PAR-2 modulates pepsinogen secretion from gastric-isolated chief cells

Stefano Fiorucci,1 Eleonora Distrutti,1 Barbara Federici,1 Barbara Palazzetti,1 Monia Baldoni,1 Antonio Morelli,1 and Giuseppe Cirino2

1Dipartimento di Medicina Clinica, Patologia, Clinica di Gastroenterologia ed Endoscopia Digestiva, Università di Perugia, 06122 Perugia, Italy, and 2Dipartimento di Farmacologia Sperimentale, Universita' di Napoli, 80131 Napoli, Italy

Submitted 10 September 2002 ; accepted in final form 3 April 2003


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 DISCLOSURES
 REFERENCES
 
In the present study, we investigated whether activation of protease-activated receptor type 2 (PAR-2) with SLIGRL (SL)NH2, a short mimetic agonistic peptide, directly stimulates pepsinogen secretion from gastric-isolated, pepsinogen-secreting (chief) cells. Immunostaining of gastric-dispersed chief cells with a specific anti-PAR-2 antibody demonstrated expression of PAR-2 receptors on membrane and cytoplasm. SL-NH2 and trypsin potently stimulated pepsinogen secretion (EC50 = 0.3 nM) and caused Ca2+ mobilization (EC50 = 0.6 nM). In contrast to SL-NH2, the scramble peptide LSIGRL-NH2 failed to stimulate pepsinogen release. Exposure to SL-NH2 also resulted in ERK1/2 phosphorylation and activation. Exposure of chief cells to phosphotyrosine kinase inhibitors and 2-(2-amino-3-methoxyphenyl)-4H-1-benzopyran-4-one, a selective MEK inhibitor, significantly reduced secretion induced by SL-NH2. Pepsinogen secretion induced by SL-NH2 was desensitized by pretreating the cells with the mimetic peptide and trypsin, and exposure to SL-NH2 abrogates pepsinogen secretion induced by carbachol and CCK-8, but not secretion induced by secretin and vasointestinal peptide. Exposure to Arg-Pro-Lys-Pro-Gln-Gln-Phe-Phe-Gly-Leu-Met-NH2 (substance P) but not to calcitonin gene-related peptide increased pepsinogen release. The neurokinin-1 receptor antagonist, N-acetyl-L-tryptophan 3,5-bis(trifluoromethyl)benzyl ester, inhibited substance P-stimulated pepsinogen secretion, whereas it did not affect secretion induced by SL-NH2. Collectively, these data indicate that PAR-2 is expressed on gastric chief cells and that its activation causes a Ca2+-ERK-dependent stimulation of pepsinogen secretion.

substance P; trypsin; ERK1; ERK2


PROTEINASES SUCH AS thrombin, trypsin, tryptase, and cathepsin G are now known to regulate target tissues via the proteolytic activation of cell surface G protein-coupled receptors called protease-activated receptors (PARs) (5, 9, 26, 27, 38). Proteases activate PARs by cleaving the NH2-terminal sequence of the extracellular exodomain. This cleavage event unmasks a new amino-terminal sequence, which in turn serves as a tethered ligand, binding the body of the receptor to trigger transmembrane signaling (9, 26). Molecular cloning has identified four PARs: PAR-1 and PAR-3, which are both preferentially activated by thrombin (38); PAR-2, which is selectively activated by trypsin (2, 29, 30); and PAR-4, which is activated by both thrombin and trypsin (38). In addition to endogenous proteases, PARs can be selectively activated by short agonistic peptides (APs) such as SLIGRL (SL)-NH2, a synthetic peptide that corresponds to the rat/mouse tethered ligand exposed after PAR-2 cleavage (1-4, 6, 8, 19-21).

Although PAR-2 is highly expressed in the gastrointestinal tract in epithelial cell, neuronal, and muscular elements (7, 11, 17, 30), the effect it exerts on gastric secretory functions is still poorly defined. In a recent study, however, Kawao et al. (20) reported that repeated injections of SL-NH2 to pylorus-ligated rats facilitates pepsinogen release in vivo, suggesting that PAR-2 acts as an endogenous mediator in pepsinogen secretion.

Pepsin is the main protein secreted by gastric epithelial cells and is released within the gastric lumen in response to a variety of neurohumoral stimuli by pepsinogen-secreting (chief) cells (10). This cell subtype represents {approx}40% of gastric epithelial cells and possesses G protein-coupled, seven-transmembrane-domain receptors that modulate pepsinogen release in response to peptide and nonpeptide agonists (10, 12-16, 31, 32, 34, 35). Binding of gastric chief cell receptors with specific agonists results in activation of at least two major intracellular pathways. Secretin, vasointestinal peptide (VIP), and PGE2 cause pepsinogen release by increasing intracellular concentrations of cAMP and PKA (10, 12-16, 32, 33, 35, 36). In contrast, gastrin, CCK, and muscarinic receptor agonists induce intracellular Ca2+ mobilization and PKC activation (10, 12-16, 32, 33, 35, 36). Although these pathways are functionally separated, a certain degree of interaction at postreceptor levels exists as demonstrated by the fact that simultaneous activation of PKA and PKC with specific agonist leads to a potentiation of pepsinogen release (16, 32). Furthermore, exposure of isolated chief cells to agonists might also result in pepsinogen secretion desensitization, a process that involves both receptor and postreceptor events, suggesting that interaction with other agonists might affect the ability of cells to respond to PAR-2 AP (12-16, 32). Because in several cell systems PAR-2 activation associates with Ca2+ mobilization and PKC activation (9, 27), we have designed a study to examine whether SL-NH2 stimulates pepsinogen release from gastric-isolated chief cells and to define intracellular messengers involved in this effect.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 DISCLOSURES
 REFERENCES
 
Materials. Male Hartley guinea pigs (200-400 g) were obtained from Charles River (Monza, Italy). DMEM was from GIBCO. HEPES, BSA fraction V, atropine, soybean trypsin inhibitor, collagenase (type I), carbamylcholine (carbachol), atropine, capsaicin, EGTA, fura 2-AM, BAPTA, and a lactate dehydrogenase (LDH) kit were from Sigma (St Louis, MO). Essential amino acid mixture and 1% essential vitamin mixture were from GIBCO (Milan, Italy); Percoll was from Pharmacia (Uppsala, Sweden); secretin and CCK-8 were from Peninsula Laboratories (St. Helens, Merceyside, UK). N-acetyl-L-tryptophan 3,5-bis(trifluoromethyl)benzyl ester (L-732,138), Arg-Pro-Lys-Pro-Gln-Gln-Phe-Phe-Gly-Leu-Met-NH2, (substance P), and Asp-Tyr-D-Trp-Val-D-Trp-D-Trp-Lys-NH2 (MEN-10,376) were from Biomol Research Laboratories (Plymouth Meeting, PA). 2-(2-Amino-3-methoxyphenyl)-4H-1-benzopyran-4-one (PD-98,059), a specific MEK inhibitor (1), was from Calbiochem (San Diego, CA). SLIGRL-NH2 and LSIGRL-NH2 (LS) were a kindly donated by Vincenzo Santagada and Giuseppe Caliendo (Department of Medicinal Chemistry, University of Naples, Italy).

Chief cell preparation. Chief cells from the guinea pig stomach were prepared as previously described (12-16) by collagenase digestion and Ca2+ chelation by omitting trypsin digestion (33). This method yields a cell population that is ~90% chief cells, 5% parietal cells, and 5% other cells. The purity of each chief cell preparation was verified daily by light microscopy, and the number of cells were counted. Chief cells were suspended in a standard incubation solution containing (in mM) 24.5 HEPES, 120 NaCl, 7.2 KCl, 1.5 Ca2Cl, 0.8 MgCl2, 2.6 KH2PO4, 14 glucose, 6 Na-pyruvate, 6 glutamate, 7 fumarate, 2 glutamine, and 0.1% (wt/vol) albumin, 1% (vol/vol) essential amino acid mixture and 1% (vol/vol) essential vitamin mixture. The pH was 7.4, and all incubations were performed at 37°C with 100% O2.

In some experiments, chief cells were prepared from guinea pigs pretreated with capsaicin. Under halothane anesthesia, animals received three doses of capsaicin (25, 50, and 50 mg/kg sc) dissolved in vehicle (10% ethanol, 10% Tween 80, sterile physiological saline) over 32 h (at 0, 6, and 32 h, respectively; 125 mg/kg total dose) or an equivalent volume of vehicle. This treatment has previously been demonstrated to deplete substance P-like and calcitonin gene-related peptide (CGRP)-like immunoreactivity from extrinsic afferent neurons in the gastrointestinal tract (18). The efficacy of the capsaicin treatment was verified by the eye-wiping test, as described previously (17, 18). All control animals received administration of vehicle. Chief cells were prepared from capsaicin-treated and control guinea pigs 7 days after the last injection of capsaicin or vehicle.

RT-PCR. After the rats were killed, stomachs were removed and immediately snap-frozen in liquid nitrogen and stored at -80°C until used. Total RNA was isolated by using TRIzol reagent (Life Technologies, Milan, Italy) as previously described (17). PCR was performed by using specific primers. For {beta}-actin (543 bp), the sense primer was 5'-TGT GAT GGT GGG AAT GGG TCA G-3' and the antisense primer was 5'-TTT GAT GTC ACG CAC GAT TTC C-3' (Stratagene, La Jolla CA). Oligonucleotide primers for guinea pig PARs were forward, 5'-CAT GTT CAG CTA CTT CCT CTC CTT-3' and reverse, 5'-GGT TTT AAC ACT GGT GGA GCT TGA-3' and were chosen to amplify a 472-bp fragment. The cDNA was amplified with a "hot start" reaction in 20 µl of reaction containing 5 µl cDNA product, 2 µl PCR buffer (200 mM Tris · HCl, pH = 8; 4,500 mM KCl), 200 µM dNTPs, 1 µM of sense and antisense primers, 1.5 mM MgCl2, 1 U platinum Taq polimerase (Life Technologies), and water in a Hybaid PCR Sprint thermocycler (Celbio, Milan, Italy). PCR was carried out as previously described (17). The size of PCR products was assessed by comparison with a 1-µg 100-bp DNA ladder (Life Technologies). The gel was photographed under ultraviolet transillumination with a Kodak Digital Science ID Image Analysis Software (Kodak, Milan, Italy), and images were digitalized. Each assay was carried out in triplicate. The {beta}-actin was used as a control for both reverse transcription and the PCR reaction itself.

Immunohistochemistry. Cytospins of gastric chief cells were air-dried overnight and fixed in acetone at 4°C for 5 min, washed in PBS, dipped in methyl alcohol with 0.3% hydrogen peroxide for 10 min for blocking of endogenous peroxidase activity, washed three times in PBS, and incubated in 0.1% albumin for 30 min. Immunostaining of PAR-2 was performed by using a rabbit polyclonal antibody (B5) that specifically recognizes PAR-2, targeted to a peptide corresponding to the cleavage/activation site of rat PAR-2 (30GPNSKGR{downarrow}SLIGRL-DT46P-YGGC) (19, 25). Sections were first incubated in 1% normal goat serum for 30 min and then in the B5 antibody at a dilution of 1:1,000 for 16 h at 4°C (19). Immunoreactivity was visualized with the use of biotinylated goat anti-rabbit IgG followed with streptavadin-conjugated peroxidase (Sigma, St. Louis, MO) and color generation with diaminobenzidine for 30 min. Smears were counterstained with hematoxylin and mounted in ACQUOVITREX (Carlo Erba, Milan, Italy), and images were recorded by digital photomicrography. Controls were obtained by incubating smears with B5 antibody preadsorbed with the synthetic peptides (19).

Effect of SL-NH2 and LS-NH2 on pepsinogen secretion. Chief cells (300,000/ml) were suspended in the standard incubation solution for 30 min alone or with concentrations of SL-NH2 or trypsin ranging from 1 pM to 100 nM. In some experiments, chief cells exposed to SL-NH2 were coincubated with maximally effective concentrations of the following agents: 30 nM secretin, 1 µM VIP, 1 µM PGE2, 30 µM carbachol, and 3 nM CCK; pepsinogen released in cell supernatants was measured. To investigate whether pepsinogen secretion induced by SL-NH2, could be desensitized by another specific agonist, cells were pretreated with maximally effective concentrations of secretin, VIP, PGE2, carbachol, and CCK for 30 min (12-16, 31, 32, 34, 35), and then, after extensive wash, cells were stimulated with 1 nM SL-NH2 and pepsinogen release in cell supernatants was measured. Pepsinogen released into the supernatant (250 µl) was assayed by using acid-denatured hemoglobin as substrate, as described previously (12-16). Each sample was both incubated and assayed in duplicate. Pepsinogen secreted during incubation is expressed as a percentage of total pepsinogen present in chief cells at the beginning of the incubation minus the pepsinogen secreted before starting incubation.

Effect of SL-NH2 on [Ca2+]i. Intracellular Ca2+ concentration ([Ca2+]i) was measured in dispersed chief cells (2 x 106/ml) loaded with fura 2-AM by using a Hitachi 2000 (Milan, Italy) fluorescence spectrophotometer (13, 14). [Ca2+]i was measured in cells stimulated with SL-NH2 (from 1 pM to 1 nM) or 1 nM trypsin. Control cells were stimulated with 30 µM carbachol. [Ca2+]i was calculated according to Grynkiewicz et al. (19). The fura 2-AM/Ca2+ signal was calibrated at the end of each recording by adding digitonin followed by EGTA as described (13, 14).

To investigate whether SL-NH2 increased Ca2+ influx, we used Mn2+ asaCa2+ surrogate (14). Fura 2-loaded cells were resuspendend in a Ca2+-free buffer and stimulated with 10 nM SL-NH2 (14). Fluorescence was excited at 360 nm, i.e., the isosbestic wavelength at which Ca2+ does not affect fura 2 fluorescence and changes in fluorescence intensity are only caused by Mn2+ quenching. Emission was recorded at 505 nm. Maximal Mn2+ quenching was calibrated in each preparation at the end of recording with digitonin (14).

Western blot analysis of ERK. Isolated chief cells were pelleted and lysed in ice in (in mM): 50 Tris · HCl (pH 8.0), 150 NaCl, 1 EGTA (pH 8.0), 100 NaF, 1% MgCL2, 1 Na3VO4, 1 PMSF, and 10% glycerol, 1% vol/vol Triton X-100, 10 µg/ml leupeptin, and 5 µg/ml aprotinin. Insoluble materials were removed by centrifugation at 12,000 g at 4°C for 10 min, and protein concentration was determined by protein assay reagent (Bio-Rad Laboratories, Hercules, CA). For Western blot analysis, 50 µg of total lysates were electrophoresed on a 11% SDS-PAGE, blotted onto nitrocellulose membrane, and incubated with an anti-phospho-ERK, an antibody that specifically recognizes the active, phosphorylated form of ERK and that reacts with both p42 and p44 isoforms (Promega, Madison, WI) (14). ERK kinase activation in response to SL-NH2 is demonstrated by the appearance of protein bands at 42 and 44 kDa.

ERK activity. ERK activity was assessed by using the p42/p44 MAP kinase assay kit (Cell Signaling Tecnologies, Beverly, MA) according to the manufacturer's indications. In this method, a monoclonal antibody against p42/p44 ERK (Thr202 and Tyr204) is used to selectively immunoprecipitate phosphorylated ERK from cell lysates. The resulting immunoprecipitate is then incubated with an Elk-1 fusion protein in the presence of ATP, allowing active ERK to phosphorylate Elk-1. Immunoprecipitates were then resolved on 10% SDS-PAGE, blotted onto nitrocellulose membrane, and incubated with a phospho-Elk-1 antibody. Membranes were then incubated with horseradish peroxidase-conjugated anti-rabbit secondary antibody. Band intensity was quantified as described previously (14) and expressed as fold of increase over basal values. To investigate whether ERK activation was required for pepsinogen release induced by PAR-2 AP, cells were incubated with 1-50 µM PD-98,059, a specific MEK inhibitor (1), and pepsinogen release induced by SL-NH2 was measured.

Statistical analysis. Data reported are means ± SE of the number of experiments indicated. The statistical analysis was carried out by using a GraphPad Prism 3 (GraphPad Software, San Diego, CA). ANOVA and Student's t-test for paired data were employed when appropriate.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 DISCLOSURES
 REFERENCES
 
Chief cells express PAR-2. At the immunohistochemical analysis, PAR-2-like immunoreactivities were found in the plasma membrane and cytosol of gastric chief cells (Fig. 1A). PAR-2-like immunoreactivities were also found in gastric mucous cells but not in parietal cells. The PAR-2 immunostaining was abolished by preabsorption of the B5 antibody with 20 µg/ml of antigenic peptide (Fig. 1B). The RT-PCR analysis (Fig. 1C) demonstrates characteristic size transcript for PAR-2 (472 bp) in this cell preparation. Exposure to SL-NH2 and trypsin has no effect on PAR-2 mRNA expression (data not shown).



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Fig. 1. Gastric-isolated chief cells express proteinase-activated (PAR)-2 receptor. A: microphotographs of immunostaining of PAR-2 on gastric-isolated chief cells (magnification, x400). B: negative control. Chief cells were stained with the anti-PAR-2 antibody preabsorbed with a peptide used for immunization (magnification, x400). C: RT-PCR demonstrating PAR-2 mRNA transcript of 3 separate PCR experiments. -, negative control (water); +, positive control (Par-2 positive cDNA); CC, chief cell lysates.

 

SL-NH2 STIMULATES PEPSINOGEN RELEASE FROM GASTRIC-ISOLATED CHIEF CELLS. As illustrated in Fig. 2, exposure to SL-NH2 resulted in a concentration- and time-dependent stimulation of pepsinogen secretion. Basal pepsinogen secretion of 2.4 ± 0.4% increased to 19.9 ± 1.3% in response to incubation with 1 nM SL-NH2 and declined slightly with higher concentrations, resulting in a bell-shaped concentration-response curve. The pepsinogen release induced by PAR-2 was significantly above basal values at 10 pM, half maximal at 0.3 nM, and maximal at 1 nM. Trypsin also stimulates pepsinogen secretion (Fig. 2B). At the concentration of 1 nM, trypsin caused a 10-fold increase of pepsinogen release. The kinetic of pepsinogen secretion induced by SL-NH2 and trypsin was biphasic (Fig. 2C). Approximately 80% of total pepsinogen secretion induced by 1 nM SL-NH2 was observed in the first 15 min of incubation, resulting in a rate of pepsinogen release of 0.76 ± 0.05% per min. Pepsinogen secretion declined thereafter (15- to 60-minute interval) to 0.16 ± 0.02% per minute. In contrast to SL-NH2, the scramble peptide LS-NH2 did not stimulate pepsinogen release. Changes in pepsinogen release were not due to alteration of cell membrane permeability. Indeed, LDH activity in cell supernatants was 2.1 ± 0.5, 2.4 ± 0.5, 2.2 ± 0.4, and 3.2 ± 0.5% of total in control cells and in cells incubated with 1 nM SL-NH2, LS-NH2, or trypsin, respectively (n = 6, not significant).



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Fig. 2. A: effect of increasing concentrations of SLIGRL (SL)-NH2 and LSIGRL (LS)-NH2 on pepsinogen release from gastric-isolated chief cells. Cells were incubated for 30 min at 37°C. B: effect of increasing concentrations of trypsin on pepsinogen release from gastric-isolated chief cells. C: time-course of pepsinogen secretion induced by 1 nM SL-NH2, LS-NH2, and trypsin. Results are means ± SE of 6 experiments. *P < 0.01 vs. basal.

 

SL-NH2 causes Ca2+ mobilization. [Ca2+]i increased from 184.4 ± 45.3 to 543.7 ± 55.6 and 689.4 ± 64.6 nM after exposure to 10 nM SL-NH2 and trypsin (Fig. 3A). Ca2+ mobilization induced by SL-NH2 was time and concentration dependent with an ED50 of {approx}0.6 nM (Fig. 3, A and B). Removal of extracellular Ca2+ attenuated but did not abolish Ca2+ signals induced by SL-NH2, indicating that PAR-2 activation in chief cells mobilizes both intra- and extracellular sources of Ca2+. Consistent with this view, exposure to SL-NH2 increased the quenching rate of the fura 2 signal in the presence of Mn2+, indicating a net Ca2+ influx during PAR-2 activation (Fig. 3C). To investigate whether pepsinogen release induced by SL-NH2 was Ca2+ dependent, chief cells were incubated with 1 mM BAPTA, an agent that binds intracellular Ca2+, in a Ca2+-free medium (Fig. 4, A and B). Pepsinogen secretion of 2.1 ± 0.3 rose to 18.5 ± 1.9% in chief cells exposed to 1 nM SL-NH2 (P < 0.05, n = 4 experiments) but dropped to 8.5 ± 0.8 in cells incubated in a Ca2+-free medium and to 5.2 ± 0.5% in cells preincubated in a Ca2+-free medium in the presence of 1 mM BAPTA, (Fig. 4, A and B; P < 0.01, n = 6). In this setting, exposure to 1 nM SL-NH2 failed to stimulate Ca2+ mobilization (P < 0.01, n = 6), indicating that pepsinogen release induced by PAR-2 activation is Ca2+ dependent and requires both Ca2+ release from intracellular pools and influx of extracellular Ca2+. In confirmation of this, we found a close correlation between pepsinogen release and [Ca2+]i in response to SL-NH2 (Fig. 4C).



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Fig. 3. PAR-2 activation causes Ca2+ mobilization. A: effect of 10 nM SL-NH2 on intracellular Ca2+ concentration ([Ca2+]i) in chief cells loaded with fura 2. Results are means ± SE from 12 determinations. B: effect of increasing concentrations of SL-NH2 on [Ca2+]i in chief cells loaded with fura 2. Results are means ± SE from 6-8 determinations. C: effect of Mn2+ addition to fura 2-loaded cells incubated with or without SL-NH2.Mn2+ was used as a substitute for Ca2+. More pronounced is the quenching of the fura 2 signal; the higher is Mn2+ (i.e., Ca2+) influx. Cells were incubated in a Ca2+-free medium and then stimulated with SL-NH2 10 nM or no agent. After 1 min of incubation, 25 µM Mn2+ was added to the cell suspension. Fluorescence intensity was normalized to 100% just before Mn2+ addition. Data are means ± SE of at 12-15 determinations.

 


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Fig. 4. Pepsinogen secretion induced by SL-NH2 is Ca2+ dependent. A and B: effect of 1 mM BAPTA and [Ca2+]i on pepsinogen secretion and [Ca2+]i induced by 1 nM SL-NH2. Results are means ± SE from 6 experiments (A) and 12 determinations (B). *P < 0.001 vs. cells treated with SL-NH2 alone. C: relationship between [Ca2+]i and pepsinogen release in chief cells exposed to SL-NH2. Data are obtained from Figs. 2A and 3B and are expressed as %maximal values of [Ca2+]i and pepsinogen release measured with 1 nM SL-NH2.

 

Secretion induced by SL-NH2 is modulated by Ca2+-mediated agonists. Because these data indicate that SL-NH2 causes Ca2+ mobilization, we then investigated whether agents that stimulate pepsinogen release through a Ca2+-dependent or -independent mechanism modulates the chief cells' response to PAR-2 activation. As shown in Table 1, we found that, although SL-NH2-stimulated secretion was additive to that induced by maximally effective concentrations of secretin, VIP, and PGE2 (12-16, 31, 32, 34, 35), secretion induced by maximally effective concentrations (12-16, 31, 32, 34, 35) of phospholipase C (PLC)-Ca2+-activating agents (carbachol, CCK-8, and thapsigargin) remained unchanged. Thus pepsinogen secretion induced by SL-NH2 is additive to secretion caused by agents that activate a cAMP-PKA pathway. To further investigate whether potentiation observed with maximally effective concentrations of agonists shown in Table 1 was maintained when submaximally effective concentrations of these agonists were used, chief cells incubated with increasing concentrations of PGE2 were challenged with 1 nM SL-NH2. As shown in Fig. 5A, in this setting, SL-NH2 still potentiates pepsinogen release induced by PGE2. The reverse was also true, because the maximally effective concentration of PGE2 (1 µM) potentiates pepsinogen release induced by submaximally effective concentrations of SL-NH2 (Fig. 5C). As illustrated in Fig. 5B, the rate of pepsinogen secretion induced by 1 nM SL-NH2 and 1 µMPGE2 was statistically different from that induced by each agent alone, resulting in a pepsinogen releasing rate of 1.5 ± 0.1% per minute in the first 15-min period (P < 0.01 vs. cells incubated with SL-NH2 or PGE2 alone, n = 5). In contrast, coincubating chief cells with submaximally effective concentrations of carbachol did not affect pepsinogen release induced by a maximally effective concentration of SL-NH2 (Fig. 5, D and E), nor did carbachol-potentiated secretion induced by submaximally effective concentrations of SL-NH2 (Fig. 5F). Furthermore, pepsinogen secretion induced by SLNH2 was not affected by the addition of 100 µM atropine to the incubation medium; pepsinogen release was 18.3 ± 2.5% in cells incubated with 1 nM SL-NH2 alone and 17.8 ± 2.8% in cells incubated with SL-NH2 plus atropine (n = 4, P > 0.05). Similarly, secretion induced by SL-NH2 was not affected by incubating chief cells with indomethacin. Thus basal secretion of 1.4 ± 0.4 rose to 17.5 ± 3.1% in response to SL-NH2 (1 nM) and to 18.3 ± 2.4% in cells incubated with SL-NH2 plus indomethacin (100 µM) (P > 0.05 compared with SL-NH2 alone). In cells incubated with indomethacin alone, secretion was 4.2 ± 1.2 (P < 0.05 vs. control, n = 4).


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Table 1. Effect of SL-NH2 on agonist-induced pepsinogen secretion in gastric isolated chief cells

 


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Fig. 5. cAMP-PKA-mediated agents potentiate pepsinogen secretion induced by SL-NH2. A: SL-NH2 potentiates pepsinogen release induced by submaximally effective concentrations of PGE2. Results are means ± SE of 6 experiments. *P < 0.01 vs. PGE2 alone. The broken line indicates pepsinogen secretion induced by 1 nM SL-NH2. B: time course of pepsinogen release induced by 1 nM SL-NH2 in combination with 1 µM PGE2. Results are means ± SE of 6 experiments. *P < 0.01 vs. SL-NH2 alone. C: the addition of 1 µM PGE2 potentiates pepsinogen release induced by submaximally effective concentrations of SL-NH2. Results are means ± SE of 6 experiments. *P < 0.01 vs. PGE2 alone. The broken line indicates pepsinogen secretion induced by 1 µM PGE2. D: SL-NH2 fails to potentiate pepsinogen secretion induced by submaximally effective concentrations of carbachol. Results are means ± SE of 6 experiments. P > 0.05 vs. SL-NH2 alone. The broken line indicates pepsinogen secretion induced by 1 nM SL-NH2. E: time course of pepsinogen release induced by 1 nM SL-NH2 in combination with 30 µM carbachol. Results are means ± SE of 6 experiments. F: the addition of 30 µM carbachol failed to potentiate pepsinogen release induced by submaximally effective concentrations of SL-NH2. Results are means ± SE of 6 experiments. P > 0.05 vs. SL-NH2 alone. The broken line indicates secretion induced by 30 µM carbachol.

 

SL-NH2 desensitizes pepsinogen secretion induced by Ca2+-mediated agonists. As illustrated in Table 2 and Fig. 6, A and B, chief cells pretreated with SL-NH2 and trypsin (1 nM) not only released significantly less pepsinogen in response to a second stimulation with the mimetic peptide, but also pepsinogen secretion induced by trypsin, carbachol, CCK-8, and thapsigargin was markedly curtailed (Table 2). In contrast, preexposure to SL-NH2 (1 nM) had no effect on secretion stimulated by secretin, VIP, and PGE2 (Table 2).


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Table 2. Effect of SL-NH2 on agonist-induced pepsinogen secretion by isolated chief cell

 


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Fig. 6. Desensitization of pepsinogen secretion induced by SL-NH2. Preincubating gastric chief cells with 1 nM SL-NH2 desensitizes pepsinogen secretion induced by a second challenge with 1 nM SLNH2 (A) or 1 nM trypsin (B). Results are means ± SE of 6 experiments. *P < 0.01 (ANOVA) compared with cell not pretreated with SL-NH2.

 

SL-NH2 causes ERK1/2 phosphorylation. Exposure of gastric-isolated chief cells to SL-NH2 caused ERK phosphorylation (Fig. 7A). Both 42- and 44-kDa ERK were activated after 5 min of incubation with 10 nM SL-NH2 and are maximally activated within 15 min. ERK activation by SL-NH2 is concentration dependent (Fig. 7B). These data were confirmed when ERK1/2 activity was assayed. As illustrated in Fig. 7C, exposure to SL-NH2 but not to LS-NH2 caused a concentration-dependent increase in ERK activity (P < 0.01 vs. control, n = 5). Because these data implied that tyrosine kinases are involved in modulating ERK activation in response to SL-NH2, we then assessed whether tyrosine kinase and ERK inhibitors modulate pepsinogen release in response to PAR-2 activation. As shown in Table 3, exposure to genistein, tyrophostin 51, staurosporine, and PD-98,059, a selective MEK inhibitor (1), caused a 40-60% inhibition of pepsinogen release induced by SL-NH2 (P < 0.01 vs. SL-NH2 alone, n = 6). As shown in Table 3, inhibition of pepsinogen secretion induced by genistein, tyrophostin 51, staurosporine, and PD-98,059 associates with inhibition of ERK phosphorylation.



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Fig. 7. PAR-2 activation causes ERK1/2 phosphorylation. A: SL-NH2 causes a time-dependent phosphorylation of ERK1/2. Chief cells were incubated with 10 nM SL-NH2 for the indicated time, and cell lysates were blotted as indicated in MATERIALS AND METHODS. Two ERK bands of 42 and 44 kDa are demonstrated. Molecular masses are shown on the left. B: SL-NH2 causes a concentration-dependent phosphorylation of on ERK1/2. Two ERK bands of 42 and 44 kDa are demonstrated. Molecular masses are shown on the left. A and B are representative of at least 3 similar experiments. C: effect of increasing concentrations of SL-NH2 and LS-NH2 on ERK1/2 activity. Results are means ± SE of 6 experiments. *P < 0.05 vs. untreated.

 

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Table 3. Effect of tyrosine kinase inhibitors and a MEK inhibitor on pepsinogen release and ERK1 and ERK2 phosphorylation induced by SL-NH2

 

Pepsinogen release induced by SL-NH2 is substance P independent. Because previous studies (17, 19-21, 33, 36) have demonstrated that in vivo administration of SL-NH2 stimulates substance P and CGRP release from enteric neurons, we then investigated whether these two neuropetides modulate secretory response to SL-NH2 in vitro. As illustrated in Fig. 8, although CGRP failed to stimulate pepsinogen secretion, exposure to 1 µM substance P caused a fourfold increase in pepsinogen release (P < 0.01 vs. medium, n = 6). Consistent with previous reports (15, 22), this effect was reversed by coincubating the cells with a neurokinin (NK)1, but not NK2, antagonist. Indeed, although exposure to 10 µM L-732,138, a selective NK1 antagonist, abolished the stimulation caused by substance P (P < 0.05, n = 6), exposure to MEN-10,376, a highly selective NK2 antagonist, failed to reduce pepsinogen secretion induced by substance P (P > 0.05, n = 6). Interestingly, both compounds failed to affect pepsinogen secretion induced by SL-NH2. In addition, as shown in Fig. 8B, capsaicin pretreatment had no effect on chief cells' response to 1 nM SL-NH2 (P > 0.05 capsaicin-pretreatment vs. naive, n = 4).



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Fig. 8. Effect of selective NK1 and NK2 antagonists on pepsinogen secretion induced by SL-NH2, substance P (SP), and calcitonin gene-related peptide (CGRP). A: isolated gastric chief cells were prepared as described in MATERIALS AND METHODS and were incubated for 30 min with 1 nM SL-NH2, 1 µM SP, and 1 µM CGRP alone or in the presence of 10 µM L-732,138 and MEN-10,376. Results are means ± SE from 6 experiments. *P < 0.01 vs. medium; **P < 0.01 vs. SL-NH2 alone. B: ablation of extrinsic afferent neurons of gastrointestinal tract in vivo by capsaicin has no effect on chief cell sensitivity to SL-NH2. Guinea pigs were pretreated in vivo with capsaicin, and chief cells were prepared from naiïve or neuron-ablated animals. Results are means ± SE from 4 experiments. *P < 0.01 vs. basal.

 


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 DISCLOSURES
 REFERENCES
 
In the present study, we have provided evidence that isolated gastric chief cells express receptor for PAR-2 and that PAR-2 activation with short mimetic peptide potently stimulates pepsinogen secretion by activating a number of intracellular messengers including protein-tyrosine kinases, ERK1/2, and Ca2+.

In other cell models, PAR-2 couples G{alpha}q/11 and phospholipase C{beta}, leading to hydrolysis of phosphatidylinositol bisphosphate, Ca2+ mobilization, and activation of PKC and ERK1/2 (8, 9, 26). Here we demonstrated that exposure of gastric chief cells to SL-NH2 causes Ca2+ mobilization but that Ca2+ is required to release pepsinogen. Indeed, incubating the cells with BAPTA, an agent that binds intracellular Ca2+, in a Ca2+-free medium abolishes pepsinogen secretion stimulated by SL-NH2 (12-14). Consistent with the view that pepsinogen release induced by PAR-2 activation is due to a Ca2+-mediated mechanism is the observation that pepsinogen release induced by SL-NH2 was additive with secretion stimulated by PGE2, secretin, and VIP (i.e., agents that stimulate pepsinogen release by cAMP/PKA-dependent pathway) but not with pepsinogen release induced by carbachol, gastrin, and thapsigargin, i.e., agents that cause pepsinogen release through a Ca2+-mediated pathway (12-16, 31, 32, 34, 35). Although these results do not allow us to determine whether SL-NH2 and cAMP-PKA activating agents act on different intracellular pools of pepsinogen, they prove that the ability of chief cells to respond to PAR-2 is largely dependent on the presence of other agonists/antagonists at the chief cell surface. Confirming the complexity of these interactions, we have also documented that SL-NH2 could desensitize pepsinogen secretion induced by carbachol and CCK-8 but not secretion induced by secretin, VIP, and PGE2 (12-16, 31).

DeFea et al. (8) have previously demonstrated that PAR-2 activates ERK1/2 through a PKC-dependent, pertussis toxin-insensitive pathway by a mechanism that requires receptor association into a multiprotein complex comprising internalized PAR-2 receptor, {beta}-arrestin, raf-1, activated ERK1/2, and perhaps other components of the MAPK pathway (8, 9, 24, 26). Here we demonstrated that exposure of gastric chief cells to SL-NH2 increases ERK1/2 activity and that tyrosine kinase inhibitors, genestein, staurosporine, and tyrophostin 5 inhibit ERK1/2 phosphorylation induced by SL-NH2 and cause a 50-60% inhibition of pepsinogen release (14). Confirming the role of ERK1/2 in regulating pepsinogen release induced by SL-NH2, exposure to PD-98,059, a selective MEK inhibitor (1), significantly reduced pepsinogen release induced by SL-NH2. However, the finding that the amount of pepsinogen secretion inhibited by PD-98,059 and tyrosine kinase inhibitors is lower than that inhibited by BAPTA suggests that ERK1/2 function to increase the amplitude of the secretory response rather than to provide an obligate signal for this response to occur (14).

Desensitization is the decrease of a biological response to repeated exposure to an agonist or to continued presence of an agonist (12-16). In the present study, we demonstrated that preexposure to SL-NH2 results in a rapid desensitization of pepsinogen secretion induced by subsequent stimulation with SL-NH2, trypsin, and other Ca2+-mediated agonists (12-16, 31, 32, 34, 35). Trypsin, the putative endogenous ligand for PAR-2, could desensitize pepsinogen secretion induced by SL-NH2 by at least three different mechanisms (3, 9, 26): 1) enzymatic cleavage of the receptor, which ensures that a single receptor molecule, once cleaved, cannot be reactivated by trypsin; 2) receptor phosphorylation and uncoupling from G proteins; and 3) inhibition of more distal steps of intracellular signaling machinery. In contrast, SL-NH2 activates PAR-2 without receptor cleavage; thus desensitization caused by this peptide could only result from G proteins uncoupling and/or inhibition of downstream signals. The finding that preexposure to SL-NH2 desensitizes pepsinogen secretion induced by carbachol and CCK-8 indicates that the main mechanism of pepsinogen secretion desensitization caused by this peptide lies downstream to the receptor (12-16, 31). This hypothesis is further supported by data obtained with thapsigargin. Indeed, exposure of gastric-isolated chief cells to thapsigargin inhibits a Ca2+-ATPase that is responsible for the reuptake of cytosolic Ca2+ by intracellular stores, resulting in sustained increase of [Ca2+]i and pepsinogen release (14). Thus the finding that SL-NH2 desensitizes pepsinogen secretion induced by thapsigargin is further evidence that PAR-2 activation stimulates a Ca2+-dependent pathway and that Ca2+ mobilization is strictly required for pepsinogen release induced by SL-NH2.

Although PAR-2 has been characterized as a trypsin-sensitive receptor (1-4, 9, 26), the potential for trypsin itself to be the preferred endogenous activator of PAR-2 in all tissues remains controversial. The high level of expression of PAR-2 in the small intestine and colon suggest the potential for direct activation of PAR-2 to occur by trypsin released from its zymogen precursor, the trypsinogen, by enteric peptidases within the intestinal lumen (1, 20, 21, 25, 27-30). In other parts of the gastrointestinal tract, including the stomach, however, it is unlikely that sufficient trypsin is generated in the lumen to directly activate PAR-2. However, trypsinogen concentrations in the blood increase physiologically after a meal, making possible blood-derived trypsin activation of gastric PAR-2, establishing an integratory feedback loop that might contribute to modulate pepsinogen release in response to food (26). Indeed, pancreatic trypsin is secreted in an episodic manner, with high levels after feeding and lower levels between meals. The finding that exposure to PAR-2 desensitizes pepsinogen secretion by a combination of effects on several components of the signaling pathway suggests that this period of cell refractoriness could be used by chief cells to replenish surface PAR-2 by mobilizing intracellular pools and/or by synthesis of new receptors.

One persistent ambiguity regarding the physiological role of PAR-2 derives from the observation that PAR-2 receptors are found on 50-60% of enteric neurons and that activation of PAR-2 by endogenous ligands or synthetic peptides stimulates capsaicin-sensitive sensory neurons to release CGRP and substance P, an endogenous NK1-preferential agonist with some NK2 activity (7, 9, 11, 26, 33, 36). Treating rodents with capsaicin to ablate extrinsic afferent innervation reduces inflammation induced by SL-NH2, demonstrating that CGRP and substance P are responsible, at least in part, for the final physiological effect of PAR-2 (4, 5, 11, 17, 20, 23, 36). With the use of a purified preparation of gastric chief cells, we have now demonstrated that PAR-2 activation directly stimulates pepsinogen release from gastric-isolated chief cells. We (15) and others (22) have previously demonstrated that gastric chief cells release pepsinogen in response to stimulation with sensory neuropetides. However, a role for these mediators in the effect exerted by PAR-2 is not similar because: 1) CGRP has no direct stimulatory effect on pepsinogen release by gastric chief cells (present study); 2) although substance P stimulates pepsinogen release, this peptide is significantly less effective than SL-NH2 (present data and Ref. 15); 3) selective NK1 and NK2 receptor antagonist did not alter pepsinogen release induced by SL-NH2 (15, 22); and 4) in vivo exposure to capsaicin fails to affect the response of gastric chief cells to SL-NH2. Together with the finding that PAR-2 is expressed on gastric chief cells and that PAR-2 activation in vivo and/or in vitro stimulates salivary (19), gastric (23), pancreatic (25, 28), and intestinal (37) secretion, our data establish a common regulatory function for this receptor through the gastrointestinal tract.

In conclusion, our results demonstrate that gastric-isolated, pepsinogen-secreting chief cells express a receptor for PAR-2 and that PAR-2 activation potently stimulates pepsinogen by activating a number of intracellular messengers, including protein-tyrosine kinase ERK1/2, and by causing Ca2+ mobilization. Together with previous findings demonstrating that PAR-2 activates salivary and pancreatic secretion, our data support the hypothesis that PAR-2 is a regulatory receptor for gastrointestinal exocrine glands.


    DISCLOSURES
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 DISCLOSURES
 REFERENCES
 
This investigation was supported, in part, by a grant from the Ministero dell' Universita' e della Ricerca Scientifica e Tecnologica, Rome, Italy.


    ACKNOWLEDGMENTS
 
We thank Dr. Morley D. Hollenberg, Dept. of Pharmacology and Therapeutics, University of Calgary, Calgary, AB, Canada, for the gift of B5 antibody.


    FOOTNOTES
 

Address for reprint requests and other correspondence: S. Fiorucci, Clinica di Gastroenterologia ed Epatologia, Policlinico Monteluce, Via E. Dal Pozzo, 06122 Perugia, Italy (E-mail: fiorucci{at}unipg.it).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


    REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 DISCLOSURES
 REFERENCES
 

  1. Alessi DR, Cuenda A, Cohen P, Dudley DT, and Saltiel A. PD 098059 is a specific inhibitor of the activation of mitogen-activated protein kinase kinase in vitro and in vivo. J Biol Chem 270: 27489-27494, 1995.[Abstract/Free Full Text]
  2. Bohm SK, Kong W, Bromme D, Smeekens SP, Anderson DC, Connolly A, Kahn M, Nelken NA, Coughlin SR, Payan DG, and Bunnett NW. Molecular cloning, expression and potential functions of the human proteinase-activated receptor-2. Biochem J 314: 1009-1016, 1996.[ISI][Medline]
  3. Bohm SK, Khitin LM, Grady EF, Aponte G, Payan DG, and Bunnett NW. Mechanisms of desensitization and resensitization of proteinase-activated receptor-2. J Biol Chem 271: 22003-22016, 1996.[Abstract/Free Full Text]
  4. Carr MJ, Schechter NM, and Undem BJ. Trypsin-induced, neurokinin-mediated contraction of guinea pig bronchus. Am J Respir Crit Care Med 162: 1662-1667, 2000.[Abstract/Free Full Text]
  5. Cirino G, Bucci M, Napoli C, and Cicala C. Inflammationcoagulation network: are serine protease receptors the knot? Trends Pharmacol Sci 21: 170-172, 2000.[ISI][Medline]
  6. Corvera CU, Dery O, McConalogue K, Bohm SK, Khitin LM, Caughey GH, Payan DG, and Bunnett NW. Mast cell tryptase regulates rat colonic myocytes through proteinase-activated receptor 2. J Clin Invest 100: 1383-1393, 1997.[Abstract/Free Full Text]
  7. D'Andrea MR, Derian CK, Leturcq D, Baker SM, Brunmark A, Ling P, Darrow AL, Santulli RJ, Brass LF, and Andrade-Gordon P. Characterization of protease-activated receptor-2 immunoreactivity in normal human tissues. J Histochem Cytochem 46: 157-164, 1998.[Abstract/Free Full Text]
  8. DeFea KA, Zalevsky J, Thoma MS, Dery O, Mullins RD, and Bunnett NW. {beta}-Arrestin-dependent endocytosis of proteinase-activated receptor 2 is required for intracellular targeting of activated ERK1/2. J Cell Biol 148: 1267-1281, 2000.[Abstract/Free Full Text]
  9. Dery O, Corvera CU, Steinhoff M, and Bunnett NW. Proteinase-activated receptors: novel mechanisms of signaling by serine proteases. Am J Physiol Cell Physiol 274: C1429-C1452, 1998.[Abstract/Free Full Text]
  10. Feldman M. Gastric secretion: normal and abnormal. In: Sleisenger & Fordtran's Gastrointestinal and Liver Disease, edited by Feldman M, Sleisenger MH, and Sharmischmidt BF. Philadelphia, PA: Saunders, 1998, p. 587-603.
  11. Fiorucci S and Distrutti E. Role of PAR-2 in pain and inflammation. Trends Pharmacol Sci 23: 153-155, 2002.[ISI][Medline]
  12. Fiorucci S, Distrutti E, Chiorean M, Cantucci L, Belia S, Fano G, DeGiorgio R, Stanghellini V, Corinaldesi R, and Morelli A. Nitric oxide modulates pepsinogen secretion induced by calcium-mediated agonist in guinea pig gastric chief cells. Gastroenterology 109: 1214-1223, 1995.[ISI][Medline]
  13. Fiorucci S, Distrutti E, Santucci L, and Morelli A. Leukotrienes stimulate pepsinogen secretion from guinea pig gastric chief cells by a nitric oxide-dependent pathway. Gastroenterology 108: 1709-1719, 1995.[ISI][Medline]
  14. Fiorucci S, Lanfrancone L, Cantucci L, Calabro A, Orsini B, Federici B, and Morelli A. Epidermal growth factor modulates pepsinogen secretion in guinea pig gastric chief cells. Gastroenterology 111: 945-958, 1996.[ISI][Medline]
  15. Fiorucci S and McArthur KE. Gastrin-releasing peptide directly releases pepsinogen from guinea pig chief cells. Am J Physiol Gastrointest Liver Physiol 259: G760-G766, 1990.[Abstract/Free Full Text]
  16. Fiorucci S and McArthur KE. Prostaglandin E2 desensitizes cAMP-mediated pepsinogen secretion in chief cells. Am J Physiol Gastrointest Liver Physiol 261: G858-G865, 1991.[Abstract/Free Full Text]
  17. Fiorucci S, Mencarelli A, Palazzetti B, Distrutti E, Vergnolle N, Hollenberg MD, Wallace JL, Morelli A, and Cirino G. Proteinase-activated receptor 2 is an anti-inflammatory signal for colonic lamina propria lymphocytes in a mouse model of colitis. Proc Natl Acad Sci USA 98: 13936-13941, 2001.[Abstract/Free Full Text]
  18. Gibbins IL, Furness JB, Costa M, MacIntyre I, Hillyard CJ, and Girgis S. Co-localization of calcitonin gene-related peptide-like immunoreactivity with substance P in cutaneous, vascular and visceral sensory neurons of guinea pigs. Neurosci Lett 57: 125-130, 1985.[ISI][Medline]
  19. Grynkiewicz G, Poeie M, and Tsien RY. A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260: 3440-3450, 1985.[Abstract]
  20. Kawao N, Sakaguchi Y, Tagome A, Kuroda R, Nishida S, Irimajiri K, Nishikawa H, Kawai K, Hollenberg MD, and Kawabata A. Protease-activated receptor-2 (PAR-2) in the rat gastric mucosa: immunolocalization and facilitation of pepsin/pepsinogen secretion. Br J Pharmacol 135: 1292-1296, 2002.[Abstract/Free Full Text]
  21. Kawabata A, Kinoshita M, Nishikawa H, Kuroda R, Nishida M, Araki H, Arizono N, Oda Y, and Kakehi K. The protease-activated receptor-2 agonist induces gastric mucus secretion and mucosal cytoprotection. J Clin Invest 107: 1443-1450, 2001.[Abstract/Free Full Text]
  22. Kawabata A, Kuroda R, Nagata N, Kawao N, Masuko T, Nishikawa H, and Kawai K. In vivo evidence that protease-activated receptors 1 and 2 modulate gastrointestinal transit in the mouse. Br J Pharmacol 133: 1213-1218, 2001.[Abstract/Free Full Text]
  23. Kawabata A, Nishikawa H, Kuroda R, Kawai K, and Hollenberg MD. Proteinase-activated receptor-2 (PAR-2): regulation of salivary and pancreatic exocrine secretion in vivo in rats and mice. Br J Pharmacol 129: 1808-1814, 2000.[Abstract/Free Full Text]
  24. Kitsukawa Y, Turner RJ, Pradhan TK, and Jensen RT. Gastric chief cells possess NK1 receptors which mediate pepsinogen secretion and are regulated by agents that increase cAMP and phospholipase C. Biochim Biophys Acta 1312: 105-116, 1996.[ISI][Medline]
  25. Khokhlatchev AV, Canagarajah B, Wilsbacher J, Robinson M, Atkinson M, Goldsmith E, and Cobb MH. Phosphorylation of the MAP kinase ERK2 promotes its homodimerization and nuclear translocation. Cell 93: 605-615, 1998.[ISI][Medline]
  26. Kong W, McConalogue K, Khitin LM, Hollenberg MD, Payan DG, Böhm SK, and Bunnett NW. Luminal trypsin may regulate enterocytes through proteinase-activated receptor 2. Proc Natl Acad Sci USA 94: 8884-8889, 1997.[Abstract/Free Full Text]
  27. McFarlane SR, Seatter MJ, Kanke T, Hunter GD, and Plevin R. Proteinase-activated receptors. Pharmacol Rev 53: 245-282, 2001.[Abstract/Free Full Text]
  28. Molino M, Barnathan ES, Numerof R, Clark J, Dreyer M, Cumashi A, Hoxie JA, Schechter N, Woolkalis M, and Brass LF. Interactions of mast cell tryptase with thrombin receptors and PAR-2. J Biol Chem 272: 4043-4049, 1997.[Abstract/Free Full Text]
  29. Nguyen TD, Moody MW, Steinhoff M, Okolo C, Koh DS, and Bunnett NW. Trypsin activates pancreatic duct epithelial cell ion channels through proteinase-activated receptor-2. J Clin Invest 103: 261-269, 1999.[Abstract/Free Full Text]
  30. Nystedt S, Emilsson K, Wahlestedt C, and Sundelin J. Molecular cloning of a potential proteinase activated receptor. Proc Natl Acad Sci USA 91: 9208-9212, 1994.[Abstract/Free Full Text]
  31. Nystedt S, Ramakrishnan V, and Sundelin J. The proteinase-activated receptor 2 is induced by inflammatory mediators in human endothelial cells. Comparison with the thrombin receptor. J Biol Chem 271: 14910-14915, 1996.[Abstract/Free Full Text]
  32. Raufman JP and Cosowsky L. Interactions between the calcium and adenylate cyclase messenger systems in dispersed chief cells from guinea pig stomach. J Biol Chem 262: 5957-5962, 1996.[Abstract/Free Full Text]
  33. Raufman JP, Sutliff VE, Kasbekar DK, Jensen RT, and Gardner JD. Pepsinogen secretion from dispersed chief cells from guinea pig stomach. Am J Physiol Gastrointest Liver Physiol 247: G95-G104, 1984.[Abstract/Free Full Text]
  34. Steinhoff M, Vergnolle N, Young SH, Tognetto M, Amadesi S, Ennes HS, Trevisani M, Hollenberg MD, Wallace JL, Caughey GH, Mitchell SE, Williams LM, Geppetti P, Mayer EA, and Bunnett NW. Agonists of proteinase-activated receptor 2 induce inflammation by a neurogenic mechanism. Nat Med 6: 151-158, 2000.[ISI][Medline]
  35. Sutliff VE, Cherner JA, Jensen RT, and Gardner JD. Binding of 125I-CCK-8 and 125I-gastrin-I to dispersed chief cells from guinea-pig stomach. Biochim Biophys Acta 1052: 9-16, 1990.[ISI][Medline]
  36. Sutliff VE, Rattan S, Gardner JD, and Jensen RT. Characterization of cholinergic receptors mediating pepsinogen secretion from chief cells. Am J Physiol Gastrointest Liver Physiol 257: G226-G234, 1989.[Abstract/Free Full Text]
  37. Vergnolle N, Bunnett NW, Sharkey KA, Brussee V, Compton SJ, Grady EF, Cirino G, Gerard N, Basbaum AI, Andrade-Gordon P, Hollenberg MD, and Wallace JL. Proteinase-activated receptor-2 and hyperalgesia: a novel pain pathway. Nat Med 7: 821-6, 2001.[ISI][Medline]
  38. Vergnolle N, MacNaughton WK, Al-Ani B, Saifeddine M, Wallace JL, and Hollenberg MD. Proteinase-activated receptor 2 (PAR-2)-activating peptides: identification of a receptor distinct from PAR-2 that regulates intestinal transport. Proc Natl Acad Sci USA 95: 7766-7771, 1998.[Abstract/Free Full Text]
  39. Vu TK, Hung DT, Wheaton VI, and Coughlin SR. Molecular cloning of a functional thrombin receptor reveals a novel proteolytic mechanism of receptor activation. Cell 64: 1057-1068, 1991.[ISI][Medline]




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