PAR1-dependent and independent increases in COX-2 and PGE2 in human colonic myofibroblasts stimulated by thrombin

Michelle L. Seymour, Nosheen F. Zaidi, Morley D. Hollenberg, and Wallace K. MacNaughton

Mucosal Inflammation Research Group, University of Calgary, Calgary, Alberta, Canada T2N 4N1


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Subepithelial myofibroblast-derived prostaglandin E2 (PGE2) regulates epithelial chloride secretion in the intestine. Thrombin is elevated in inflammatory conditions of the bowel. Therefore, we sought to determine a role for thrombin in regulating PGE2 synthesis by colonic myofibroblasts. Incubation of cultured CCD-18Co colonic myofibroblasts with thrombin, the proteinase-activated receptor 1 (PAR1)-activating peptide (Cit-NH2), and peptides corresponding to 2 noncatalytic regions of thrombin (TP367 and TP508) for 18 h increased both cyclooxygenase (COX)-2 expression (immunocytochemistry) and PGE2 synthesis (enzyme immunoassay). Inhibition of thrombin by D-Phe-Pro-Arg-chloromethylketone (PPACK) did not significantly reduce PGE2 synthesis, which remained elevated compared with control. We also investigated the basic fibroblast growth factor (bFGF) dependence of thrombin-induced PGE2 elevations. Recombinant human bFGF concentration dependently increased PGE2 synthesis, and a bFGF neutralizing antibody inhibited PGE2 synthesis induced by TP367 and TP508 (~40%) and by thrombin (~20%) (but not Cit-NH2). Thrombin, therefore, upregulates COX-2-derived PGE2 synthesis by both catalytic cleavage of PAR1 and bFGF-dependent noncatalytic activity. This presents a novel mechanism by which intestinal myofibroblasts might regulate epithelial chloride secretion.

cyclooxygenase; proteinase-activated receptor 1; prostaglandin E2


    INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

MYOFIBROBLASTS ARE LOCATED in the lamina propria of the intestine in close proximity to epithelial cells and have ultrastructural features of both smooth muscle cells and fibroblasts (38). They are paracrine cells that regulate a number of fundamental biological processes in the intestine, including epithelial chloride secretion (11), which, in turn, controls the secretion of water into the intestinal lumen. Coculture studies indicate that the myofibroblast regulates epithelial cell chloride secretion by producing prostaglandin E2 (PGE2) (11) via cyclooxygenase (COX)-2 (25). In intestinal tissue, this regulation represents a rapidly activated host defense mechanism where PGE2-induced chloride secretion leads to water secretion that flushes away invading bacteria (10).

Epithelial chloride secretion can be induced by thrombin via the direct activation of proteinase-activated receptor 1 (PAR1) on intestinal epithelial cells (16). Thrombin has also been shown to increase both COX-2 expression and PGE2 release in a hamster fibroblast cell line by proteolytic cleavage of PAR1 (20). Therefore, because myofibroblast-derived PGE2 is important in regulating epithelial chloride secretion (11), the ability of thrombin to upregulate PGE2 synthesis in lamina propria myofibroblasts represents a mechanism by which thrombin might further amplify epithelial secretory responses on a longer term basis and contribute to intestinal host defense. An increase in the generation of thrombin from prothrombin has been associated with inflammation in both Crohn's disease (18) and ulcerative colitis (35).

Aside from its classic role in the cleavage of fibrinogen to active fibrin in the clotting cascade, thrombin has the ability to regulate a number of cellular functions (19), such as proliferation (5, 8, 17, 23, 26), platelet aggregation (40), and endothelial cell gene upregulation (21). These actions of thrombin are mediated in a number of ways. The first and best characterized is via the cleavage of the amino terminus of PAR1 to reveal a tethered ligand with the ability to interact with and activate the main body of the receptor (32, 40). The effect of this tethered ligand in the activation of PARs can be mimicked by short peptides corresponding to, or relating to, the amino acid sequence of the tethered ligand.

The second and third sites possessing biological activity are regions of thrombin that are independent of its catalytic activity and that regulate cellular functions, such as monocyte chemotaxis (6, 7), cell adhesion (4, 9), wound healing (36), and proliferation of fibroblasts (17, 26), macrophages (8), and endothelial cells (24). Residues 367-380 of human prothrombin that contain the "loop B" insertion sequence may be mimicked by the thrombin-derived peptide TP367 (YPPWNKNFTENDL) (8, 26) and residues 508-530 of human prothrombin containing an arginine-glycine-aspartic acid (RGD) sequence common to adhesion molecules such as fibronectin and vitronectin, which may be mimicked by the thrombin-derived peptide TP508 (AGYKPDEGKRGDACEGDSGGPFV) (9, 26).

A synergistic relationship between PAR activation and some of the noncatalytic effects of thrombin has been established (17, 26), and, as such, some cellular effects may be the result of multiple signals from thrombin. The relative contributions of the different active regions of the thrombin molecule to intestinal function are not known.

A number of reports suggest that thrombin causes its cellular effects in part by inducing the autocrine release of basic fibroblast growth factor (bFGF) (1, 24, 39) and that this thrombin-induced bFGF release might be independent of thrombin's catalytic activity (24, 39). We therefore hypothesized that in colonic myofibroblasts, thrombin might cause an upregulation of COX-2 and subsequent PGE2 release, both via the catalytic cleavage of PAR1 and via the noncatalytic activity of thrombin, and that this upregulation might be mediated, in part, by bFGF.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
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REFERENCES

Cell lines and culture methods. The myofibroblast-like cell line CCD-18Co, derived from a colonic mucosal biopsy of a 2-mo-old human female, was obtained from the American Type Culture Collection (ATCC, Manassas, VA; no. CRL-1459). Cells were grown in minimum essential medium (Eagle) (MEM) (ATCC) supplemented with 10% fetal bovine serum (Medicorp, Montreal, QC, Canada) and 1% penicillin/streptomycin (Sigma, Oakville, ON, Canada). Cells were subcultured at a 1:2 split ratio following dissociation with 1.5 × trypsin-ETDA (Sigma).

CCD-18Co were incubated for the given times at the stated concentrations with thrombin (human plasma, 2,974 NIH U/mg protein) (Calbiochem), bFGF (Upstate Biotechnology, Lake Placid, NY), anti-bFGF neutralizing antibody (bFM-1) (Upstate Biotechnology), the PAR1-activating peptides (PAR-APs) AparafluoroFRCyclohexylACitY-NH2 (Cit-NH2) and TFLLR-NH2 synthesized by Dr. Dennis McMaster (University of Calgary, Peptide Synthesis Facility), and the thrombin-derived peptides TP367 (YPPWNKNFTRNDLL) and TP508 (AGYKPDEGKRGDACEGDSGGPFV) synthesized by Biochem Therapeutic (Dorval, QC, Canada). All peptides were added to the cell culture in the presence of amastatin (10 µM) (Sigma) to prevent proteolysis of the peptides. Inhibition of COX-2 was achieved by preincubating cells for 20 min with the selective COX-2 inhibitors celecoxib (1 µM) and NS398 (30 µM) and applying the agonists in the continued presence of these inhibitors.

RT-PCR. CCD-18Co were grown as described above on 75-cm2 Falcon tissue culture flasks (VWR, Edmonton, AB, Canada) and scraped from the base of the flask in the presence of the RNEasy lysis buffer (Qiagen, Mississauga, ON, Canada). Total RNA was extracted from the cells using RNeasy (Qiagen) and measured on a Gene Quant II nucleic acid analyzer (Amersham Pharmacia Biotech, Baie d'Urfé, QC, Canada). RNA (2 µg) was subjected to first-strand cDNA synthesis using Superscript RNase H Reverse Transcriptase (300 U) (GIBCO, Toronto, ON, Canada) in the presence of 1 mM dNTPs (Amersham Pharmacia Biotech), 35.4 µg/ml random hexamer nucleotide primers (Amersham Pharmacia Biotech), 17 U RNAGuard (Amersham Pharmacia Biotech), and buffer giving 50 mM KCl, 1.5 mM MgCl2, and 10 mM Tris · HCl in a final volume of 20 µl. In a DNA Engine PTC-200 Peltier thermal cycler (MJ Research, Waltham, MA), the RT reaction was performed for 10 min at 25°C, and then at 42°C for 50 min and at 95°C for 10 min.

PCR for PAR1 and PAR4 (also activated by thrombin) was performed on cDNA from the RT reaction using the forward (5' CAC GGA TCC TAT TTT TCC GGC AGT GAT TGG 3') and reverse (5' CAC GAA TTC TCA AAT GAT AGA CAC ATA ACA GAC CCT 3') primers for human PAR1 (2) and forward (5' AAT GGG CTG GCG CTG TGG GTG 3') and reverse (5' CGG AAG GTC TGC CG (C/A) TGCA 3') primers for human PAR4. The PCR reaction contained 20 U/ml Taq polymerase (GIBCO), 0.2 mM dNTPs, 20 pmol each of forward and reverse primers, 2 µl cDNA, and PCR buffer giving 50 mM KCl, 1.5 mM MgCl2, and 10 mM Tris · HCl in a final volume of 50 µl and was cycled for 35 cycles (denaturation at 94°C for 1 min, annealing at 55°C for 30 s, and elongation at 72°C for 1 min). PCR products were separated on a 1.3% agarose gel and visualized using a GelDoc 2000 ultraviolet (UV) imager (BioRad, Mississauga, ON, Canada). Products were sequenced (DNA Sequencing Lab, University of Calgary) to confirm identity.

Calcium imaging. CCD-18Co were grown on 12-mm diameter glass coverslips (Warner Instrument, Hamden, CT). Cells were washed twice with MEM containing 25 mM HEPES (Sigma) and 10 mM tetramethyl ammonium hydroxide (TMA) (Sigma) and incubated with 5 µM fura 2-AM and 0.01% pluronic F-127 (Molecular Probes, Eugene, OR) in MEM/HEPES/TMA for 45 min at room temperature in the dark. Coverslips were mounted in a perfusion chamber (Warner Instruments) on a Zeiss Axiovert 135 microscope. Cells were stimulated with PAR agonists by replacement of the 500 µl of perfusion buffer (145 mM NaCl, 0.01 M HEPES, 0.4 mM tetramethyl ammonium hydroxide, 0.01 M D-glucose, and 0.1 mM CaCl2) with agonists prediluted in perfusion buffer. Specificity of the responses to PAR-1 activation was shown using receptor desensitization, as we have previously demonstrated for PAR-1-induced epithelial chloride secretion (16). Fura 2 fluorescence was measured through a Fluor ×20 objective and a Chroma filter set using ImageMaster System software and DeltaRAM rapid wavelength-switching illuminator (Photon Technology International, London, ON, Canada). Fura 2 fluorescence was expressed as the ratio of emission at 510 nm with excitation at 340 and 380 nm.

Immunocytochemistry. CCD-18Co fibroblast cells were grown on LabtekII chamber slides (VWR, Mississauga, ON, Canada) and incubated with the above agents at 37°C for 18 h before staining. Cells were washed with Tris-buffered saline (TBS) (0.15 M NaCl and 50 µM Tris, pH 7.6) and then fixed on ice with methanol for 20 min. To inhibit endogenous peroxidases, cells were incubated in 0.1% sodium azide (Sigma) containing 3% hydrogen peroxide (Sigma) for 30 min. Slides were then washed three times with TBS with 5-min intervals separating each wash. Nonspecific binding was blocked with Dulbecco's MEM (DMEM) (Sigma) containing 10% fetal bovine serum and 3 mg/ml of bovine serum albumin (Sigma) for 30 min. The slide was then drained but not washed. Cells were incubated with the COX-2 mouse monoclonal primary antibody (Cayman Chemical, Ann Arbor, MI) at a concentration of 5 µg/ml, diluted in TBS at room temperature for 1 h, and washed 3 times. A biotin-labeled rabbit anti-mouse secondary antibody (DAKO, Mississauga, ON, Canada) was applied to the cells at a dilution of 1:1,000 at room temperature for 1 h and washed 3 times. To amplify the staining signal, a streptavidin-biotin-horseradish peroxidase complex (SAB-HRP) was applied to the cells. SAB-HRP was used at a 1:200 dilution in Tris · HCl (pH 7.6), incubated at room temperature for 1 h, and washed 3 times. Staining was visualized with aminoethylcarbazole (AEC), which produces a red reaction product using an AEC kit (DAKO), prepared following the manufacturer's instructions. Cells were then washed once with TBS and once with distilled water. The cells were counterstained with Mayer's hematoxylin for 2 min and washed in water for 5 min. The slide was mounted with AquaPerm Mounting Medium (Shandon, PA) and dried overnight. Coverslips were placed over the slides using DPX Mountant (Fluka, Oakville, ON, Canada). The slides were examined under light microscopy at ×400 magnification. A positive cell was distinguished as having red, cell-associated staining. Positive cells were counted from a minimum of 200 cells and expressed as percent positive. Cell counts were verified by an observer unaware of the treatment group.

PGE2 enzyme immunoassay. CCD-18Co were incubated for 18 h at 37°C in the presence or absence of the agonists listed above. Cells were washed in Dulbecco's phosphate-buffered saline (PBS) (Sigma) and incubated for 15 min in fresh, prewarmed PBS at 37°C. The resulting supernatant was collected and immediately frozen at -80°C. The cells were dissociated with 1.5× trypsin-ethylenediaminetetraacetic acid (EDTA) (Sigma), and the number of cells in each well was counted on an Improved Neubauer Hemacytometer. PGE2 levels were determined in the appropriately diluted supernatants using a PGE2 enzyme immunoassay kit (Cayman Chemical) according to the manufacturer's instructions. The results were expressed as nanograms per million cells to reduce any error produced by agents possessing mitogenic activity and, where necessary, normalized to the vehicle control. According to the manufacturer, the antibody used in the PGE2 immunoassay crossreacts 100% with PGE2 ethanolamide, 43% with PGE3, 18.7% with PGE1, and 1% with 6-keto PGF1alpha , but at negligible levels with other eicosanoid pathway products.

Thrombin activity assay. Thrombin activity was measured using the thrombin substrate S-2238 (Diapharm, West Chester, OH) in a volume of 125 µl. Thrombin (100 U/ml-final concentration) was incubated with or without D-Phe-Pro-Arg-chloromethylketone (PPACK; Calbiochem, La Jolla, CA; 100 µM final concentration) on ice for 30 min before assay. One hundred microliters of the chromogenic substrate S-2238 (1 mM) were added, and the change in absorbance at 405 nm was measured over 5 min using a UVmax kinetic microplate reader (Molecular Devices, Sunnyvale, CA) and expressed as milli optical density units per minute.

Statistics. Unless otherwise stated, PGE2 data are calculated as nanograms per million cells and expressed as a percentage of control (100%). Control values are designated such that the addition of the agonist is the only variable, thus eliminating effects of pretreatment of cells with amastatin, PPACK, anti-bFGF, or combinations thereof on baseline PGE2 values. Paired sample comparisons were performed using the paired Student's t-test. When the Student's t-test was used on normalized data, a Kolmogorov-Smirnov test for normality was performed. In each case, the data passed this test. A paired test is justified because each experiment was done on cells from the same passage number and grown in the same large flask before splitting into smaller flasks for experimental manipulation; thus they are considered to be from the same population. In concentration dependency experiments, the ANOVA multiple comparisons test was performed with a Bonferroni posttest. Statistical tests were performed using INSTAT statistical software (Graphpad Software, San Diego, CA) . In all cases, P <=  0.05 was considered significant.


    RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

PAR1 expression. CCD-18Co expression of PAR1 was investigated by RT-PCR and desensitizing calcium responses to PAR1 agonists. PAR1 mRNA expression was observed by RT-PCR and produced a defined band at 401 bases (Fig. 1A). Its identity as human PAR1 was confirmed by sequencing of the PCR product and comparison to the GenBank database (accession no. NM-001992.2) Calcium mobilization assays were performed on monolayers of CCD-18Co using thrombin, TFLLR-NH2, and Cit-NH2. Thrombin at 100 U/ml produced a robust fluorescence from fura 2-AM loaded cells, which desensitized the response to both TFLLR-NH2 (100 µM) and Cit-NH2 (50 µM), although these responses were not fully abrogated (Fig. 1B, a-c). TFLLR-NH2 (100 µM) and Cit-NH2 (50 µM) fully desensitized the response to the other PAR1-AP (Fig. 1B, c and d) and the response to thrombin (100 U/ml) (Fig. 1B, e and f).


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Fig. 1.   Expression of proteinase-activating receptor 1 (PAR1) in CCD-18Co. A: agarose gel of RT-PCR products from CCD-18Co. Lane 1: PAR1 expression in CCD-18Co. Lane 2: absence of PAR4 mRNA in CCD-18Co compared with mRNA from a PAR4-transfected cell line (lane 3). Lanes 4 and 5: PCR for PAR1 and PAR4 in the absence of RT product. B: representative traces from calcium mobilization experiments for CCD-18Co using fura 2-AM fluoresence ratio (340/380 nm). a: addition of thrombin (100 U/ml) reduced the subsequent response to the specific PAR1AP Cit-NH2 (50 µM) when compared with the response from naïve cells (shown in c). b: addition of thrombin (100 U/ml) reduced the subsequent response to the specific PAR1AP TFLLR-NH2 (100 µM) when compared with the response from naïve cells (shown in d). c: addition of the PAR1-activating peptide Cit-NH2 (50 µM) abrogated the subsequent response to TFLLR-NH2 (100 µM) when compared with the response from naïve cells (shown in d). d: addition of TFLLR-NH2 (100 µM) reduced the subsequent response to Cit-NH2 (50 µM) when compared with the response from naïve cells (shown in c). e: addition of Cit-NH2 (50 µM) fully abrogated the subsequent response to thrombin (100 U/ml) when compared with the response from naïve cells (shown in a). f: addition of TFLLR-NH2 (100 µM) reduced the subsequent response to thrombin (100 U/ml) when compared with the response from naïve cells (shown in a).

In the calcium desensitization studies, there was no residual thrombin response after the addition of either Cit-NH2 or TFLLR-NH2 (Fig. 1B), suggesting that there is no PAR4 expressed on CCD-18Co and that the effects of thrombin are not, therefore, PAR4 mediated. This result was substantiated by RT-PCR; CCD-18Co cDNA showed no signal for PAR4 when compared with cDNA from a PAR4-transfected cell line (Fig. 1A).

Effect of thrombin on PGE2 release. In experiments measuring basal PGE2 release after 18 h incubation with thrombin (25-100 U/ml), PGE2 released in a 15-min period was elevated in a concentration-dependent manner by thrombin (P < 0.05, n = 5), and a >3-fold increase was observed at 100 U/ml (P < 0.05, n = 5) (Fig. 2A). Because of the substantial baseline level of PGE2 synthesis (Fig. 2A), experiments were repeated in which CCD-18Co cells were grown for 24 h in media containing no serum to eliminate the possible effects of serum on COX-2 expression. Serum-starved cells did not show altered basal COX-2 expression as measured by immunocytochemistry (data not shown). PGE2 synthesis was lower in cells grown in media containing serum than in serum-starved cells (serum-starved, 1,786 ± 224 pg/106 cells vs. serum-fed, 250 ± 69 pg/106 cells; P > 0.05). Using low-dose indomethacin (100 nM) to block COX-1 did not affect baseline PGE2 synthesis in serum-starved cells (1,395 ± 363 pg/106 cells; P > 0.05 vs. serum-starved cells not receiving indomethacin). Increased PGE2 synthesis in serum-starved cells may represent a stress response in this cell line.


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Fig. 2.   A: effect of thrombin 0-100 U/ml on prostaglandin E2 (PGE2) synthesis. Thrombin induced a concentration-dependent increase in PGE2 synthesis (n = 5). * P < 0.05 vs. 0 U/ml thrombin. B: effect of Cit-NH2 (50 µM), TP367 (100 µM), and TP508 (100 µM) in the presence of amastatin (10 µM) on PGE2 synthesis. Data are calculated as ng/million cells and expressed as % of control (amastatin 10 µM). * P < 0.05 vs. control.

Effect of PAR1 activation on PGE2 release. In PAR1 desensitization experiments (calcium measurements), Cit-NH2 (50 µM) produced a calcium response equivalent to that evoked by thrombin (100 U/ml) (Fig. 1) and, therefore, was added to the cultures at this concentration in the presence of the amino peptidase inhibitor amastatin (10 µM) for 18 h. Cit-NH2 caused a 2.5-fold increase in PGE2 release compared with control (P < 0.05, n = 10) (Fig. 2B), suggesting that thrombin activation of PAR1 is involved in the upregulation of COX-2-derived PGE2 synthesis. The addition of amastatin in these experiments did not significantly alter PGE2 release compared with untreated cells (data not shown).

The effect of thrombin-derived peptides TP367 and TP508 on PGE2 release. The addition of peptides corresponding to noncatalytically active regions of thrombin, TP367 (100 µM) and TP508 (100 µM), to the CCD-18Co cultures in the presence of amastatin (10 µM) for 18 h caused significant increases in PGE2 synthesis to 176 ± 29% of control samples (treated with amastatin alone) (P < 0.05, n = 8) and 141 ± 12% of control samples (amastatin alone) (P < 0.01, n = 8), respectively (Fig. 2B).

COX-2 immunocytochemistry. Expression of COX-2 by CCD-18Co was measured by immunocytochemistry and expressed as the percentage of cells staining positively for COX-2. In untreated cells, 29 ± 8% of cells had constitutive expression of COX-2. After treatment with thrombin (100 U/ml) for 19 h, an additional 18 ± 8% of cells expressed COX-2 such that 47 ± 10% of cells were COX-2 positive (P < 0.05, n = 11) (Fig. 3A). The addition of Cit-NH2 with amastatin increased COX-2 expression by 11 ± 6% (rising from 11 ± 3% to 23 ± 8%; P < 0.05; n = 6) (Fig. 3A). The thrombin-derived peptides TP367 and TP508 with amastatin also increased COX-2 expression by 18 ± 7% (rising from 16 ± 7% to 34 ± 10%; P < 0.05; n = 12) and 16 ± 8% (rising from 9 ± 2% to 25 ± 9%; P < 0.05; n = 10), respectively (Fig. 3A).


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Fig. 3.   A: increase in cyclooxygenase (COX)-2 expression in CCD-18Co after treatment for 18 h with thrombin (100 U/ml), Cit-NH2 (50 µM), TP367 (100 µM), and TP508 (100 µM) after subtraction of controls. Thrombin was compared with untreated cells, and peptides were added in the presence of amastatin (10 µM) and compared with amastatin controls. * P < 0.05 vs. control. B: effect of celecoxib (1 µM) (cells are pretreated for 20 min before collection of cell supernatants) on thrombin (100 U/ml)-, Cit-NH2 (50 µM)-, TP367 (100 µM)-, and TP508 (100 µM)-induced PGE2 synthesis. Data are calculated as PGE2 ng/million cells and expressed as % of control (controls are untreated cells for thrombin and amastatin 10 µM for peptides). * P < 0.05 vs. control. # P < 0.05 vs. agonist in the absence of celecoxib.

Selective COX-2 inhibition. PGE2 production by myofibroblasts has previously been shown to be predominantly COX-2 derived (25). We have found that in thrombin-treated CCD-18Co, subsequent production of PGE2 is inhibited by 80% (P < 0.05, n = 3) by the selective COX-2 inhibitor celecoxib (1 µM), confirming that the PGE2 is COX-2 derived (Fig. 3B). Similar results were obtained with NS398 (30 µM) (data not shown). Selective inhibition of COX-2 by celecoxib also significantly inhibited PGE2 synthesis in Cit-NH2 treated cells by 65% (P < 0.01, n = 7) (Fig. 3B). Celecoxib inhibited the PGE2 synthesis in cells treated with TP367 and TP508 by 47% (P < 0.01, n = 9) and 74% (P < 0.05, n = 8), respectively (Fig. 3B).

Noncatalytic activity of thrombin. The simultaneous addition of Cit-NH2 and TP508 in PGE2 release experiments further increased the PGE2 released from the cells above that achieved with Cit-NH2 (P < 0.05) (Fig. 4A) or TP508 alone (P < 0.05, n = 7) (Fig. 4A). The combination of Cit-NH2 and TP367 was unable to elevate PGE2 release above that of Cit-NH2 (n = 5) (Fig. 4A). The effect of the combinations of Cit-NH2 and TP508 may indicate an additive relationship between PAR1 activation and the TP508 "receptor."


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Fig. 4.   A: effect of adding Cit-NH2 (50 µM) in combination with either TP367 (100 µM) or TP508 (100 µM) for 18 h on PGE2 release. * P < 0.05 vs. Cit-NH2, # P < 0.05 vs. TP508. B: effect of D-Phe-Pro-Arg-chloromethylketone (PPACK; 100 µM) (preincubated with agonist >30 min before addition to cells) on PGE2 synthesis induced by thrombin (100 U/ml). * P < 0.05 vs. control (100%) (controls are untreated cells ± PPACK 100 µM for thrombin). NS, not significantly different from each other.

The inhibition of the catalytic activity of thrombin by preincubation for 30 min with PPACK (100 µM) for 30 min before the addition to cultures for 18 h did not significantly reduce PGE2 release compared with catalytically active thrombin (P > 0.1, n = 9) (Fig. 4). There was a residual increase in PGE2 release after incubation with PPACK-thrombin that remained significantly greater than control cultures (P < 0.05, n = 9) (Fig. 4B). PPACK alone did not have any effect on PGE2 synthesis (P > 0.05). As such, data were expressed as a percentage of control in the presence of PPACK where appropriate. PPACK-thrombin was confirmed to be completely catalytically inactive (retaining 0% activity) compared with thrombin by enzyme activity assay using the chromogenic thrombin substrate S-2238.

The role of bFGF in the thrombin/peptide-induced increase in COX-2 expression and PGE2 release. Some effects of thrombin have been attributed to the secondary release of bFGF (39, 42), and bFGF has been implicated in the upregulation of COX-2 in several cell types (27, 28, 33, 41). Thus we sought to evaluate a potential role for bFGF in the induction of PGE2 synthesis by thrombin. The addition of bFGF (2.5-25 ng/ml) to CCD-18Co cultures for 18 h caused a concentration-dependent increase in PGE2 release (P < 0.05, n = 5) with a maximal twofold increase at 10 ng/ml (P < 0.05) (Fig. 5A). Celecoxib significantly inhibited the PGE2 synthesis in bFGF-treated cells by 88% (P < 0.05, n = 3) (from 46.4 ± 12.6 to 5.9 ± 0.6 ng/million cells), confirming that the PGE2 is primarily COX-2 derived.


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Fig. 5.   A: effect of basic fibroblast growth factor (bFGF) 0-100 ng/ml on PGE2 synthesis by CCD-18Co. An 18-h incubation of cells with bFGF increased PGE2 synthesis in a concentration-dependent manner (n = 5). * P < 0.05 vs. control (untreated cells). B: effect of the anti-bFGF neutralizing antibody (20 µg/ml) on PGE2 synthesis induced by thrombin (100 U/ml), Cit-NH2 (50 µM), TP367 (100 µM), and TP508 (100 µM). Data are calculated as % of control (controls are untreated cells ± anti-bFGF or amastatin ± anti-bFGF as appropriate) and expressed as % inhibition). * P < 0.05 vs. agonists in the absence of anti-bFGF.

The PGE2 release after incubation with TP367 and TP508 was significantly reduced by the addition of an anti-bFGF neutralizing antibody (20 µg/ml) by 45% (P < 0.05, n = 4) and 40% (P < 0.05, n = 4, respectively) (Fig. 5B). In contrast, the PGE2 release stimulated by the PAR1 agonist Cit-NH2 was not reduced by anti-bFGF (P > 0.4, n = 6) (Fig. 5B). The response to thrombin was significantly inhibited (18%) by the addition of anti-bFGF (P = 0.05, n = 8) (Fig. 5B).


    DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The role of fibroblast-derived PGE2 in regulating the secretory responses of the epithelium in the intestinal mucosa is well established (11). In this paper, we have found a dual mechanism by which thrombin could induce epithelial chloride secretion via both catalytic and noncatalytic mechanisms in addition to, but independently from, its direct action on epithelial PAR1 (16).

Our results are in keeping with previous reports that thrombin causes an increase in COX-2 expression and subsequent PGE2 synthesis in hamster fibroblasts (20). In the cited study, the PAR1-activating peptide SFLLRN increased both COX-2 expression and PGE2 synthesis, suggesting that thrombin acts through the proteolytic cleavage of PAR1 (20). In the current study, using human colonic myofibroblasts, we observed significant increases in PGE2 synthesis not only via activation of PAR1 by the specific PAR1AP Cit-NH2 but also by the actions of peptides corresponding to regions of the thrombin sequence that exert their effects independently of its catalytic activity. Catalytically inactive PPACK-thrombin was also able to increase PGE2 synthesis by about 50%, suggesting that the increase in PGE2 synthesis induced by thrombin has both a PAR1-mediated component and one that is not reliant on the catalytic activity of thrombin. This conclusion is consistent with the ability of either Cit-NH2 or the thrombin-derived peptides TP367 and TP508 to mimic the response to thrombin and that TP508 and Cit-NH2, in combination, augment the increase in PGE2 synthesis toward the levels caused by thrombin alone. Because PPACK is a short peptide inhibitor that reacts covalently with the catalytic site on thrombin, it is unlikely to interact with or inhibit the noncatalytic regions of thrombin. In contrast, the specific thrombin inhibitors antithrombin (AT) III and hirudin (7) could be expected to mask thrombin's noncatalytic domains. This issue is of particular consequence for the region corresponding to TP367, which is located in close proximity to (or containing) residues involved in both the catalytic activity of thrombin and the hirudin and AT III binding sites (7, 8). This proximity to the catalytic site is apparently not the case for TP508 (9).

This dual effect (catalytic and noncatalytic) of thrombin has been shown previously in the proliferative responses of hamster fibroblasts (17, 26) and endothelial cells (24), where the action of thrombin can only be fully mimicked by activation of PAR1 (by PAR1-AP or gamma -thrombin) in the presence of the TP367 peptide (17) or catalytically inactive alpha -thrombin (17, 24).

The presence of bFGF has been shown to be a requisite factor in thrombin-induced mitogenesis in both hamster lung fibroblasts (39) and human vascular smooth muscle cells (42). We therefore examined the role of bFGF in mediating our thrombin-induced increases in COX-2 expression and PGE2 release. Our data show that, indeed, the addition of bFGF to our cultures concentration dependently increased PGE2 release and that the addition of anti-bFGF-neutralizing antibody concurrently with our stimuli significantly inhibited the responses to both TP367 and TP508 by ~40%. Interestingly, the response to thrombin was inhibited by about 15-20%, while there was no change in the response to Cit-NH2. These results suggest that a component (~20%) of the thrombin response is both PAR1 independent and bFGF dependent. This concept is supported by the work of Vouret-Craviari and colleagues (39), who observed that the PAR1-activating peptide was unable to mimic the thrombin-induced mitogenesis in the absence of bFGF (39). Because we have determined a dependence of the actions of TP367 and TP508 on bFGF, this result might indicate that the bFGF-dependent component of the response to thrombin is linked to its noncatalytic domains. The dissociation of the bFGF effect from a PAR-1 or catalytically mediated effect likely explains the incomplete inhibition of PGE2 synthesis obtained by pretreatment of the cells with anti-bFGF antibody.

The receptor(s) for the noncatalytic regions of thrombin or TP367 or TP508 have not been characterized or identified, and their signaling pathways are largely unknown. However, reports indicate that the "high affinity" receptor for TP508 might be a member of the beta 3-integrin family based on the presence in TP508 of the "RGD" sequence (4, 9). It has been suggested that TP508 signals via a transient increase in cAMP (22). The "loop B"-containing noncatalytic region of thrombin or TP367 peptide has some structural homology to colony-stimulating factors and may act through receptors for these polypeptides (8). In a study by Sower et al. (34), TP508, when applied to human fibroblasts, initiates signals distinct from those induced by PAR1 activation and results in differential gene expression.

Three-dimensional models of thrombin suggest that the loop B-containing region of thrombin (as mimicked by TP367) is surface exposed and might have the opportunity to interact with receptors on the cell membrane (13). However, the RGD-containing sequence (as mimicked by TP508) is thought not to be surface exposed in native thrombin (14) and, as a result, might require autoproteolysis for surface exposure (9, 12) The presence of heparan sulphate proteoglycan and plasmin has been implicated in the exposure of this sequence of thrombin (3, 4). Thus, in our cultures, the noncatalytic activity of thrombin may be a result of the loop B-containing region only, or, indeed, both regions may be active after autoproteolysis of thrombin in the culture system.

Previous studies have shown that in IL-1-treated fibroblasts, the COX-2 pathway accounts for 95% of total PGE2 synthesis (25). This result is confirmed for thrombin in our studies by the use of the selective COX-2 inhibitor celecoxib at a concentration (1 µM) specific for the COX-2 isoform. At this concentration of celecoxib, we observed a 45-80% reduction in PGE2 synthesis in thrombin-, Cit-NH2-, TP367-, and TP508-treated cells but no significant reduction in nontreated cells. This COX-2 inhibition would support the reports that in treated myofibroblasts, PGE2 is predominantly COX-2 derived but also suggests that there is a COX-1-mediated component, particularly in untreated cells. The prostaglandin E synthases (PGES) downstream of COX (37) may also be important in regulating PGE2 synthesis.

In our studies, the percent increase in PGE2 synthesis appears larger than the relatively modest increase in COX-2 expression. Our measurement of COX-2 expression, however, only takes into account the percentage of cells expressing COX-2 and not the amount of active COX-2 expressed per cell. Because the latter is also likely to increase after exposure to thrombin, TP peptides, or Cit-NH2, it is not surprising that the increase in PGE2 is not proportionate to the increased number of COX-2-positive cells. In addition, we observed that basal PGE2 synthesis was relatively high and was not blocked by either low dose indomethacin (to block COX-1) or celecoxib (to block COX-2). This observation remains to be investigated but does not detract from our main observation of thrombin-induced COX-2 expression and activity.

The induction of epithelial chloride secretion by elevated PGE2 synthesis is a host defense mechanism (11) induced by the presence of bacterial products and allergens, thus preventing tissue invasion and injury (10, 31). In these circumstances, the luminal bacteria and their products (as mimicked by LPS) induce an inflammatory response that includes increased vascular permeability (15, 30). Because the myofibroblast is closely associated with the capillary network of intestinal villi (29), elevations in mucosal levels of thrombin resulting from vascular leakage will, according to our data, increase COX-2-derived PGE2 release from the myofibroblast and thereby induce epithelial chloride secretion. This action of thrombin, therefore, represents an important host defense mechanism. As such, the use of selective COX-2 inhibitors may have a deleterious effect on the ability of the intestine to protect itself against injurious agents.

Our study, therefore, documents a novel mechanism whereby epithelial chloride secretion and host defense may be regulated by extravasated thrombin in two ways. First, thrombin acts catalytically through activation of PAR1. Second, thrombin acts via its noncatalytic domains to stimulate bFGF. Both actions of thrombin induce COX-2 expression. Because COX-2 produces the myofibroblast-derived PGE2, which regulates the host defense mechanism of epithelial chloride secretion, this mechanism may have therapeutic implications for the use of both selective COX-2 inhibitors and classic NSAIDs.


    ACKNOWLEDGEMENTS

We thank Dr. Jonathon Lytton and Stephen Leech for use of, and assistance with, the calcium imaging equipment, Patricia Andrade-Gordon for the use of the PAR4-transfected cell line, and Kelly Cushing for technical assistance.


    FOOTNOTES

M. L. Seymour is the recipient of a fellowship from the Canadian Association of Gastroenterology in association with Canadian Institutes of Health Research, and Solvay Pharma. N. F. Zaidi is the recipient of a Canadian Association of Gastroenterology summer studentship. W. K. MacNaughton is an Alberta Heritage Foundation for Medical Research Scholar.

This work was funded by the Canadian Institutes of Health Research (to W. K. MacNaughton and M. D. Hollenberg).

Address for reprint requests and other correspondence: W. K. MacNaughton, Mucosal Inflammation Research Group, Univ. of Calgary, 3330 Hospital Drive NW, Calgary, Alberta, Canada T2N 4N1 (E-mail: wmacnaug{at}ucalgary.ca).

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.

First published December 27, 2002;10.1152/ajpcell.00126.2002

Received 19 March 2002; accepted in final form 17 December 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
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