A role for proteinase-activated receptor 2 and PKC-{epsilon} in thrombin-mediated induction of decay-accelerating factor on human endothelial cells

Elaine A. Lidington,1 Rivka Steinberg,1 Anne R. Kinderlerer,1 R. Clive Landis,1 Motoi Ohba,2 Allen Samarel,3 Dorian O. Haskard,1 and Justin C. Mason1

1British Heart Foundation Cardiovascular Medicine Unit, Eric Bywaters Centre, Imperial College London, Hammersmith Hospital, London, United Kingdom; 2Institute of Molecular Oncology, Showa University, Tokyo, Japan; and 3Cardiovascular Institute, Stritch School of Medicine, Loyola University Chicago, Maywood, Illinois

Submitted 13 October 2004 ; accepted in final form 2 August 2005


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Thrombin, an important mediator of thrombosis and inflammation, may also enhance vascular cytoprotection. Thus thrombin induces expression of the complement-inhibitory protein decay-accelerating factor (DAF) in human umbilical vein endothelial cells (HUVECs), thus increasing protection against complement-mediated injury. Using PKC isozyme-specific peptide antagonists and adenoviral constructs, we have shown in the present study that PKC-{epsilon} is the primary isozyme involved in DAF induction by thrombin. Experiments with proteinase-activated receptor-1 (PAR1) and PAR2 activating peptides (APs) showed that DAF expression induced by PAR1-AP was PKC-{alpha}-dependent; in contrast, PAR2-AP induction of DAF required activation of PKC-{epsilon}. PAR1-AP and PAR2-AP in combination exerted an additive effect on DAF protein expression, which was equivalent to that observed with thrombin alone. These data implied a specific role for PAR2 in DAF induction, which was supported by the observation that upregulation of endothelial cell (EC) PAR2-enhanced DAF induction by thrombin. ERK1/2, p38, and JNK MAPK were also involved in thrombin-induced DAF upregulation, with evidence of interdependence between ERK1/2 and JNK. A role for transactivation of PAR2 by PAR1 was suggested by partial inhibition of thrombin-induced DAF expression by the PAR1 signaling antagonists BMS-200261 and SCH79797, whereas inhibition of thrombin-induced cleavage of PAR1 by specific MAbs or hirudin completely abrogated the response. Together, these data imply that the predominant pathway for thrombin-induced DAF expression involves transactivation of PAR2 by PAR1 and signaling via PKC-{epsilon}/MAPK. This may represent an important, novel pathway for endothelial cytoprotection during inflammation and angiogenesis and suggests that PAR2 may play a central role in some thrombin-induced responses.

cytoprotection; proteinase-activated receptor 1


THROMBIN, A MULTIFUNCTIONAL serine proteinase best known for its ability to cleave fibrinogen to fibrin, also exerts important effects on endothelial cells (ECs) (9). Thrombin-mediated signaling results in a coordinated combination of responses, including cytoskeletal reorganization, increased permeability, hemostasis, angiogenesis, and leukocyte adhesion and transmigration. Many of these actions, including the secretion of chemokines (8) and the synthesis of matrix metalloproteinases (MMPs) (11), growth factors (57), or membrane-bound molecules such as E-selectin and vascular cell adhesion molecule-1 (VCAM-1) (24, 25, 63), are protein synthesis dependent. Although the proinflammatory actions of thrombin are well recognized, thrombin's potential as a cytoprotective factor is a relatively novel concept (30, 41). Thrombin has been reported to be antiapoptotic (6) and proangiogenic (63), and we have shown that thrombin upregulates DAF expression on the EC surface to levels that significantly enhance resistance to complement-mediated injury (30).

Cell surface receptors for thrombin are members of the proteinase-activated receptor (PAR) family of G protein-coupled, seven-transmembrane receptors activated after proteolytic cleavage and exposure of a tethered ligand (9). Of the four members of the PAR family, only PAR1, PAR3, and PAR4 are cleaved by thrombin. PAR1 is the primary thrombin receptor, although it is also cleaved by factor Xa and activated protein C (54). PAR2, while not cleaved directly by thrombin, can be activated by proteinases, including trypsin and mast cell tryptase. In addition, it has been reported that PAR2 can be transactivated by the thrombin-cleaved, PAR1-tethered ligand, at least in the presence of PAR1 antagonists (47). Expression of PAR2, but not PAR1, is upregulated in ECs by TNF-{alpha}, IL-1{beta}, and LPS (46). PAR3 is poorly expressed on ECs and PAR4, is undetectable on human umbilical vein ECs (HUVECs), and is expressed by human arterial ECs (15), where expression is regulated by proinflammatory cytokines (17). Although no function has been established in human ECs, PAR4 plays a critical role in human platelet responses to a high concentration of thrombin (23) and in thrombin-induced responses in rodent ECs (26).

PAR-mediated signaling via G proteins activates multiple downstream pathways, including those regulated by MAPKs, PKC, and phosphatidylinositol 3-kinase (PI3-kinase)/Akt, which may act sequentially or in parallel (30, 41). Thrombin induces rapid phosphorylation of ERK1/2, p38 MAPK, and JNK, all of which have been implicated in thrombin-mediated responses (34, 49). Thrombin has been shown to use distinct PKC isozymes, including PKC-{alpha}, -{delta}, -{epsilon}, and -{zeta}. Thus thrombin-induced intercellular adhesion molecule-1 (ICAM-1) is PKC-{delta} dependent (52), thrombin-induced VCAM-1 requires activation of both PKC-{delta} and -{zeta}, and thrombin-mediated changes in vascular permeability follow activation of PKC-{alpha} (39). Using specific agonist peptides, EC PAR1 and PAR2 have been reported to use the same signaling pathways. However, functional differences have been identified downstream; for example, only ligation of PAR1 induces monocyte chemoattractant protein-1 (MCP-1) (54) and alters vascular permeability (28), reflecting prolonged activation of RhoA downstream of PAR1 but not PAR2 (66).

Located at the blood-tissue interface, vascular endothelium is continually exposed to autologous bystander injury by complement components, a risk that is enhanced during inflammation. Furthermore, complement activation has an established role in acute coronary syndromes (20) and may also be involved in the pathogenesis of atherosclerosis and myocardial infarction (45, 62). To combat this pathology, both constitutive and inducible cytoprotective mechanisms have evolved to limit complement-mediated injury, including both cell surface and soluble regulatory proteins (32). The membrane-bound complement-inhibitory proteins DAF (CD55), membrane cofactor protein (MCP) (CD46), and cluster of differentiation (CD59) use distinct mechanisms for the regulation of complement and may act cooperatively (32). DAF is a multifunctional, glycosyl phosphatidylinositol-anchored cell surface glycoprotein, which, by acting at the level of the C3 convertase, interferes with the pivotal step of the complement cascade. The cytoprotective importance of DAF is revealed by the increased susceptibility of DAF-deficient mice to glomerulonephritis and ischemia-reperfusion injury (31, 60, 67). In recent EC studies, we have demonstrated the induction of DAF by TNF-{alpha} and IFN-{gamma} (38), thrombin (30), VEGF, and basic FGF (36, 37) and by 3-hydroxy-3-methylglutaryl-CoA reductase inhibitors (35). The upregulation of DAF protected ECs against C3 deposition and complement-mediated cell lysis.

Although evidence for the induction of EC cytoprotective genes by thrombin is emerging, the identity of the specific PARs and downstream signaling pathways facilitating these responses remain to be defined completely. However, PAR1 and PAR2 have been linked to protective responses in murine models of colitis, bronchoconstriction, and distal ischemia (7, 13, 16, 40). In this study, we have used thrombin-induced DAF expression in ECs as a means by which to explore the role of the PARs and PKC in thrombin-mediated cytoprotection against complement activation. We provide further evidence in support of the hypothesis that cleaved PAR1 may transactivate PAR2, and we describe a novel, thrombin-activated, PAR2/PKC-{epsilon}/MAPK-cytoprotective pathway in human vascular endothelium.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Reagents. Anti-DAF MAb 1H4 was a gift from Dr. D. Lublin (Washington University School of Medicine, St. Louis, MO), and biotinylated anti-DAF (IA10) was purchased from BD Biosciences Pharmingen (San Diego, CA). PAR1-blocking antibodies WEDE15 and ATAP2 were obtained from Beckman Coulter (Luton, UK) and Santa Cruz Biotechnology (Santa Cruz, CA), respectively. MAb 1.2B6 (anti-E- and -P-selectin) was generated in house. Thrombin from human plasma and the aminopeptidase inhibitor amastatin were purchased from Sigma (Poole, UK). APs for PAR1 (SFLLRN-OH and TFLLR-NH2), PAR2 (SLIGKV-OH), PAR4 (AYPGKF-OH), and control peptide (LSIGRL-NH2) were generated by the Imperial College Peptide Synthesis Service. PAR1 signaling peptide inhibitor BMS-200261 was a gift from S. Traynelis (Emory University, Atlanta, GA), and PAR1 antagonist SCH79797 was obtained from Tocris Cookson (Ellisville, MO). GM 6001 and its negative control compound, Gö6976, GF-109203X, U0126, SB-202190, JNK inhibitor II (SP-600125), and JNK inhibitor I peptide (H-Gly-Arg-Lys-Lys-Arg-Agr-Gln-Arg-Arg-Arg-Pro-Pro-Arg-Pro-Lys-Arg-Pro-Thr-Thr-Leu-Asn-Leu-Phe-Pro-Gln-Val-Pro-Arg-Ser-Gln-Asp-Thr-NH2), were purchased from Merck Biosciences (Nottingham, UK). The PKC-{beta} inhibitor LY-379196 was a gift from Dr. K. Ways (Eli Lilly, Indianapolis, IN). Myristoylated (myr)-PKC peptide inhibitors (myr-{Psi}-PKC) (myr-Arg-Phe-Ala-Arg-Lys-Gly-Ala-Leu-Arg-Gln-Lys-Asn-Val) and PKC-{epsilon} v1-2 (myr-Glu-Ala-Val-Ser-Leu-Lys-Pro-Thr) were purchased from Promega (Southampton, UK) and Biomol (Plymouth Meeting, PA), respectively. Myr-PKC-{zeta} (myr-Ser-Ile-Tyr-Arg-Arg-Gly-Ala-Arg-Arg-Trp-Arg-Lys-Leu) was obtained from Merck. Antibodies for immunoblot analysis of phosphorylated and nonphosphorylated ERK1/2, p38, and JNK MAPK were purchased from Cell Signaling (Beverly, MA), and antibodies against PKC isozymes were obtained from Santa Cruz Biotechnology.

Cell isolation and culture. Our human tissue protocols were approved by the hospital Research Ethics Committee. HUVECs were isolated from umbilical cords as described previously (30) and cultured in gelatin-coated tissue culture flasks Costar (Cambridge, MA) in medium-199 (M-199) supplemented with 20% FBS, 100 IU/ml penicillin, 0.1 mg/ml streptomycin, 2 mM L-glutamine (Invitrogen, Paisley, UK), 10 U/ml heparin, and 30 µg/ml EC growth supplement (ECGS; Sigma). For each experiment, ECs were plated in M-199 containing 10% FBS, 5 U/ml heparin, and 15 µg/ml ECGS for 24 h, and the medium was changed to M-199 containing 5% FBS, 2.5 U/ml heparin, and 7.5 µg/ml ECGS for the duration of the experiment.

Flow cytometry. Flow cytometry was performed as previously described (30). Pharmacological antagonists were added 60 min before the addition of agonist, and samples were analyzed using an Epics XL-MCL flow cytometer (Beckman Coulter), which enabled us to count 5,000 cells/sample. Results are expressed as the relative fluorescence intensity (RFI), which represents the mean fluorescence intensity (MFI) with test MAb divided by the MFI using an isotype-matched irrelevant MAb. To allow the results of multiple experiments to be used, the data in some figures are expressed as the percentage of the RFI for thrombin-treated ECs or for ECs treated with inhibitor alone.

Adenovirus infection. The generation of adenovirus (Adv) expression vectors for dominant-negative (DN) and constitutively active (CA) PKC isozymes was described previously (19, 59). DN-PKC-{alpha}, DN-PKC-{epsilon}, CA-PKC-{epsilon}, and {beta}-gal-Adv were amplified in human embryonic kidney HEK-293A cells and purified using the BD Adeno-X purification kit (BD Biosciences, Oxford, UK). Viral titers were estimated using the BD Adeno-X Rapid Titer kit. HUVECs were infected at between 50 and 200 infectious units (IFU)/cell in serum-free M-199 for 2 h before the medium was replaced with M-199–5% FBS, 2.5 U/ml heparin, and 7.5 µg/ml ECGS. Infected ECs were stimulated with thrombin and PAR peptides 24 h postinfection. Infection of HUVECs with a {beta}-gal control Adv demonstrated a transfection efficiency of 95%.

Western blot analysis. HUVECs were cultured for 2 h in 1% BSA to reduce basal phosphorylation and preincubated with MAPK or PKC antagonists for 1 h before stimulation with thrombin for 7 min and lysis in 4 mM EDTA, 50 mM Tris·HCl, pH 7.4, in 150 mM NaCl with 25 mM sodium deoxycholic acid, 200 µM sodium orthovanadate, 10 mM sodium pyrophosphate, 100 mM sodium fluoride, 1% Triton X-100, 1 mM PMSF, and 5% protease inhibitor cocktail (Sigma). After being subjected to SDS-PAGE, separated proteins were transferred to Immobilon P membranes (Millipore, Bedford, MA) and blocked for 1 h at room temperature in BSA-0.1% Tween 20 (vol/vol) with 5% milk powder wt/vol. The membrane was incubated with primary Abs overnight at 4°C, washed in TBS-0.1% Tween 20, and incubated for 1 h at room temperature with appropriate peroxidase-labeled secondary Abs developed with an ECL substrate (Amersham Pharmacia Biotech, Little Chalfont, UK), and exposed to autoradiographic film. When appropriate, blots were stripped and reprobed for detection of the nonphosphorylated form of MAPK. Equal loading was achieved by estimating the lysate protein content using the Bio-Rad Dc protein assay (Bio-Rad, Hercules, CA) and Ponceau staining of membranes before analysis. Integrated density values were measured using the ChemiImager 5500 (Alpha Innotech, San Leandro, CA).

Statistics. Differences between results were evaluated by performing ANOVA with the Bonferroni multiple-comparison test using GraphPad Prism 4.0 software (GraphPad Software, San Diego, CA). Differences of P < 0.05 were considered statistically significant.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Thrombin-induced DAF upregulation is PKC-{epsilon} dependent. In a previous article, we reported a dose-responsive induction of DAF expression in ECs by thrombin, with maximal DAF expression and cytoprotection against complement-mediated injury observed after treatment with 10 U/ml thrombin for 24 h (30). Under these conditions, the upregulation of DAF expression by thrombin was sensitive to the inhibition of PKC (30). However, although thrombin is known to activate PKC-{alpha}, -{delta}, -{epsilon}, and -{zeta} in ECs, the precise PKC isozymes required for DAF induction are unknown. Using optimal conditions for DAF expression by thrombin, we investigated the effect of PKC-specific pharmacological antagonists (Fig. 1A). Initial immunoblotting experiments demonstrated that HUVECs expressed PKC-{alpha}, -{beta}II, -{delta}, -{epsilon}, -{theta}, and -{zeta}, whereas dose-response experiments identified the optimal concentrations of each inhibitor (data not shown). GF-109203X inhibited basal and thrombin-induced DAF expression, suggesting that both constitutive turnover and upregulation of DAF by thrombin require activation of classical (cPKC) and/or novel PKC (nPKC) isozymes. Gö6976, an inhibitor of cPKC isozymes, reduced DAF upregulation by only 20%, whereas LY-397196, a PKC-{beta}-specific antagonist, had no significant effect, suggesting that the action of Gö6976 was mediated by inhibition of PKC-{alpha}. The role of PKC-{alpha} was explored further by using a specific myristoylated cell-permeable peptide PKC-{alpha} and -{beta} antagonist, myr-{Psi}-PKC, and by infection with an adenovirus expressing a PKC-{alpha} dominant-negative construct (DN-PKC-{alpha}). As shown in Fig. 1, B and C, these approaches similarly reduced thrombin-induced DAF expression by only 20%, supporting the involvement of PKC isozymes in addition to PKC-{alpha}.



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Fig. 1. Effect of PKC antagonists on thrombin-induced protein decay-accelerating factor (DAF) expression. A: human umbilical vein endothelial cells (HUVECs) were preincubated with the PKC antagonists GF-109203X (5 µM), Gö6976 (5 µM), or LY-379196 (60 nM) for 1 h before the addition of 10 U/ml thrombin for 20 h (solid bars) or plain HUVEC medium alone (open bars). Endothelial cells ECs were harvested, and DAF expression was analyzed using flow cytometry with MAb 1H4. Data are expressed as percentages of DAF expression in thrombin-treated cells ± SE (n = 3 experiments). B: HUVECs were preincubated with PKC isozyme-specific peptide inhibitors myristoylated (myr)-PKC-{epsilon} v1-2 (myr-{epsilon}), myr-{Psi}-PKC (myr-{alpha}/-{beta}), or myr-PKC-{zeta} (myr-{zeta}) (100 µM) for 1 h before the addition of plain medium alone (Unstim) or 10 U/ml thrombin for 20 h and analysis of DAF expression by flow cytometry (n = 3 experiments). C: HUVECs were infected for 2 h in serum-free medium with adenoviruses (Adv) expressing dominant-negative (DN)-PKC-{epsilon}, DN-PKC-{alpha}, -{epsilon}, or {beta}-gal control Adv [200 infectious units (IFU)/cell] 24 h before the addition of plain HUVEC medium alone (Unstim) or 10 U/ml thrombin in plain medium for 20 h and analysis of DAF expression using flow cytometry (n = 5 experiments). To account for any effect of myristoylated peptides or adenoviruses alone on constitutive DAF expression, the data in B and C are shown as percentages of DAF expression on the untreated, myristoylated peptide- or adenovirus alone-treated ECs, as appropriate, ± SE. *P < 0.05 and **P < 0.005 thrombin + PKC antagonist-treated ECs vs. ECs treated with thrombin alone. D: HUVECs were infected with adenoviruses expressing DN-PKC-{alpha}, DN-PKC-{epsilon}, or {beta}-gal control adenovirus (from 10 to 500 IFU/cell) 24 h before lysis. Lysates were separated using SDS-PAGE, transblotted, and incubated with specific antibodies against PKC-{alpha} or PKC-{epsilon}. Blots were subsequently stripped and reprobed with the opposite PKC antibody to demonstrate the specificity of the DN-PKC adenoviruses.

 
To explore the role of PKC-{epsilon}, ECs were pretreated with the myristoylated PKC-{epsilon} v1-2 peptide inhibitor or by infection with a DN-PKC-{epsilon} adenovirus. Inhibition of PKC-{epsilon} led to complete abrogation of thrombin-induced DAF expression, whereas the {beta}-gal control adenovirus had no effect (Fig. 1, B and C). Moreover, infection of EC with both DN-PKC-{alpha} and DN-PKC-{epsilon} had no greater inhibitory effect than that of DN-PKC-{epsilon} alone. Subsequent experiments investigated the role of PKC-{delta} and PKC-{zeta} in thrombin-induced DAF expression. The PKC-{delta} antagonist rottlerin, used at concentration known to inhibit thrombin-induced ICAM-1 expression in HUVECs (52), did not significantly inhibit thrombin-induced DAF upregulation (data not shown). The inhibition of PKC-{zeta} using a myristoylated peptide reduced thrombin-induced DAF expression by 20%, suggesting that this atypical PKC isozyme may also play a minor role in DAF regulation. The efficacy of myr-{Psi}-PKC and myr-PKC-{zeta} was confirmed by their ability to inhibit VEGF-induced EC proliferation and TNF-{alpha}-induced ICAM-1 upregulation, respectively (data not shown). The myristoylated peptides and adenoviruses had no consistent effect on basal DAF expression. In all of the adenovirus experiments described, the optimal multiplicity of infection for each adenovirus and PKC specificity were predetermined using Western blot analysis with PKC isozyme-specific antibodies (Fig. 1D).

The importance of PKC-{epsilon} in DAF induction was confirmed by infecting HUVECs with an adenovirus expressing a constitutively active form of PKC-{epsilon} (CA-PKC-{epsilon}). CA-PKC-{epsilon} significantly upregulated DAF expression (Fig. 2), and this effect was inhibited by the presence of the MEK-1 antagonist U0126 (data not shown). The specificity of the response was confirmed using Western blot analysis, which demonstrated that overexpression of PKC-{epsilon} increased the activated, phosphorylated form of PKC-{epsilon} in a dose-dependent manner without altering the expression of other PKC isozymes (data not shown). In addition, overexpression of a constitutively active form of PKC-{delta} did not induce DAF expression.



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Fig. 2. Overexpression of PKC-{epsilon} induces DAF expression. A: HUVECs incubated for 2 h in serum-free medium-199 (M-199) alone [unstimulated (US)], in the presence of a constitutively active (CA) PKC-{epsilon} adenovirus (CA-{epsilon}) (50 IFU/cell), or in the presence of a {beta}-gal adenovirus ({beta}-gal) (50 IFU/cell). The medium was then replaced with plain HUVEC medium alone or containing 10 U/ml thrombin (Thr) for 20 h. ECs were harvested, and DAF expression was analyzed using flow cytometry with MAb 1H4. Data are expressed as relative fluorescence intensity (RFI) ± SE (n = 4 experiments). *P < 0.01 and **P < 0.001 vs. US cells.

 
PAR1- and PAR2-specific peptides induce DAF expression. Signaling via PAR1 is thought to regulate the majority of thrombin-induced responses in vascular endothelium. However, it has been reported that PAR2 can be transactivated by thrombin-cleaved PAR1 (47) and that platelet responses to a high concentration of thrombin can be mediated by PAR4. Therefore, the effect of activating specific PARs on HUVECs was initially investigated using the peptide agonists SFLLRN and TFLLR-NH2 (PAR1), SLIGKV (PAR2), and AYPGKF (PAR4), both alone and in combination (Fig. 3). Although SFLLRN and TFLLR-NH2 induced a comparable and significant increase in DAF expression, this increase was significantly less than that observed with the use of thrombin alone (Fig. 3A). The PAR2 agonist peptide SLIGKV induced a modest but significant increase in DAF expression in resting ECs (Fig. 3B). However, in combination with either SFLLRN or TFLLR-NH2, the response was equivalent to that observed with thrombin alone and the level of DAF induction observed with the combination of PAR1 and PAR2 peptides was significantly higher than that demonstrated with either peptide alone (Fig. 3, A and B). In contrast, the PAR4 agonist peptide AYPGKF had no effect on DAF expression and a combination of SLIGKV and AYPGKF revealed no additive effect, whereas AYPGKF alone was able to aggregate platelets (data not shown). It should be noted that SFLLR-derived PAR1 agonist peptides can activate both PAR1 and PAR2 (2, 27), whereas TFLLR-NH2 is more PAR1 specific. However, the upregulation of DAF on ECs induced by SFLLRN and TFLLR-NH2 was directly comparable to and further increased by the presence of the PAR2 peptide SLIGKV (Fig. 3B), suggesting that SFLLRN in this system acts via PAR1.



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Fig. 3. Effect of proteinase-activated receptor (PAR) agonist peptides on DAF expression. A: HUVECs were treated with thrombin (10 U/ml), agonist peptides (400 µM) for PAR1 SFLLRN (SF) and TFLLR-NH2 (TF), and PAR4 AYPGKF (AY), or they were left unstimulated for 20 h before being analyzed for DAF expression using flow cytometry. B: HUVECs were treated with thrombin (10 U/ml) or with PAR2 agonist peptide SLIGKV (SL) alone or in combination with SFLLRN (SF), TFLLR-NH2 (TF), or AYPGKF (AY) (400 µM), or they were left unstimulated for 20 h before being analyzed for DAF expression using flow cytometry. Data are expressed as percentages of DAF expression in untreated cells ± SE (n = 3 experiments with separate ECs). *P < 0.05 for ECs treated with single peptides vs. US cells; **P < 0.05 for ECs treated with PAR1 and PAR2 peptides vs. PAR1 peptide alone. C: HUVECs were left unstimulated in plain medium or treated with TFLLR-NH2 (TF) or LSIGRL-NH2 (LS) at the concentrations shown in the presence or absence of the aminopeptidase inhibitor amastatin (Ama; 10 µM) for 20 h before being analyzed for DAF expression using flow cytometry. Data are expressed as percentages of DAF expression in US cells ± SE (n = 3 experiments with separate ECs). *P < 0.05 for ECs treated with PAR1 vs. US or Ama alone. D: HUVECs were pretreated with TNF-{alpha} (10 ng/ml) (filled bars) or plain HUVEC medium alone (open bars) for 24 h. HUVECs were then treated with thrombin (Thr), SFLLRN (SF), TFLLR-NH2 (TF), SLIGKV (SL), or LSIGRL-NH2 (LS) or were left unstimulated for a further 20 h before being analyzed for DAF expression using flow cytometry. Data are expressed as percentages of DAF expression on US cells ± SE (n = 4 experiments with separate ECs). *P < 0.05 for ECs treated with TNF-{alpha} vs. ECs pretreated with plain HUVEC medium alone.

 
Dose-response and time-course experiments demonstrated a significant upregulation of DAF expression by PAR1 and PAR2 peptides 20 h posttreatment, with responses first observed at 100 µM and maximally at 400 µM (Table 1). The relatively high concentration of peptide required to induce a maximal response reflects the prolonged time course required for the induction of surface proteins and the probable degradation of the peptide during this time. We therefore looked at the concentrations of the PAR peptides required to induce DAF expression in the presence of the aminopeptidase inhibitor amastatin. Amastatin alone and the control peptide LSIGRL-NH2 had no effect on basal DAF expression. However, amastatin reduced the concentration of TFLLR-NH2 required to induce a significant upregulation of DAF from 400 to 4 µM (Fig. 3C) and that of SFLLRN from 400 to 100 µM (Table 1).


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Table 1. Amastatin reduces the concentration of PAR-activating peptide required to induce DAF expression

 
The expression of PAR2, but not of PAR1, is regulated by TNF-{alpha}, with increased levels of protein detected 24 h poststimulation (46). To investigate the effect of PAR2 upregulation on the subsequent induction of DAF by thrombin and the PAR2 peptide, HUVEC monolayers were stimulated with TNF-{alpha} for 24 h before being treated with control peptide LSIGRL-NH2, PAR peptides, or thrombin for another 20 h. Flow cytometric analysis revealed that basal DAF expression was not significantly altered by TNF-{alpha} pretreatment for 24 h (Fig. 3D). However, DAF upregulation in response to thrombin was significantly increased. At the same time, although DAF induction by the PAR1 peptides SFLLRN and TFLLR-NH2 was not enhanced by pretreatment with TNF-{alpha}, the response to ligation of PAR2 by SLIGKV was significantly increased (Fig. 3D). The response to the combination of PAR1 and PAR2 peptides was also significantly enhanced by pretreatment with TNF-{alpha} (data not shown). These data suggest that the induction of PAR2 surface expression is responsible for the increased DAF upregulation observed in response to thrombin after pretreatment with TNF-{alpha}. Moreover, the failure of TNF-{alpha} pretreatment to increase the response observed with SFLLRN further supports the conclusion that SFLLRN acts in a PAR1-specific manner in this in vitro assay.

To investigate the possibility that thrombin-mediated activation of pro-MMP is involved in DAF upregulation (29), further experiments were performed in the presence of the MMP antagonist GM 6001 and its matched negative control. At concentrations up to 1 µM, GM 6001 failed to inhibit thrombin-induced DAF expression with RFI ± SE for untreated ECs, 26.5 ± 1.3; for thrombin-treated ECs, 63.3 ± 3.0; for thrombin with GM 6001, 61.5 ± 2.5; and for thrombin with GM 6001 negative control, 62.0 ± 2.7. The activity of GM 6001 was confirmed by its ability, at all concentrations used, to inhibit EC MMP-2 activation by phorbol dibutyrate as assessed using gelatin zymography (data not shown).

PAR1 and PAR2 use distinct PKC isozymes for DAF upregulation. Dominant-negative adenoviral constructs were used to further investigate the role of PKC isozymes downstream of PAR1 and PAR2 in DAF regulation. HUVECs were incubated with Adv for 2 h in serum-free medium before culture overnight in HUVEC medium supplemented with TNF-{alpha} to upregulate PAR2 expression as appropriate. ECs were then exposed to thrombin, SFLLRN, and SLIGKV, alone or in combination, and DAF expression was analyzed using flow cytometry after an additional 20 h. TNF-{alpha} alone had no significant effect on DAF expression (data not shown). Thrombin and PAR2-induced DAF upregulation were significantly reduced by DN-PKC-{epsilon} (Fig. 4A). Surprisingly, however, the induction of DAF by the PAR1 peptide was not significantly inhibited, suggesting the use of an alternative PKC isozyme. In subsequent experiments, we sought to identify the PKC isozymes involved in PAR1-mediated upregulation of DAF. Significant inhibition of this response was observed after expression of the DN-PKC-{alpha} construct in HUVECs (Fig. 4B). In contrast, no reduction of PAR2-induced DAF was observed. For all experiments, the data are presented as a percentage of appropriately matched controls to account for any changes in basal DAF expression. In addition, a {beta}-gal control Adv was included that had no consistent effect on either basal or induced DAF expression. The results of these experiments provide further evidence for a role for PAR2, as well as for PAR1, in the regulation of DAF expression by thrombin.



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Fig. 4. Role of PKC-{alpha} and PKC-{epsilon} in PAR peptide-induced DAF expression. HUVECs were infected with Adv (200 IFU/cell) expressing DN-PKC-{epsilon} (A; gray bars), DN-PKC-{alpha} (B; gray bars), or {beta}-gal control Adv (filled bars), or they were left untreated (open bars). After 24 h, ECs were treated with 10 U/ml thrombin, SFLLRN (SF), or SLIGKV (SL) (400 µM), or they were left unstimulated for 20 h. To optimize the DAF response induced by SLIGKV, relevant wells and matched controls were pretreated with TNF-{alpha} to induce PAR2 expression. DAF expression was analyzed using flow cytometry. To account for any effect of Adv on constitutive DAF expression, data are shown as percentages of DAF expression on the untreated, TNF-{alpha}-, or Adv alone-treated ECs as appropriate and presented as RFI ± SE (n = 5 experiments with separate ECs). *P < 0.05.

 
Thrombin-induced upregulation of DAF involves transactivation of PAR2 by PAR1. We next sought to establish whether the transactivation of PAR2 by PAR1, first proposed by O'Brien et al. (47), plays a role in DAF induction by thrombin. To investigate the role of PAR1, we used a combination of MAbs, WEDE15 and ATAP2, to block the binding of thrombin to PAR1 and thus prevent cleavage (47). These MAbs inhibited thrombin-induced DAF expression on ECs, whereas treatment with an isotype-matched control MAb (1.2B6) had no effect (Fig. 5A). Furthermore, the leech product hirudin, which inhibits the proteolytic action of thrombin, thus preventing both activation of PAR1 and transactivation of PAR2, completely inhibited DAF upregulation by thrombin (data not shown) as we previously reported (30). A combination of WEDE15 and ATAP2 also inhibited thrombin-induced DAF expression in ECs pretreated for 24 h with TNF-{alpha} to induce PAR2 expression (Fig. 5B). Treatment of ECs with MAbs alone had no effect on basal DAF expression (data not shown).



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Fig. 5. Effects of inhibition of PAR1 binding and signaling on DAF induction. A: HUVECs were preincubated with plain medium, PAR1-blocking MAbs WEDE15 and ATAP2 (W/A), or isotype-matched control MAb 1.2B6 for 30 min before being treated with 10 U/ml thrombin or plain medium alone for 16 h. B: HUVECs were treated with TNF-{alpha} (10 ng/ml) for 24 h before being preincubated with plain medium, PAR1-blocking MAbs WEDE15 and ATAP2 (W/A), or isotype-matched control MAb 1.2B6 for 30 min. This procedure was followed by the addition of 10 U/ml thrombin or carrier control and 16 h of further incubation. DAF expression was analyzed using flow cytometry with biotinylated MAb IA10. Data are expressed as percentage increases in DAF expression above unstimulated ECs ± SE (n = 3 experiments with separate ECs). *P < 0.001 vs. thrombin-treated controls. C: HUVECs were preincubated with 50 µM BMS-200261 (BMS) or carrier control (US) for 1 h before the addition of 10 U/ml thrombin or SFLLRN (SF) for 20 h. *P < 0.05 vs. untreated ECs (US). D: HUVECs were preincubated with 3 µM SCH79797 (SCH) or carrier control (US) for 1 h before the addition of thrombin, TFLLR-NH2 (TF), SFLLRN (SF), or SLIGKV (SL) (400 µM) for 20 h. DAF expression was analyzed using flow cytometry with MAb 1H4. Data are expressed as percentages of DAF expression above US control ± SE (n = 3 experiments with separate ECs). *P < 0.05 vs. peptide alone.

 
To investigate a role for PAR1-mediated signaling in the thrombin induction of DAF, BMS-200261, a peptide mimetic of the PAR1 AP, was used (5). This compound prevents PAR1 signaling by thrombin or PAR1 APs via inhibition of the binding of the tethered ligand to the body of the receptor. Although the presence of BMS-200261 led to a small reduction in thrombin-induced DAF upregulation, this reduction failed to reach significance, whereas SFLLRN (PAR1)-induced expression was reduced to baseline (Fig. 5C). In contrast, BMS-200261 failed to inhibit the induction of DAF expression by the PAR2 agonist peptide SLIGKV on both resting ECs and those pretreated with TNF-{alpha} (data not shown). Because BMS-200261 may activate PAR2, further experiments were performed with a PAR1-selective nonpeptide antagonist, SCH79797 (1). Although SCH79797 significantly inhibited DAF upregulation induced by PAR1 agonist TFLLR-NH2, it had no significant effect on either thrombin-induced or PAR2 peptide-induced responses (Fig. 5D). Taken together, our data strongly suggest that DAF upregulation by thrombin is not mediated by signaling pathways downstream of PAR1 but is mediated via an alternative mechanism that is likely to involve cleavage of PAR1 and subsequent transactivation of PAR2 by the PAR1-tethered ligand.

Thrombin-induced MAPK pathways. Thrombin may activate ERK1/2, p38, and JNK MAPK. However, the relationship between these pathways is complex, may vary between different cell types, and remains to be defined fully (50). In previous work, we demonstrated that thrombin-induced DAF expression is abrogated by pretreatment with pharmacological inhibitors of MEK-1 (PD-98059 or U0126), which prevent ERK1/2 activation, and by SB-202190, which inhibits p38 MAPK (30). Furthermore, as shown in Fig. 6A, pretreatment of HUVECs with pharmacological antagonists of JNK, JNK inhibitor II (SP-600125), and JNK inhibitor I, a cell-permeable peptide inhibitor, significantly inhibited thrombin-induced DAF expression. In addition, the ERK1/2, p38, and JNK MAPK activation inhibitors also abrogated the DAF upregulation observed after treatment with the PAR1 and PAR2 APs (data not shown).



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Fig. 6. MAPK phosphorylation by thrombin. A: HUVEC monolayers were preincubated with JNK inhibitor I peptide (25 µM; JI-1 pep) or JNK inhibitor II (SP-600125; JNKII) (50 µM) for 1 h. HUVECs were then left unstimulated (Unstim) in plain HUVEC medium (open bars) or were treated with 10 U/ml thrombin (filled bars) and cultured for 20 h before DAF expression was analyzed using flow cytometry. Data are expressed as percentages of thrombin-induced DAF expression ± SE (n = 3 experiments). *P < 0.05 and **P < 0.001 vs. thrombin-treated ECs. BD: HUVECs were preincubated with inhibitors for 1 h before stimulation with 10 U/ml thrombin for 7 min and then were lysed. Lysates were separated by SDS-PAGE, transblotted, and incubated with phosphorylation-specific antibodies against p38 MAPK, JNK, or ERK1/2. B: HUVECs were preincubated with plain medium alone (US) (lanes 1 and 2) or GF-109203X (GF) (5 µM; lanes 3 and 4) and stimulated with thrombin (lanes 2 and 4). C: HUVECs were preincubated with plain medium alone (US) (lanes 1 and 2), U0126 (U0) (1 µM; lanes 3 and 4), or GF-109203X (5 µM; lanes 5 and 6) and stimulated with thrombin (lanes 2, 4, and 6). D: HUVECs were preincubated with plain medium alone (US) (lanes 1 and 2), U0126 (1 µM; lanes 3 and 4), SB-202190 (SB) (25 µM; lanes 5 and 6), JNK inhibitor II (SP-600125, JNK, 50 µM; lanes 7 and 8) or GF-109203X (5 µM; lanes 9 and 10) and stimulated with thrombin (lanes 2, 4, 6, 8, and 10). Results are representative of 3 experiments and were quantified using densitometry with integrated density values (IDV) of phosphorylation-specific bands expressed as percentages of those obtained with antibodies against nonphosphorylated MAPKs.

 
Western blot analysis using phosphorylation-specific antibodies was performed to explore the relationship between the signaling pathways downstream of thrombin, PAR1, and PAR2 in ECs. As shown in Fig. 6, ERK1/2, p38, and JNK MAPK were rapidly phosphorylated by thrombin. However, these responses were not prevented by the inhibition of PKC with GF-109203X. Thrombin-induced phosphorylation of the MAPK enzymes was inhibited by specific antagonists of their activation, ERK1/2 (U0126) (Fig. 6D), p38 (SB-202190) (data not shown), and JNK (SP-600125 and JNK inhibitor I peptide) (data not shown). However, thrombin-induced JNK phosphorylation was also sensitive to the inhibition of ERK1/2 activation by U0126 (Fig. 6C), and, likewise, ERK1/2 phosphorylation was inhibited by the JNK antagonists (Fig. 6D). Despite the potential limitations of pharmacological antagonists in terms of strict target selectivity, these data may be interpreted as demonstrating the interdependence of JNK and ERK1/2, an observation that has not been reported previously.


    DISCUSSION
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 DISCUSSION
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Thrombin exerts pleiotropic effects on vascular endothelium. Many of these effects are considered proinflammatory, such as increased vascular permeability (33) and the induction of cellular adhesion molecules and leukocyte adhesion (24, 25, 63). Moreover, the detection of thrombin at sites of inflammation, including atherosclerotic plaques (44), areas of vascular injury (18), and the rheumatoid synovium (48), has often led to consideration of thrombin as a deleterious influence in these settings. However, thrombin may also induce antiapoptotic genes (6, 54), upregulate VEGF receptor expression (63), and induce local secretion of VEGF by ECs (53), resulting in cytoprotective and proangiogenic effects. Thus the role of thrombin at sites of inflammation appears to be complex, on the one hand inducing proinflammatory genes and on the other enhancing mechanisms for vascular protection.

The complement-inhibitory protein DAF interferes with the pivotal step of the complement cascade, in which an absolute excess of C3 is required for further activation of the cascade. We have previously shown that expression of DAF on human ECs is increased up to threefold by thrombin, resulting in a significant reduction in C3 deposition and subsequent complement-mediated lysis, thus providing enhanced resistance against complement-mediated injury (30). These observations, and the data presented herein, indicate the presence of an additional protective mechanism activated by thrombin and mediated by the transactivation of PAR2 and signaling via PKC-{epsilon}. However, the net effect of thrombin on the vascular endothelium is influenced by a variety of factors, including the local microenvironment and the balance between proinflammatory and cytoprotective pathways (Fig. 7).



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Fig. 7. Schematic of proposed mechanism for upregulation of DAF by thrombin. Thrombin is able to bind PAR1 but not PAR2. Binding to PAR1 cleaves the receptor and exposes a tethered ligand that activates a PAR1-PKC-{alpha} signaling pathway, which results in increased vascular permeability (39) and monocyte chemoattractant protein (MCP-1) production (54). The PAR1-tethered ligand may also transactivate PAR2, resulting in activation of PKC-{epsilon}, upregulation of DAF, and protection against complement-mediated injury.

 
PKC consists of three families of isozymes: cPKC, nPKC, and atypical PKC. The role of PKC in DAF regulation by thrombin was initially confirmed by using pharmacological antagonists and myristoylated pseudosubstrate peptides, which indicated a role for PKC-{epsilon} and, to a lesser extent, PKC-{alpha}. Taking into account concerns regarding the specificity of pharmacological antagonists, we sought to establish the roles of PKC-{alpha} and PKC-{epsilon} by using validated DN constructs expressed in adenoviruses. This approach confirmed the roles of these isozymes, with nearly complete inhibition of thrombin-induced DAF upregulation by DN-PKC-{epsilon} and only partial inhibition by DN-PKC-{alpha}.

The importance of the MAPK enzymes ERK1/2, p38, and JNK in thrombin signaling pathways led us to explore further their role in thrombin-induced DAF regulation and their relationship to PKC. Thrombin induced the phosphorylation of these MAPKs in HUVECs, and inhibition of the activation of JNK, ERK1/2, and p38 MAPK prevented DAF induction (30). Cross-inhibition studies revealed interdependence between ERK1/2 and JNK because inhibition of either kinase also prevented the activation of the other by thrombin. Although we cannot completely exclude a lack of specificity, the antagonists of ERK1/2 and JNK activation have not been reported to inhibit other MAPK enzymes significantly at the concentrations used. Moreover, evidence for complex, activating, or inhibitory cross-talk pathways between MAPK enzymes has been identified previously. Inhibition of ERK1/2 also prevents VEGF and thrombin-mediated activation of JNK in ECs (50). Furthermore, cross-talk between JNK and ERK1/2, and between ERK1/2 and p38 MAPK, has been reported in cellular responses to IL-1{beta} and TNF-{alpha}, respectively (21, 58).

Our data gathered to date show that inhibition of either the PKC or MAPK pathway is sufficient to prevent thrombin-induced DAF upregulation, suggesting that PKC and MAPK form part of a linear signaling pathway similar to that described for the induction of ICAM-1 by thrombin (52). We show herein that the overexpression of a constitutively active form of PKC-{epsilon} capable of ERK1/2 activation (19) also induces a significant increase in DAF expression, which is inhibited by the MEK-1 antagonist U0126. However, the broad spectrum PKC antagonist GF-109203X failed to inhibit thrombin-induced phosphorylation of the MAPK enzymes. This may reflect the inability of GF-109203X to inhibit atypical isozymes such as PKC-{zeta}. Alternatively, PKC and MAPK may not exhibit a simple linear relationship in this setting, and the pathways may integrate downstream or activate distinct transcription factors involved in DAF regulation.

HUVECs express PAR1, PAR2, and PAR3, but not PAR4 (47, 55), which can be detected in human arterial ECs (15). Although PAR1 is the predominant thrombin receptor on vascular endothelium, it remains to be determined whether PAR1 is responsible for all thrombin-mediated signaling in human ECs. In murine ECs, PAR1 and PAR4 may contribute to thrombin signaling (26). However, no role for PAR4 in thrombin-mediated responses has been found in human ECs (15). PAR2, which is expressed in human ECs and upregulated by proinflammatory cytokines (46), does not bind thrombin and may be activated by trypsin and mast cell tryptase. Early studies demonstrating the failure of PAR1-blocking antibodies to completely inhibit thrombin responses suggested the involvement of additional receptors for thrombin (3). Moreover, the response of HUVECs to thrombin can be reduced when PAR2 is first activated with a selective agonist peptide (42, 43). O'Brien et al. (47) subsequently reported that when PAR1 signaling is inhibited, PAR2 may be transactivated by cleaved PAR1, thereby providing a mechanism for indirect activation of PAR2 by thrombin. We sought to extend these findings by looking for evidence of a role for PAR2 transactivation in thrombin-mediated DAF upregulation in HUVECs.

PAR1 and PAR2 APs induced DAF expression in HUVECs, whereas peptides targeting PAR3 (data not shown) and PAR4 had no effect. The relatively high concentration of PAR1 and PAR2 peptides required to induce a response is likely to be a consequence of the long time course of the assay and the presence of associated peptide degradation. The latter hypothesis was confirmed by the ability of the aminopeptidase inhibitor amastatin to reduce substantially the concentration of TFLLR-NH2 required to induce DAF expression. The upregulation of DAF by PAR1 and PAR2 peptides in combination was equivalent to that observed with thrombin alone. In addition, the response to thrombin and the PAR2 peptide was enhanced by pretreating ECs with TNF-{alpha} to upregulate PAR2 expression before exposure to the AP. These data suggest that PAR2 might play a role in thrombin-induced DAF expression.

In the absence of known antagonists for PAR2, we used the PAR1 signaling antagonists BMS-200261 and SCH79797 to further investigate the hypothesis that transactivation of PAR2 by thrombin-cleaved PAR1 contributes to DAF upregulation. BMS-200261 and SCH79797 prevent binding of the PAR1-tethered ligand to the receptor without interfering with signaling via PAR2 or transactivation of PAR2 by PAR1 (1, 5, 47). As previously reported for thrombin-induced Ca2+ flux in ECs (47), inhibition of PAR1 with BMS-200261 or SCH79797 failed to significantly inhibit thrombin-mediated DAF upregulation while it completely prevented the upregulation by the PAR1 agonists SFLLRN and TFLLR-NH2. In contrast, hirudin, which binds thrombin and inhibits its proteolytic action, and the MAbs WEDE15 and ATAP2, which inhibit thrombin-mediated PAR1 cleavage, completely abrogated thrombin-induced DAF upregulation, whereas the MMP antagonist GM 6001 had no effect. SFLLR-derived peptides at the concentrations used can activate PAR2 in addition to PAR1 (2, 27). We therefore compared the responses observed with SFLLRN to those obtained with TFLLR-NH2, which has more selectivity for PAR1 than for PAR2 (27). SFLLRN and TFLLR-NH2 had directly comparable effects on DAF induction, which were inhibited by the PAR1 antagonists BMS-200261 and SCH79797. In addition, pretreatment of ECs with TNF-{alpha} to increase PAR2 expression had no effect on the upregulation of DAF by SFLLRN. These data suggest that in this in vitro system, SFLLRN acts in a PAR1-specific manner and that any binding to PAR2 by the soluble peptide is less efficient than that of the orientation adapted by the tethered ligand itself.

Taken as a whole, our data suggest that thrombin cleaves PAR1, which transactivates PAR2, resulting in increased DAF expression to levels that we have previously shown to be capable of enhancing protection against complement-mediated injury (30). Although in the absence of specific PAR2 antagonists we cannot entirely exclude a role for other as yet unidentified EC thrombin receptors requiring PAR1 cleavage for their activation, we suggest that DAF upregulation is a functionally important outcome of the PAR1-mediated transactivation of PAR2 on ECs first proposed by O'Brien et al. (47).

Investigation of the signaling pathways downstream of PAR1 and PAR2 revealed no differences in the utilization of MAPK. However, the predominant PKC isozymes used by these receptors for the induction of DAF expression were different. Inhibition of PKC-{epsilon} with a DN-Adv construct significantly inhibited DAF upregulation by the PAR2 but not the PAR1 APs. In contrast, DN-PKC-{alpha} inhibited DAF induction downstream of PAR1 but not downstream from PAR2. Furthermore, the PAR1 signaling antagonists blocked the PAR1/PKC-{alpha}-dependent pathway but had no effect on PAR2/PKC-{epsilon}-induced DAF upregulation. Although PAR1 and PAR2 agonist peptides typically produce similar downstream responses, differences are emerging. Thus MCP-1 mRNA (54) and increased vascular permeability (28, 66) are induced by PAR1 but not PAR2 agonists. This may reflect differences in the ability of PAR1 and PAR2 ligation to activate the PKC-{alpha} and Rho-GTPase signaling pathway involved in regulating vascular permeability (28, 39, 66), a hypothesis supported by the recent report that PAR1-induced activation of PKC-{alpha} is involved in P-selectin expression on microvascular ECs (14). The link between PAR2 and PKC-{epsilon} described herein is likely to be important in vascular cytoprotection, and it is relevant that PKC-{epsilon} has been associated with a variety of other cytoprotective molecules, including endothelial nitric oxide synthase and hemoxygenase-1 (51).

In vascular endothelium, PAR1 activation by thrombin helps to coordinate the response to tissue injury (9). This activation includes proinflammatory actions such as the upregulation of cellular adhesion molecules including E-selectin, VCAM-1, and ICAM-1 and the release of the chemokines IL-8 and MCP-1, which in turn direct leukocyte migration toward sites of inflammation (24, 25, 30, 41). However, PAR1 may also activate a variety of reparative responses, including the release of PDGF, connective tissue growth factor, and vascular remodeling (10, 16, 65). The role of PAR2 in this setting remains not fully understood, and the lack of developmental defects in PAR2-deficient mice suggests that its role may be more restricted than that of PAR1 (10, 65). Ligation of PAR2 can induce neurogenic and cutaneous inflammation (22, 56, 61). Moreover, PAR2-deficient mice are protected against collagen-induced arthritis (12). Notwithstanding this finding, PAR2 may also exert anti-inflammatory, cytoprotective effects in a murine model of colitis and can act to reduce bronchoconstriction and induce angiogenesis, thus accelerating recovery in the ischemic hindlimb (4, 7, 13, 40). This suggests that activation of PAR2, although it is important in the initiation of inflammation, may also facilitate the resolution of inflammation, the initiation of repair, and the protection of surrounding tissues (64).

We propose that thrombin may utilize the transactivation of PAR2 by PAR1 even when PAR1 signaling is intact and subsequently activate a PKC-{epsilon}/MAPK-regulated pathway to increase the expression of the complement-inhibitory protein DAF and thus help to maintain vascular integrity. Moreover, the ability of TNF-{alpha} to increase PAR2 expression, thus enhancing DAF upregulation by thrombin, suggests that during inflammation, when the risk of autologous tissue damage by complement is greatest, cytoprotection is maximal. Our data emphasize the importance of establishing a detailed understanding of thrombin- and/or PAR-mediated signaling pathways, which may in turn help to identify novel targets through which vascular inflammation can be controlled and vascular cytoprotection can be preserved.


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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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This study was funded by the British Heart Foundation and the Arthritis Research Campaign. J. C. Mason is an Arthritis Research Campaign senior fellow.


    ACKNOWLEDGMENTS
 
We thank Dr. S. Traynelis for the generous gift of BMS-200261 and Dr. H. Yarwood for performing gelatin zymography.


    FOOTNOTES
 

Address for reprint requests and other correspondence: J. C. Mason, Cardiovascular Medicine Unit, Imperial College, Hammersmith Hospital, DuCane Road, London W12 ONN, UK (e-mail: justin.mason{at}imperial.ac.uk)

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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