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
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ABSTRACT
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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
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.
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MATERIALS AND METHODS
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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
-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
-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
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.
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RESULTS
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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.
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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.
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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
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.
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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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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.
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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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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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|
View this table:
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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
|
---|
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
q/11 and
phospholipase C
, 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,
-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
|
---|
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.
 |
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