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GASTROINTESTINAL, HEPATIC, PULMONARY, AND RENAL
Division of Physiology and Pathophysiology (N.K., M.N., K.N., S.K., F.S., A.K.) and Division of Biological Chemistry (T.W., S.I.), School of Pharmaceutical Sciences, Kinki University, Higashi-Osaka, Japan; Department of Pharmacology and Therapeutics (M.D.H.) and Department of Physiology and Biophysics (K.C., W.K.M.), Faculty of Medicine, University of Calgary, Calgary, Alberta, Canada; and Research and Development Centre, Fuso Pharmaceutical Industries Ltd., Osaka, Japan (H.N.)
Received May 13, 2005; accepted August 23, 2005.
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; Ossovskaya and Bunnett, 2004). Although PAR1, PAR3, and PAR4 are thrombin receptors, PAR2 cannot be activated by thrombin but instead by trypsin, tryptase, coagulation factors VIIa and Xa, activated neutrophil-derived proteinase 3, membrane-bound membrane-type serine proteinase-1, and acrosin (Sekiguchi and Kawabata, 2004). PAR2 is extensively but unevenly distributed in the mammalian body, including the nervous, circulatory, gastrointestinal, and respiratory systems, modulating a variety of functions under physiological and/or pathological conditions (Cocks et al., 1999; Cicala et al., 2001a; Kawabata et al., 2001a,b, 2004; Vergnolle et al., 2001; Ossovskaya and Bunnett, 2004; Sekiguchi and Kawabata, 2004a,b; Kawabata and Kawao, 2005). PAR2, like other PARs, couples to Gq/11 protein that mediates activation of phospholipase C
followed by formation of inositol triphosphate and diacylglycerol and also seems to trigger activation of other multiple signaling pathways, including the mitogen-activated protein kinase (MAPK) cascades in distinct cell types (Ossovskaya and Bunnett, 2004; Sekiguchi and Kawabata, 2004). Recent data suggest that PAR2 activation can cause trans-activation of the epidermal growth factor (EGF) receptor via a matrix metalloproteinase (MMP)-dependent release of transforming growth factor-
(TGF-) (Darmoul et al., 2004).
In the respiratory system, PAR2 plays a dual role, being both anti-inflammatory/protective and proinflammatory (Cocks et al., 1999; Cicala et al., 2001b; Moffatt et al., 2002; Schmidlin et al., 2002; Kawabata and Kawao, 2005). PAR2 is expressed in airway epithelial cells, and stimulation of PAR2 with PAR2-activating peptides or trypsin causes epithelial prostaglandin E2 (PGE2)-dependent relaxation in the isolated tracheal and bronchial tissues from humans, guinea pigs, rats, and mice (Cocks et al., 1999; Lan et al., 2001). The involvement of PAR2 in agonist-evoked airway relaxation has been substantiated by the experiments employing PAR2-knock-out mice (Kawabata et al., 2004a). PAR2 is also expressed in human bronchial (BEAS-2B) and alveolar (A549) epithelial cell lines. Stimulation of these cells with PAR2 agonists for 24 h causes release and/or production of PGE2 and interleukins 6 and 8 (Asokananthan et al., 2002). In cultured alveolar A549 cells, PAR2 stimulation also elicits the release and/or expression of granulocyte macrophage colony stimulating factor (Sun et al., 2001; Vliagoftis et al., 2001) and MMP-9 (Vliagoftis et al., 2000). Thus, PAR2 upon activation seems to trigger or promote the release and/or production of various proinflammatory mediators in human lung epithelial cell lines. Nevertheless, the detailed signal transduction mechanisms responsible for the PAR2-mediated release/production of the mediators in those cells have yet to be studied in any depth. In this context, the present study aimed at identifying the intracellular signal transduction pathways underlying PAR2-mediated PGE2 formation in A549 cells.
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Materials and Methods |
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Cell Culture and Determination of PGE2 Formation. Cells from the human pulmonary type II-like adenocarcinoma cell line A549 were cultured in Dulbecco's modified Eagle's medium (Sigma) supplemented with 10% heat-inactivated fetal bovine serum (Sigma or Thermo Electron Corporation, Waltham, MA) and antibiotics. The cells were grown to 90% confluence in a CO2 incubator maintained at 5% CO2 and 37°C. A549 cells (1.5 x 105 cells/well) were then placed and grown in the above medium for 24 h in each well (33 mm in diameter) of a 6-well culture plate and then cultured in the serum-free medium overnight. After another change of the medium with a fresh serum-free medium (1 ml), the cells were incubated for 1 to 1.5 h and then stimulated with SLIGRL-NH2, a PAR2-activating peptide, at 0.1 to 300 µM; LSIGRL-NH2, a PAR2-inactive control peptide, at 300 µM; TFLLR-NH2, a PAR1-activating peptide, at 300 µM; and thrombin, an endogenous agonist proteinase for PAR1, PAR3, and PAR4, at 100 nM. Samples in a volume of 6 µl were repeatedly collected from the supernatant of the culture medium after stimulation for 0, 0.5, 3, and 18 h. In inhibition experiments, various inhibitors/chemicals or vehicle were added to the culture medium 0.5 h (0.5 or 18 h for GM6001; 24 h for doxycycline) before stimulation. The collected samples were diluted 10 times, and the amount of PGE2 in the diluted samples was determined using an enzyme immunoassay kit (Cayman Chemical). In brief, 96-well plates coated with the goat anti-mouse IgG antibody were loaded with 50 µl of samples or standards, 50 µl of PGE2-linked acetylcholinesterase tracer, and 50 µl of the mouse anti-PGE2 monoclonal antibody and incubated for 18 h at 4°C. After five washes, 200 µl of Ellman's reagent was added into each well. After 90-min incubation for development of color, the optical density at a wavelength of 405 nm was determined. The amount of PGE2 formed and released in response to stimuli was calculated by subtracting the basal value at time 0 from the value at each time point.
Assay of Cytosolic Ca2+ Mobilization. A549 cells (1.5 x 105 cells) were seeded and grown for 24 h on four round glass coverslips (13.2 mm in diameter) coated with collagen (Cellmatrix Type I-A; Nitta Gelatin Inc., Osaka, Japan) in a tissue culture dish (35 mm in diameter) containing the above-mentioned standard medium. After additional 24-h incubation in the serum-free medium, the cells were loaded with 10 µM of Fura 2-AM (Dojindo, Kumamoto, Japan) for 1 h at room temperature in a HEPES buffer of the following composition: 150 mM NaCl, 3 mM KCl, 1.5 mM CaCl2, 1.0 mM MgCl2, 20 mM HEPES, and 10 mM D-glucose. The cells were then washed with a fresh HEPES buffer, and the glass coverslip with the cells was set on a holder. The holder was then mounted in a quartz cuvette containing 2 ml of a gassed (95% O2/5% CO2) HEPES buffer, which was placed in an Intracellular Ion Analyzer (CAF-110; Japan Spectroscopic Co., Tokyo, Japan) and maintained at 25°C with constant stirring throughout the experiment. The cytosolic Ca2+ level was measured as a ratio of fluorescence intensity at an emission wavelength of 500 nm with excitation at wavelengths of 340 and 380 nm applied alternately at a frequency of 128 Hz (F340/F380). The data were recorded on a computer at an acquisition rate of 100 Hz using a PowerLab Recording System (AD Instruments, New South Wales, Australia). The maximal increase in cytosolic Ca2+ levels was observed at the end of each experiment by adding 20 µM ionomycin. Changes in cytosolic Ca2+ levels caused by various stimuli are expressed as the percentage of the response to ionomycin at 20 µM.
Detection of Phosphorylation of ERK, p38 MAPK, and EGF Receptors by Western Blot Analysis. A549 cells grown in a 6-well plate, as described above in the PGE2 formation assay, were stimulated with the PAR2-activating peptide SLIGRL-NH2 at 0.01 to 100 µM for 5, 15, 30, or 120 min in the serum-free medium. Inhibitors were added 30 min before agonist challenge. After aspirating the medium at scheduled time points, the cells were rinsed with Ca2+-/Mg2+-free phosphate-buffered saline (PBS), and lysed in 100 µl of a lysis buffer containing 2% SDS, 62.5 mM Tris-HCl, and 10% glycerol (pH 6.8). The cell lysate was harvested with a cell scraper and exposed to freeze-thawing. The samples were then sonicated for 10 s and denatured by heating at 95°C for 5 min. Protein samples (10-30 µg/lane) were separated by electrophoresis on a 12.5% SDS-polyacrylamide gel (Daiichi Pure Chemicals, Tokyo, Japan) and transferred onto polyvinylidene difluoride membrane (Immobilon-P; Millipore Corporation, Billerica, MA). The membrane was blocked with a blocking solution containing 5% skim milk, 137 mM NaCl, 0.1% Tween 20, and 20 mM Tris-HCl (pH 7.6) for 1 h at room temperature. After being washed three times with Tris-buffered saline (TBS) containing 0.1% Tween 20, the membrane was incubated with the primary polyclonal antibodies at appropriate dilution with gentle agitation overnight at 4°C. The primary antibodies employed were: rabbit anti-p44/42 MAPK (dilution 1:2000), rabbit anti-phospho-p44/42 MAPK (Thr202/Tyr204) antibody (dilution 1:1000), rabbit anti-p38 MAPK antibody (dilution 1:2000), rabbit anti-phospho-p38 MAPK (Thr180/Tyr182) antibody (dilution 1:1000), rabbit anti-EGF receptor antibody (1:2000), and rabbit anti-phospho-EGF receptor (Tyr1068) antibody (1:100) (Cell Signaling, Beverly, MA). After incubation with the primary antibody, the membrane was washed three times with the above-mentioned TBS solution and then incubated with biotinylated goat anti-rabbit IgG antibody (Vector Laboratories, Burlingame, CA) as the secondary antibody for 30 min at room temperature. The membrane was washed three times with the TBS solution again, and positive bands were detected by the ABC method (Vectastain ABC kit, Vector Laboratories) followed by enhanced chemiluminescence staining (GE Healthcare, Little Chalfont, Buckinghamshire, UK).
Detection of Cyclooxygenase-2, Cyclooxygenase-1, and Microsomal PGE Synthase-1 by Western Blot and/or Immunocytochemical Analyses. A549 cells (1.2 x 106 cells) were grown in a tissue culture dish (100 mm in diameter) containing the standard medium for 24 h and then cultured in the serum-free medium for an appropriate time followed by stimulation with the PAR2 agonist SLIGRL-NH2 at 100 µM for 3 h. In inhibition experiments, various inhibitors or vehicle were added to the culture medium 30 min before agonist challenge. After 2- or 3-h agonist stimulation, the cells were washed with PBS and lysed in 200 µl of radioimmune precipitation buffer [PBS, 1% Igepal CA-630 (Sigma-Aldrich), 0.5% sodium deoxycholate, and 0.1% SDS] containing 0.1 mg/ml phenylmethylsulfonyl fluoride, 0.15 U/ml aprotinin, and 1 mM sodium orthovanadate. The cell lysate was harvested with a cell scraper and passed through a 21-gauge needle followed by 60-min incubation on ice. After centrifugation at 10,000g for 10 min at 4°C, the supernatant was heated for 3 min at 95°C. Proteins were separated by electrophoresis and transferred onto the membrane, as mentioned above. The membrane was probed for 1 h at room temperature with an affinity-purified goat polyclonal antibody raised against a peptide mapping at the carboxyl terminus of human cyclooxygenase (COX)-2 (dilution 1:100) (Santa Cruz Biotechnology, Santa Cruz, CA), a monoclonal antibody against human COX-1 (dilution 1:200) (Wako Pure Chemicals), and a purified rabbit polyclonal antibody against a peptide corresponding to amino acids 59 to 75 of human microsomal PGE synthase-1 (mPGES-1) (dilution 1:1000) (Cayman Chemical). After washing, as described above, the membrane was incubated for 45 (COX-2) or 60 min (COX-1, mPGES-1) at room temperature with a horseradish peroxidase (HRP)-conjugated anti-goat IgG antibody (Chemicon International, Temecula, CA) for COX-2, a HRP-linked anti-mouse IgG antibody (Cell Signaling) for COX-1, or an HRP-linked anti-rabbit IgG antibody (Chemicon International) for mPGES-1. Positive bands were visualized by enhanced chemiluminescence detection, as mentioned above.
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Analysis of mRNA Levels for COX-2 and COX-1 by Quantitative Real Time Polymerase Chain Reaction. A549 cells grown in a tissue culture dish (100 mm in diameter) were stimulated with SLIGRL-NH2 at 100 µM for 1 to 3 h in the absence or presence of inhibitors, as described above. After washing with PBS, the cells were lysed and harvested in the TRIzol reagent (Invitrogen, Carlsbad, CA) using a cell scraper for extraction of total RNA. Then, mRNA was reverse-transcribed with avian myeloblastosis virus-derived reverse transcriptase (Takara, Kyoto, Japan) at 42°C for 50 min. Quantitative PCR was performed with an ABI Prism 7000 Sequence Detection System (Applied Biosystems, Foster, CA) using Assay-On-DemandTM Gene Expression Products for human COX-2 and COX-1 and for eukaryotic 18S rRNA (Hs00153133_m1, Hs00377721_m1, and Hs99999901_s1, respectively; Applied Biosystems). PCR amplification proceeded for 40 cycles of incubation at 95°C for 15 s (denaturing) and at 60°C for 60 s (annealing/extending). Sequence-specific amplification was monitored as a real time increase in fluorescence signals. The sequence-specific mRNA levels for COX-2 and COX-1 were determined using appropriate standard curves followed by normalization with rRNA levels.
Detection of mRNAs for EGF Receptor Ligands in A549 Cells. The cells were collected, and the extracted total RNAs were reverse-transcribed, as described above. PCR amplification was performed using the RNA LA PCR kit (AMV), version 1.1 (Takara, Otsu, Japan) and allowed to proceed for 35 or 40 cycles (genes of EGF receptor ligands) and for 30 cycles (housekeeping gene) (a cycle: denaturation at 94°C for 30 s; reannealing at 55°C for 30 s; primer extension at 72°C for 1 min). PCR primers employed were: 5'-TCA GTT CTG CTT CCA TGG AAC C-3' (sense) and 5'-TTT CTG AGT GGC AGC AAG CG-3' (antisense), amplifying 317-bp fragments for transforming growth factor- (TGF-); 5'-ACA AGG AGG AGC ACG GGA AAA G-3' (sense) and 5'-CGA TGA CCA GCA GAC AGA CAG ATG-3' (antisense), amplifying 276-bp fragments for heparin-binding EGF (HB-EGF); and 5'-TGT GAA CTG ATC ATG TTT ATG-3' (sense) and 5'-GTC CAC CAC CCT GTT GCT GTA GCC-3' (antisense), amplifying 876-bp fragments for a housekeeping gene, glyceraldehydes-3-phosphate dehydrogenase. PCR products were verified by electrophoresis on 2% agarose gels and visualized under UV with ethidium bromide.
Cell Proliferation Assay. Cell proliferation was determined by the MTT method. In brief, A549 cells (5000 cells/well) were grown in 100 µl of the standard Dulbecco's modified Eagle's medium for 24 h using a 96-well plate and then cultured in 50 µl of the serum-free medium for 1 h before the experiment. Cells were stimulated with SLIGRL-NH2 at 100 µM for 24 h in the absence or presence of indomethacin at 10 µM, and then 10 µl of MTT solution was added to each well. After 4-h incubation at 37°C, the cells were lysed by addition of 100 µl of isopropyl alcohol containing 0.04 M HCl and left to stand at a room temperature for 18 h. Absorbance at two distinct wavelengths of 595 and 670 nm was measured. The A595 to A670 was taken as the index of cell count. The proliferation rate is shown as a percentage of the basal value before agonist stimulation.
Statistics. Data are represented as mean ± S.E.M. Statistical significance was analyzed by Student's t test for two-group data and by Tukey's test for multiple comparisons. Significance was set at a P < 0.05 level.
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Results |
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PGE2 (in picograms) was 24.2 ± 18.7 and 43.4 ± 24.7 in the cells treated with vehicle and SLIGRL-NH2 at 300 µM, respectively, for 10 min (n = 8). The stimulatory effect of SLIGRL-NH2 on the release of PGE2 was concentration-dependent in a range of 1 to 300 µM (Fig. 1C). Of particular note is that LSIGRL-NH2, a standard control PAR2-inactive scrambled peptide at 300 µM was incapable of mimicking the effect of SLIGRL-NH2 (Fig. 1C). In contrast, neither thrombin, an activator for PAR1, PAR3, and PAR4, at 0.1 µM (10 U/ml) nor TFLLR-NH2, the PAR1-activating peptide, at 300 µM caused an increase in PGE2 formation in the A549 cells.
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PGE2 (in picograms) was -33.8 ± 24.1 and 48.6 ± 24.2 in the cells treated with vehicle and SLIGRL-NH2 at 100 µM, respectively, for 30 min. Therefore, we examined the effect of the COX inhibitors on the increased PGE2 production after 30-min incubation with SLIGRL-NH2 at 300 µM and found that NS-398 significantly (P < 0.05) and strongly suppressed the PGE2 formation and that SC-560 also exhibited a tendency toward inhibition; i.e., PGE2 (in picograms) was 26.6 ± 13.1 in nonstimulated cells and 127.2 ± 26.0, 51.2 ± 32.8, and 10.1 ± 24.0 in SLIGRL-NH2 (300 µM, 30 min)-simulated cells in the presence of DMSO, SC-560, and NS-398, respectively (n = 10-11).
Effects of Various Inhibitors of Signal Transduction Pathways on PGE2 Formation after PAR2 Stimulation in A549 Cells. Because PAR2 stimulation is known to activate multiple MAPK pathways, including phosphorylation of ERK (Macfarlane et al., 2001; Hollenberg and Compton, 2002; Ossovskaya and Bunnett, 2004), we first evaluated involvement of MAPK pathways in the PAR2-mediated PGE2 formation. Two selective inhibitors of MAPK/ERK kinase (MEK), U0126 at 10 µM and PD98059 at 50 µM, blocked 100 µM SLIGRL-NH2-induced PGE2 production (Fig. 3A). Likewise, SB203580, an inhibitor of p38 MAPK, at 1 to 10 µM markedly reduced the PAR2-mediated increase in PGE2 production (Fig. 3A). The formation of PGE2 after PAR2 stimulation was also significantly reduced by genistein (30 µM), a nonselective inhibitor of tyrosine kinases, and PP2 (1 µM), a selective inhibitor of the Src family tyrosine kinase (Fig. 3B). We then examined whether trans-activation of the EGF receptor kinase (EGFRK) possibly via a MMP-mediated mechanism might contribute to PGE2 production caused by PAR2 stimulation. The involvement of the EGFRK itself was evident from the ability of the potent and selective EGFRK inhibitor, PD153035, to block the SLIGRL-NH2-evoked formation of PGE2 (Fig. 4). Surprisingly, however, GM6001, an effective inhibitor of MMPs, even at a supramaximal inhibitory concentration of 25 µM (0.5-h or 18-h preincubation), did not affect PAR2-stimulated PGE2 production (Fig. 4). Likewise, doxycycline, a nonselective inhibitor of MMP production, at 25 µM (24-h preincubation) did not show significant inhibition (Fig. 4). The PAR2-mediated increase in PGE2 production was also abolished by BAPTA/AM (30 µM), an intracellular Ca2+ chelator, and was partially blocked by GF109203X (1 µM), a protein kinase C inhibitor (Fig. 5). In contrast, wortmannin (0.1 µM), an inhibitor of phosphatidylinositol 3-kinase (PI3-kinase), failed to suppress PGE2 release because of PAR2 activation (Fig. 5). Collectively, the data indicated that PAR2-mediated PGE2 formation involves an elevation of intracellular calcium, both the MEK-ERK and p38 MAPK pathways, Src family kinase, an MMP-independent trans-activation of the EGFRK, and to a certain extent, protein kinase C.
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Phosphorylation of ERK and p38 MAPK by PAR2 Stimulation. The PAR2-activating peptide SLIGRL-NH2 at 100 µM caused a prompt activation of both ERK1 and ERK2 at 5 min. The activation, inferred from the increased phosphorylated forms of ERK1/2, was transient, decreasing to a lower but persistent level over a 30-min time period (Fig. 6A). The SLIGRL-NH2-evoked ERK1/2 phosphorylation at 5 min was concentration-dependent over a wide concentration range, 0.001 to 100 µM (Fig. 6B). The MEK inhibitor PD98059 at 50 µM abolished ERK1/2 phosphorylation triggered by PAR2 stimulation with SLIGRL-NH2 at 100 µM for 5 min and also reduced the constitutive phosphorylation of ERK1/2 observed in the absence of PAR2 activation (Fig. 6C). Neither GF109203X (1 µM), a PKC inhibitor, nor genistein (30 µM), a nonselective tyrosine kinase inhibitor, reduced the activation of ERK1/2 caused by SLIGRL-NH2 at 100 or 10 µM (Fig. 6, D and E). However, these concentrations of either GF109203X or genistein clearly attenuated ERK1/2 phosphorylation caused by a low concentration (0.1 µM) of SLIGRL-NH2 (Fig. 6, D and E). In contrast, PP2, a Src inhibitor, at 1 µM produced no inhibition on ERK activation by SLIGRL-NH2 at 0.1 µM (Fig. 6F). The selective and potent EGFRK inhibitor PD153035 at 1 µM failed to suppress ERK activation caused by PAR2 activation with SLIGRL-NH2 at either high (100 µM) or low (0.1 µM) concentrations (Fig. 6G). Taken together, protein kinase C and a non-Src tyrosine kinase would seem to be involved in part in the prompt activation of ERK after PAR2 activation with SLIGRL-NH2 at low but not at high concentrations, whereas trans-activation of the EGFRK does not seem to be upstream of PAR2-mediated ERK activation.
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Increased phosphorylation of p38 MAPK was observed after PAR2 stimulation with SLIGRL-NH2 at 100 µM with a delayed time course (15-30 min), relative to ERK1/2 (increased at 5 min), that persisted for 2-h stimulation (Fig. 7A). The phosphorylation of p38 MAPK caused by SLIGRL-NH2 was blocked by SB203580 (Fig. 7B), an inhibitor that is known to block the enzymatic activity of p38 MAPK, possibly concurrently altering its conformation in order to not to be efficiently phosphorylated by its upstream activating kinase. Both PP2 and PD153035 also attenuated SLIGRL-NH2-induced phosphorylation of p38 MAPK (Fig. 7B), implying that activation of Src and EGF receptors is upstream of p38 MAPK activation. Of note is that both the early and late phases of p38 MAPK phosphorylation caused by PAR2 stimulation for 15 min and 2 h, respectively, were not affected by either the COX-1 inhibitor SC-560 or the COX-2 inhibitor NS-398 (Fig. 7C), indicating that cyclooxygenases products themselves do not contribute to either the prompt or delayed activation of p38 MAPK.
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Phosphorylation of EGF Receptors by PAR2 Stimulation and Expression of mRNAs for EGF Receptor Ligands in A549 Cells. We then examined whether PAR2 stimulation could cause trans-phosphorylation/activation of EGF receptors in A549 cells. The PAR2-activating peptide SLIGRL-NH2 at 100 µM produced a rapid phosphorylation of the EGFRK at 5 and 15 min, but not at 30 min (Fig. 8A). EGF receptor activation by PAR2 stimulation thus occurred more rapidly than p38 MAPK activation (see Fig. 7A). Interestingly, the SLIGRL-NH2-evoked rapid phosphorylation of EGF receptors was attenuated by the Src inhibitor PP2 (Fig. 8B), implying a role for Src in the trans-phosphorylation/activation of EGF receptors. In reverse transcription-PCR analyses of mRNAs for EGF receptor ligands, TGF- and HB-EGF employing appropriate primers that had been confirmed to work in other human cells, HB-EGF mRNA, was detected after PCR amplification of 40 cycles, but not 35 cycles, in A549 cells, although mRNA for TGF- was not detectable even after 40-cycle amplification (Fig. 8C).
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Expression of COX Isoforms after PAR2 Stimulation in A549 Cells. The cellular content of the COX-1 and COX-2 enzymes was documented by monitoring both the increase in intensity of staining in individual cells and by estimating by morphometric analysis, the proportion of COX-expressing cells (% COX-positive) counted in multiple microscopic fields. A549 cells expressed a low degree of immunoreactive COX-1 in the cytosol and/or adjacent organelles in approximately 80% of the cells (Fig. 10A). PAR2 stimulation with SLIGRL-NH2 at 100 µM for 3 h did not significantly increase the proportion of cells in which immunoreactive COX-1 was detected (Fig. 10A). Immunoreactive COX-2 was barely detectable in nonstimulated cells (Fig. 9, A and B). However, upon stimulation with SLIGRL-NH2 for 3 h, there was a dramatic increase not only in the intensity of COX-2 staining in individual cells (Fig. 9, A and B) but also in the proportion of cells expressing COX-2 (Fig. 10A). The Western blot analysis also confirmed that COX-2 protein was up-regulated by PAR2 activation with SLIGRL-NH2 for 3 h, whereas COX-1 protein did not considerably increase after PAR2 stimulation (Fig. 10B).
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30%) attenuated the SLIGRL-NH2-evoked increase in the COX-2-positive cell number (data not shown). In summary, COX-2 up-regulation after PAR2 stimulation involves activation of p38 MAPK, EGF receptors and Src, and also non-PGE2 prostanoids that are produced via COX-2 itself.
Next, we examined whether PAR2 stimulation could induce up-regulation of COX-2 mRNA in A549 cells. EGF, known to cause COX-2 induction (Pawliczak et al., 2004), at 50 nM caused a significant increase in COX mRNA expression at 3 h. Likewise, SLIGRL-NH2 at 100 µM caused a significant increase in COX-2 mRNA levels at 2 and 3 h (Fig. 11B). PAR2 activation also caused a small but significant increase in COX-1 mRNA levels at 1 and 2 h in A549 cells (Fig. 11A). The increased COX-2 mRNA levels were in keeping with the up-regulation of COX-2 protein levels (see Figs. 9 and 10, A and B). The enhancement of COX-2 mRNA levels after PAR2 activation was blocked by the EGFRK inhibitor PD153035 (Fig. 11C).
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). We also found that PAR2 stimulation for 2 h (peak time) could cause up-regulation of mPGES-1 in A549 cells (Fig. 12). Most interestingly, the COX-1 inhibitor SC-560, but not the COX-2 inhibitor NS-398, strongly suppressed the PAR2-triggered mPGES-1 induction (Fig. 12).
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; Hung et al., 2000), we tested whether PAR2 activation could facilitate proliferation of A549 cells and determined the possible involvement of endogenous prostanoids. PAR2 stimulation with SLIGRL-NH2 at 100 µM for 24 h significantly (P < 0.05) increased the proliferation rate in A549 cells, an effect that was resistant to pretreatment with indomethacin, a nonselective COX inhibitor. The proliferation rates (in percentage) in the absence and presence of indomethacin were 187.2 ± 9.4 and 205.8 ± 10.0 in nonstimulated cells and 233.2 ± 8.0 and 237.0 ± 14.2 in SLIGRL-NH2-stimulated cells, respectively (n = 5). |
Discussion |
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), but that study did not examine the early time course of PGE2 production (only a 24-h time point was evaluated) and did not assess in any detail the signal transduction pathways that might be involved. We found that at early time points (up to 3 h) neither the PAR1-activating peptide TFLLR-NH2 nor thrombin caused an increase in PGE2 production (Fig. 1D), whereas activation of PAR2 by SLIGRL-NH2 (but not the PAR2 standard inactive peptide, LSIGRL-NH2) caused a substantial increase (Fig. 1C). These distinct effects of activating PAR1 versus PAR2 were mirrored by the calcium signaling assay wherein PAR1 activation (TFLLR-NH2 stimulus) elicited only a slight Ca2+ signal compared with PAR2 (SLIGRL-NH2 stimulus; Fig. 1A).
Our data obtained using a variety of signal pathway inhibitors indicate that, for PAR2-stimulated increases in A549 cell PGE2 production, there is an involvement of cytosolic Ca2+ mobilization, cPLA2, COX-1, COX-2, both MEK-ERK and p38 MAPK pathways, Src, protein kinase C, and an MMP-independent trans-activation of the EGFRK. MEK-ERK activation would seem to be key for the increased enzymatic formation of PGE2, whereas p38 MAPK (but not MEK-ERK) is essential for the up-regulation of COX-2 enzyme. A trans-activation of the EGFRK, leading to p38 MAPK activation, is also implicated in the up-regulation of COX-2 mRNA. Table 1 summarizes the effects of the various inhibitors on: 1) the PAR2-triggered increase of PGE2 production and 2) the up-regulation of COX-2 enzyme. Taken together with COX-1-dependent induction of mPGES-1, the hypothetical signal transduction mechanisms involved in this process, based on our results, are shown in Fig. 13.
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Activation of PAR2 in alveolar type II-derived A549 cells triggered multiple signaling pathways, including protein kinase C and non-Src genistein-sensitive tyrosine kinases that are considered to be upstream of activation of the MEK-ERK pathway (see Fig. 6, D, E, and F). It is well established that mobilization of cytosolic Ca2+ and a direct phosphorylation of cPLA2 by activated ERK can cause trans-location of cPLA2 from the cytosol to the membrane, thereby triggering arachidonic acid release (Wissing et al., 1997). In turn, the released arachidonic acid can be rapidly metabolized by cyclooxygenases to yield prostaglandins. cPLA2 is also known to be a target for phosphorylation by p38 MAPK (Kramer et al., 1996). In keeping with the proposed mechanisms for the activation of cPLA2, we found that preventing an elevation of cytosolic calcium with BAPTA and blocking MAPK activation with the MEK-MAPK inhibitors PD98059 and U0126 abrogated the PAR2-mediated increases in PGE2. It is of importance in this regard that two chemically distinct MEK inhibitors were effective in blocking PGE2 production (see Figs. 3A and 5), because PD98059, but not U0126, might potentially have blocked PGE2 formation by inhibiting cyclooxygenase activity per se, rather than by blocking MEK (Borsch-Haubold et al., 1998). Likewise, a concentration of SB203580 lower than that required for maximal cyclooxygenase inhibition was able to block the PGE2 production (Fig. 3A) and the up-regulation of COX-2 itself (Figs. 9 and 10), implicating p38 MAPK in this process, rather than an inhibition of COX-2 itself. As described previously (Croxtal et al., 1996), a role for protein kinase C in the release of PGE2 from A549 cells via activation of cPLA2 was also implicated in view of the ability of GF109203X to block PAR2-stimulated A549 cell ERK activation/phosphorylation at lower SLIGRL-NH2 concentrations. Although the EGFRK inhibitor PD153035 blocked both the increase in PGE2 and the increase in COX-2 protein, activation of EGF receptors per se was found not to be upstream of the MEK-ERK-cPLA2 pathway, because ERK phosphorylation after stimulation with SLIGRL-NH2 was unaffected by the EGFRK inhibitor (see Fig. 6G). Rather, trans-activation of the EGFRK was upstream of p38 MAPK activation (see below).
We found unexpectedly that both COX-1 and COX-2 isoforms seem to contribute synergistically to the PAR2-triggered up-regulation of PGE2 formation, because either the COX-1 inhibitor, SC-560, or the selective COX-2-inhibitor, NS-398, employed at appropriately selective concentrations (Miralpeix et al., 1997; Kato et al., 2001) inhibited the increase in PGE2 production caused by PAR2 activation (see Fig. 2). In this regard, it was striking that COX-2, but not COX-1, activity was essential for the up-regulation of COX-2 protein itself (Fig. 10, C and F), implying a role for small amounts of COX-2 prostanoid products, potentially activating A549 EP3 and EP4 receptors in this process (Yano et al., 2002). In contrast, the up-regulation of mPGES-1 by PAR2 stimulation is attributable to possibly non-PGE2 prostanoids produced through COX-1 but not COX-2 (see Fig. 12). Of note is that both COX-1 and COX-2 have been reported to be involved together in other biological events after activation of PARs (Buresi et al., 2002; Kawabata et al., 2004).
The PAR2-triggered up-regulation of COX-2 at the mRNA and protein levels was linked directly to the activation of p38 MAPK downstream of the trans-activation of the EGFRK (see Figs. 7B; 9, A and B; 10, C, D, and E; and 11C). This up-regulation of COX-2 protein was not dependent upon MAPK activation. In this regard, the activation/phosphorylation of EGF receptors temporally preceded the phosphorylation/activation of p38 MAPK (see Figs. 7A and 8A). However, the activation of p38 MAPK was not linked to a potential action of COX metabolic products via EP receptors, because COX inhibitors attenuated neither the early (15 min) nor the delayed (2 h) phosphorylation/activation of p38 MAPK after PAR2 activation (see Fig. 7C). It is an open question whether activation of protein kinase A via EP4 receptors (Yano et al., 2002) might be involved in the positive feedback regulation of COX-2 by PGE2 formed in A549 cells stimulated with a PAR2 agonist.
It is now established that a number of G protein-coupled receptors, including PAR2, are capable of trans-activating EGF receptors through a metalloproteinase-dependent release of EGF receptor-activating ligands (Prenzel et al., 1999; Pai et al., 2002; Darmoul et al., 2004). In this context, it was important to note that the EGF receptor tyrosine kinase inhibitor PD153035, but not the MMP inhibitor GM6001, blocked PGE2 formation after PAR2 activation (see Fig. 4). Thus, a MMP-induced trans-activation of EGF receptors was not involved. It is unlikely that PAR2 stimulation would have caused an activation of EGF receptors through transcriptional up-regulation of EGF receptor ligands such as EGF, HB-EGF, and TGF- because of the rapid (5 min) phosphorylation-activation of the EGFRK caused by PAR2 activation (see Fig. 8). Rather, our inhibition experiments using the Src family tyrosine kinase inhibitor PP2 (see Figs. 7B, 8B, and 10, C and G) imply that activation of Src directly or indirectly contributes to the trans-activation of the EGFRK. This trans-activation would be followed by the downstream activation of p38 MAPK and a subsequent up-regulation of COX-2. The precise mechanisms other than a Src kinase link responsible for the cross-talk between PAR2 and the EGFRK remain to be investigated.
The physiological significance of PAR2-triggered PGE2 formation in A549 cells remains an open question. In isolated airway preparations, stimulation of epithelial PAR2 produces bronchial and tracheal relaxation within minutes via PGE2 release (Cocks et al., 1999; Lan et al., 2001; Kawabata et al., 2004). However, in the A549 cell line representing alveolar type-II epithelial cells, the increase in PGE2 release after PAR2 stimulation took place over 1 to 3 h, involving a coordinated activation of cPLA2 and an up-regulation of COX-2 protein. In this context, our present data reflect the delayed up-regulation of this enzyme associated with inflammatory processes. In this regard, PAR2 can be seen to play both proinflammatory and anti-inflammatory roles in cells of the respiratory system (Cicala et al., 2001b; Moffatt et al., 2002; Schmidlin et al., 2002; Kawabata and Kawao, 2005).
PAR2 stimulation led to a proliferative response of the A549 cells. However, this effect was distinct from the ability of PAR2 activation to increase PGE2 production in terms of the lack of effect of indomethacin on the proliferative response. Of note is that PGE2 may participate in angiogenesis that is critical for wound healing and cancer growth/metastasis (Pai et al., 2001; Salcedo et al., 2003).
In summary, as outlined in Fig. 13, our data indicate that PAR2 activation via a coordinated and distinct activation of both ERK and p38 MAPK leads to an increase in PGE2 production involving cPLA2 activation and an up-regulation of COX-2 and mPGES-1. This process is linked to the production of both COX-1 and COX-2 metabolites and is driven by an increase in cytosolic Ca2+, activation of a non-Src genistein-sensitive tyrosine kinase, protein kinase C, and a Src-mediated trans-activation of the EGFRK. The activation of the EGFRK in turn activates p38 MAPK to up-regulate COX-2 mRNA and protein.
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Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
ABBREVIATIONS: PAR, proteinase-activated receptor; MAPK, mitogen-activated protein kinase; EGF, epidermal growth factor; MMP, matrix metalloproteinase; TGF-, transforming growth factor-; PGE2, prostaglandin E2; mPGES-1, microsomal prostaglandin E synthase-1; ERK, extracellular signal-regulated kinase; EGFRK, EGF receptor tyrosine kinase; PBS, phosphate-buffered saline; TBS, Tris-buffered saline; COX, cyclooxygenase; HRP, horseradish peroxidase; PCR, polymerase chain reaction; HB-EGF; heparin-binding EGF; Igepal CA-630, (octylphenoxy)-polyethoxyethanol; iPLA2, Ca2+-independent phospholipase A2; cPLA2, cytosolic Ca2+-dependent phospholipase A2; BEL, bromoenol lactone; PI3-kinase, phosphatidylinositol 3-kinase; AACOCF3, arachidonyl trifluoromethyl ketone; MEK, MAPK/ERK kinase; DMSO, dimethyl sulfoxide; SLIGRL-NH2, Ser-Leu-Ile-Gly-Arg-Leu-amide; TFLLR-NH2, Thr-Phe-Leu-Leu-Arg-amide; U0126, 1,4-diamino-2,3-dicyano-1,4-bis[2-aminophenylthio]butadiene; SC-560, 5-(4-chlorophenyl)-1-(4-methoxyphenyl)-3-trifluoromethylpyrazole); NS-398, N-(2-cyclohexyloxy-4-nitrophenyl)methanesulfonamide); PD98059, 2-(2'-amino-3'-methoxyphenyl)-oxanaphthalen-4-one); SB203580, 4-(4-fluorophenyl)-2-(4-methylsulfonylphenyl)-5(4-pyridyl)imidazole); GF109203X, 3-(1-(3-(dimethylamino)propyl)-1H-indol-3-yl)-4-(1H-indol-3-yl)-1H-pyrrole-2,5-dione monohydrochloride; PP2 4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine; PD153035, 4-(3-bromoanilino)-6,7-dimethoxyquinazoline; GM6001, N-[2(R)-2-(hydroxamido carbonylmethyl)-4-methylpentanoyl]-L-tryptophane methylamide; BAPTA-AM, 1,2-bis(2-aminophenoxyethane)-N,N,N',N'-tetraacetic acid/acetomethoxy ester; MTT, 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide.
Address correspondence to: Dr. Atsufumi Kawabata, Division of Physiology and Pathophysiology, School of Pharmaceutical Sciences, Kinki University, 3-4-1 Kowakae, Higashi-Osaka 577-8502, Japan. E-mail: kawabata{at}phar.kindai.ac.jp
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