Fig. 3). Similar reactivity was observed in smooth muscles at the level of the intestine.

View larger version (181K): [in this window] [in a new window]   Fig. 2.   Immunolocalization of ATP diphosphohydrolase (ATPDase) in stomach: cardia region. A: very weak signal on mucus-producing cells (arrowheads) and strong reaction on gastric glands (body and fundus). B: higher magnification shows reaction product concentrated on plasma membrane of chief cells (arrowheads) with strong reactions in parietal cells (arrows). C: control with preimmune serum. Magnification: ×200 (A and C), ×500 (B).

View larger version (220K): [in this window] [in a new window]   Fig. 3.   Immunolocalization of ATPDase in stomach: fundus region and muscularis externa. A: high immunoreactivity of parietal cells. B: higher magnification showing intense signal in parietal cells (arrows): transverse section. C: overview showing exceptionally high intensity of the immunoreaction on smooth muscles (arrow). D: higher magnification showing immunostaining on plasma membrane (arrowheads). Notice variability in the signal. Some cells appear to be devoid of reaction (arrows). Magnification: ×200 (A and C), ×500 (B and D).

Intestine. In the duodenum, ATPDase was found on columnar epithelial cells. Some cells of the lamina propria were also highly reactive (Fig. 4, A, B, and D), including lymphocytes, which appear to migrate into the epithelial layer (Fig. 4C). In the jejunum, ATPDase distribution was essentially similar, although the majority of reticular cells of the lamina propria were also highly reactive (Fig. 4, E and F). In the ileum, the enzyme was found in both columnar and reticular cells, and Peyer's patches showed particularly intense reactions (Fig. 4G).

View larger version (177K): [in this window] [in a new window]   Fig. 4.   Immunolocalization of ATPDase in the intestine. A: duodenum columnar cells are positive (arrowheads) together with some cells in the lamina propria (arrow). B: immunoreactive lymphocytes (arrowheads). C: small lymphocytes described above show high immunoreactivity (arrowheads). Notice different levels of migration through epithelial layer. D: a dense immunoreaction is observed on reticular cells of the duodenum lamina propria (arrow) (transverse section). E: in jejunum, columnar cells are positive (arrowheads), with some immunoreactivity in the lamina propria (arrows). F: the majority of reticular cells are highly reactive (arrow). G: immunolocalization of ATPDase in the ileum. Highly reactive columnar cells (arrowheads), reticular cells (arrows), and Peyer's patch (large arrowhead). Magnification: ×200 (A, E, and G), ×500 (B-D and F).

Pancreas. Pancreas ATPDase immunoreactivity was quite high considering the low specific activity of the enzyme measured in the particulate fraction (see Table 1). Careful examination of the sections revealed that the immunoreaction products were localized on basal and lateral membranes as well as on the apical membrane of acinar cells, whereas the signal over zymogen granules appeared less intense (Fig. 5, A-C). ATPDase was also found on the ductal epithelium, with much less reactivity associated with mucus-producing cells (Fig. 5D).

View larger version (209K): [in this window] [in a new window]   Fig. 5.   Immunolocalization of ATPDase in the pancreas. A: high level of immunoreactivity associated with acinar cells (arrowheads). B: at high magnification, immunoreaction seems to be associated with basal (arrows) and lateral membranes (arrowheads). C: immunoreaction observed on apical membrane (arrowheads). D: control with preimmune serum. E: immunolocalization of ATPDase at the level of ducts. The majority of ductal cells react strongly, but some are devoid of reaction (arrow). Magnification: ×500 (A, D, and E), ×800 (B and C).

Parotid. As shown in Fig. 6, the level of ATPDase was very high in ductal epithelium and blood vessels, in contrast to pancreas acinar cells. Parotid acinar cells were devoid of immunoreactivity, except for a very light signal at the level of the apical membrane, which suggests a shedding of the enzyme in the acinar lumen. Myoepithelial cells, also identified as "basket cells," which are dispersed throughout the tissue, were also positive.

View larger version (165K): [in this window] [in a new window]   Fig. 6.   Immunolocalization of ATPDase in the parotid gland. A: all ducts (arrows) and blood vessels show strong immunoreactivity (arrowheads). B: strong immunoreactivity on duct cells (arrow) with a light signal in acinar lumen (arrowheads). Myoepithelial cells (basket cells) (wavy arrows) are also positive. C: control with preimmune serum. Notice acini (arrowheads) and secretory duct cells (arrow). Magnification: ×125 (A), ×500 (B and C).

	DISCUSSION

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This study reports the immunolocalization of ATPDase in the different cells and tissues of the mammalian digestive system. Intense reactions were found in stomach and intestine, in agreement with enzyme assays. More specifically, ATPDase was found on epithelial cells, smooth muscle cells, reticular cells, and lymphocytes in these organs. Reactivity of the gastric glands was remarkably high in both chief cells and parietal cells. In addition, ATPDase was localized on the ductal systems of glands associated with the digestive tract. Similar to the parenchymal cells of the gastric glands, pancreatic acini were highly reactive, whereas parotid gland acini did not produce significant levels of reaction. This wide distribution of ATPDase, joined to the existence of a variety of purinoceptors, supports the view that this enzyme plays a key role in the regulation of purinergic actions and many vital processes throughout the digestive tract. Obviously, there are still many missing links in this complex puzzle of purine signaling. Perhaps the most crucial questions are the source and the mechanisms of ATP release. There is a lack of knowledge regarding these processes. Nevertheless, we explored the role of ATPDase by looking at purinoceptor distribution and the physiological effects on different cellular systems of the gastrointestinal tract.

Stomach

High levels of ATPDase were measured in the stomach, and immunolocalization indicated that the enzyme is mainly associated with parietal cells, chief cells, and smooth muscle cells. In contrast, ATPDase was barely detectable in mucus-producing cells. Several reports have provided evidence that gastric acid secretion may be modulated by adenosine receptors in rabbit and rat stomach (1, 25, 36) and that it is mediated by an A₂ receptor (1, 25). There is also some evidence for a P_2Y receptor in gastric gland plasma membrane (35) and a P_2U receptor on mucus-producing cells (26). Indeed, ATP (but not adenosine) induced a glycoprotein secretory response of isolated rabbit gastric mucous cells in primary culture (26). In the rabbit antral mucosa, blood flow is increased by adenosine, an effect mediated by an A₂ subtype purinoceptor (27). In rat gastric fundus, nerve terminals presumably release ATP and thereby influence smooth muscle activity. Indeed, in vitro studies on longitudinal muscle strips showed that adenosine produced relaxation, whereas ATP caused a phasic relaxation followed by a maintained spasm (22). Ohno et al. (24) observed that on smooth muscle cells of the guinea pig stomach fundus a transmural nerve stimulation evoked an excitatory junction potential that could be blocked by suramin, which is a putative P₂ purinoceptor inhibitor. These observations, together with our immunolocalization of ATPDase, indicate that this enzyme influences gastric acid and pepsin secretions, mucus production, and contractility of the stomach.

Intestine

High levels of ATPDase activity were found in the pig small intestine, in agreement with recent observations of ecto-ATPase on rat intestine (30). Columnar epithelial cells and reticular cells of the mucosa were highly reactive, and some significant immunoreactivity was found on smooth muscle cells. ATPDase appears to be strategically located on the luminal side of the intestine to modulate extracellular concentrations of ATP and other triphospho- and diphosphonucleosides, which could potentially come with chyme, including bile. ATPDase could also modulate the response to the release of ATP from the nervous system at the neuromuscular junction, thereby influencing intestinal motility. Finally, the presence of the enzyme on blood vessels indicates that it can influence the concentration of nucleotides in the circulatory system. Hence important roles for the ATPDase can be postulated, since purines exert major influences on peristalsis, electrolyte secretion, and blood flow (12, 15, 21, 27, 31). Concerning purinoceptor distribution and the physiological effects associated with these purinoceptors, there is evidence for an A₁-mediated inhibition of peristalsis (12) in rat jejunum, whereas in the smooth muscles of the same species, both adenosine and ATP enhanced spontaneous mechanical activity (21). The effects of extracellular purines on intestinal electrolyte secretion are supported by pharmacological studies and Northern blot analysis (31) on the T84 cell line, which have indicated that adenosine-stimulated Cl secretion of human intestinal epithelia is mediated by an A_2b purinoceptor. Purines can also alter intestinal blood flow. Indeed, sympathetic nerve stimulation causes a constriction of submucosal arterioles of guinea pig ileum that is mediated by ATP acting on P_2X receptors (11). Pennanen et al. (27) found a differential effect of adenosine on blood flow to subregions of the upper gastrointestinal tract of the rabbit, an effect that was mediated by A₂ purinoceptors. After ischemia-reperfusion of the rat intestine, adenosine arrests most of the inflammatory changes associated with reperfusion (15). These observations have potentially important implications for the treatment of intestinal diseases, including diarrhea.

Pancreas

ATPDase was originally localized in the pig pancreas by a rather nonspecific cytochemical approach (3). In the present study, the presence of ATPDase was confirmed in basolateral and apical membranes of acinar cells and zymogen granules, in agreement with our original observations. Intriguingly, a strong signal was observed on the ductal epithelium. Very little is known about the role of purines at this level of the gland. Activation of Cl and K⁺ conductances was observed after exposure of CFPAC-1 cells to ATP and UTP. These cells respond to nucleotide receptor activation with a transient increase in intracellular Ca²⁺ that stimulates these ionic currents (10). The role of ATPDase in zymogen granules remains a matter of speculation. Why the 78-kDa protein is truncated to a 54-kDa form is another intriguing question that remains unanswered (32). Moreover, the sectioned fragment remains associated with the zymogen granule membrane (unpublished observations).

Parotid

In the pig parotid gland, ATPDase was mainly associated with duct epithelium, with very little, if any, immunoreactivity on acinar cells. Myoepithelial cells dispersed among acini produced a significant signal. There have been reports of Ca²⁺-dependent ATPases associated with isolated parotid acinar cells, but as judged by their biochemical properties, these ATPases are different from ATPDases. However, a nucleoside triphosphatase described by Sato et al. (29) in bovine parotid gland and an apyrase described by Valenzuela et al. (34) in a microsomal fraction of rat salivary gland appear to correspond to the ATPDase described in this study and hence could well be associated with ductal epithelial and myoepithelial cells. In agreement with these findings, some studies on rat parotid acini show very little ecto-ATPase activity (9). Some P_2Z purinoceptors have been described on rat parotid acinar cells, which respond to ATP and mediate a Ca²⁺ increase caused by both an influx and a mobilization from intracellular stores (13). Another receptor, P_2X4, an ATP-gated ion channel, is expressed in serosal cells of salivary glands (6). Xu et al. (37) characterized and localized Ca²⁺ signaling receptors in rat submandibular salivary gland ducts (37). They showed that the ATP receptors were localized in the luminal membrane of the epithelial cell. If a similar localization for these ATP receptors exists in the pig parotid gland, the ATPDase could play a key role in the transport of electrolytes by modulating the extracellular ATP concentration in the salivary gland ducts.

Our study shows that ATPDase is present in large amounts in secretory epithelia from stomach, intestine, pancreas, and parotid gland. These results, in conjunction with the presence of purine-gated channels, lead us to believe that the enzyme modulates electrolyte secretion all along the digestive tract. The presence of the enzyme and P₂ purinoceptors on smooth muscle cells also suggests that the enzyme is involved in digestive tract motility and blood flow. Localization of the ATPDase supports the view that this ectonucleotidase is a prominent modulator of the action of extracellular purines and provides new clues for the interpretation of the role of ectonucleotidases in the digestive system.

	ACKNOWLEDGEMENTS

We thank Johanne Proulx for technical assistance.

	FOOTNOTES

This work was supported by grants from Fonds pour la Formation de Chercheurs et l'Aide à la Recherche du Québec (FCAR) and from the Natural Sciences and Engineering Research Council of Canada (NSERC). J. Sévigny received studentships from FCAR and from the Heart and Stroke Foundation of Canada.

Address for reprint requests: A. R. Beaudoin, Département de Biologie, Faculté des Sciences, Université de Sherbrooke, Sherbrooke, Québec J1K 2R1, Canada.

Received 1 December 1997; accepted in final form 5 May 1998.

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