Published online before print January 23, 2003, doi: 10.1161/01.RES.0000057386.15390.A3
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
© 2003 American Heart Association, Inc.
Molecular Medicine |
Differential Actions of PAR2 and PAR1 in Stimulating Human Endothelial Cell Exocytosis and Permeability
The Role of Rho-GTPases
From the Department of Medicine (S.W.K., A.C., R.C.N., A.G.M.), University of Alberta, Edmonton, Alberta; and the Departments of Pharmacology & Therapeutics and Medicine (M.D.H.), University of Calgary, Calgary, Alberta, Canada.
Correspondence to Allan G. Murray, Dept of Medicine, Rm 260F HMRC, University of Alberta, Edmonton, AB, Canada T6G 2S2. E-mail allan.murray{at}ualberta.ca
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Endothelial cell proteinase activated receptors (PARs) belong to a family of heterotrimeric G protein-coupled receptors that are implicated in leukocyte accumulation and potentiation of reperfusion injury. We characterized the effect and the signal transduction pathways recruited after stimulation of endothelial PAR2. We used von Willebrand Factor (vWF) release and monolayer permeability to peroxidase to report Weibel-Palade body (WPB) exocytosis and pore formation, respectively. Human umbilical vein endothelial cells (HUVECs) were stimulated with the selective PAR2 agonist peptide SLIGRL-NH2 or PAR1 agonist peptide TFLLR-NH2. PAR2 stimulation resulted in WPB exocytosis like PAR1 stimulation but, unlike PAR1, failed to increase monolayer permeability. BAPTA-AM inhibited PAR2-induced exocytosis, indicating a PAR2 calcium-dependent signal in ECs. Moreover, PAR2-like PAR1-stimulated exocytosis requires actin cytoskeleton remodeling, because vWF release is inhibited if the cells were pretreated with Jasplakinolide. Rho-GTPase activity is required for PAR-stimulated exocytosis, because inactivation of this family of actin-regulatory proteins with Clostridium difficile toxin B blocked exocytosis. Expression of dominant-negative mutant Cdc4217N inhibited exocytosis whereas neither dominant-negative Rac17N expression nor C3 exotoxin treatment affected vWF release. PAR2 stimulated RhoA-GTP weakly compared with the PAR1 agonist. We conclude that both PAR2 and PAR1 elicit WP body exocytosis in a calcium and Cdc42 GTPase-dependent manner. In contrast, the differential effect of PAR1 versus PAR2 activation to increase monolayer permeability correlates with weak RhoA activation by the PAR2 agonist.
Key Words: vascular endothelium • reperfusion injury • thrombin receptors • Rho-GTP-binding proteins
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Reperfusion of tissue after revascularization of an arterial thrombosis or preservation of a solid-organ allograft is accompanied by microvascular endothelial cell injury, local activation of the coagulation cascade, and diminished endothelial anticoagulant properties.1–3 In addition, platelet and leukocyte adhesion to the endothelium is thought to contribute to inhomogeneous perfusion and persistent ischemic injury of the parenchymal cells of the tissue.4–6 Activation of specific receptors on the microvascular endothelial cells by the serine proteinases of the coagulation cascade appears to be an important mechanistic link between tissue injury and the early recruitment of leukocytes.2
The proteinase activated receptors (PARs) are a novel family of G protein-coupled receptors that require proteolytic cleavage at the amino terminus to stimulate signaling (see review7). Cleavage allows the newly exposed tethered ligand to initiate an intramolecular interaction necessary for signal transduction, which can be mimicked by peptide analogues of the newly exposed N-terminus. The PAR family members, PAR1–4, are expressed constitutively on endothelial cells, and PAR2 and PAR4 are further upregulated on inflamed endothelium.8,9 PAR1, PAR3, and PAR4 are substrates for thrombin, whereas PAR2 can be stimulated by the activity of tryptase, a product of mast cell degranulation, and coagulation Factors VIIa and Xa.10,11
Stimulation of endothelial cells with thrombin or peptide analogues of the PAR1-tethered ligand (eg, TRAP or SFLLRN-NH2), results in reorganization of the cytoskeleton and discharge of endothelial storage granules, or Weibel-Palade bodies (WPBs). Exocytosis of WPBs plays a major role in the recruitment of leukocytes to sites of inflammation.12 WP body exocytosis releases stored IL-8 and mobilizes the adhesion molecule P-selectin to the lumenal surface of the endothelial cell.13,14 In addition, WPB exocytosis promotes platelet adhesion through the release of von Willebrand’s factor (vWF).13 Thrombin stimulation elicits capillary leak in vivo and increased endothelial monolayer permeability in vitro.15,16 PAR stimulation, then, promotes leukocyte adhesion to vascular endothelium and links inflammation and coagulation in a variety of pathological settings.
Stimulation of the PAR2 is also proinflammatory in vivo. Injection of selective peptide analogues of the PAR2 receptor tethered ligand is sufficient to initiate inflammation in vivo.17 Moreover, impaired inflammatory responses are observed in the PAR2-deficient mouse.18 In vitro, trypsin, like thrombin or TRAP, treatment elicits WPB exocytosis,19 but fails to induce monolayer permeability.20 However, the proteinases may act on surface proteins other than PARs,21 and the PAR1 agonist peptides modeled on the PAR1 tethered ligand activate both PAR1 and PAR2.22 Hence, characterization of the effects of selective endothelial PAR2 stimulation, and the signaling pathways recruited by PAR2 agonists to stimulate the endothelial cell remain to be explored.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Reagents
PAR agonist peptides (AP) TFFLR-NH2, and SLIGRL-NH2 were obtained from the peptide synthesis facility of the Health Sciences Center of the University of Calgary or from the Alberta Peptide Institute and were >95% pure by HPLC. Jasplakinolide was obtained from Molecular Probes. ELISA reagents for vWF quantitation, were obtained from Cedarlane Laboratories, and a vWF standard through Precision Biologic. Ionomycin and C3 exotoxin were obtained from Calbiochem. The rho-kinase inhibitor Y-27632 was a gift of Welfide Corporation (Osaka, Japan). The Clostridium difficile toxin B (TcdB) was obtained from List Biochemicals. The rhotekin Rho binding domain (RBD)-GST fusion protein cDNA was a generous gift of Dr M. Schwartz (Scripps Research Institute, San Diego, Calif). Anti-RhoA mAb 26C4, and phospho-specific myosin light chain kinase pAb, were obtained from Santa Cruz Biotechnology. Peroxidase-conjugated secondary pAb were from Jackson Immunoresearch. M199, FBS, and Hanks balanced salt solution were from Invitrogen. Cell dissociation solution, cytochalasin D, H-7, and all other reagents were from Sigma.
Endothelial Cell Culture
Human umbilical vein endothelial cells (HUVECs) were isolated from several umbilical cords, pooled, and cultured in complete media (M199 with 20% FBS, penicillin, streptomycin, and glutamine).23 Cells were serially passed in complete media supplemented with ECGS. HUVECs were cultured on a gelatin matrix and used in the experiments at passage 2 to 5. The passaged endothelial cells were harvested by washing twice with Hanks’, then incubated with a nonenzymatic Cell Dissociation Solution (Sigma) until the ECs were seen to lift off the plate. For use in an experiment, the harvested cells were then replated onto fibronectin (Fn)-coated C6 wells and incubated at 37°C in complete medium for 24 to 48 hours until confluent.
Where indicated, Cytochalasin D (1 µmol/L), jasplakinolide (45 nmol/L), or DMSO vehicle were diluted in supplemented M199 and added to the existing media. After 1 hour of incubation at 37°C, the media was removed and discarded, and fresh complete media with the agonist or vehicle diluted to the desired concentration was applied to the HUVECs. To inhibit Rho-GTPases, HUVECs were pretreated with 50 µg/mL C3 exotoxin for 24 hours and a mobility shift of endothelial Rho on SDS-PAGE was confirmed as described previously.24 To inhibit rho-kinase, endothelial cells were pretreated with H-7 (0.5 µmol/L) or Y-27632 (15 µmol/L) for 120 minutes before stimulation. To stimulate WP body exocytosis, the following agonists were used: phorbal 12-myristate 13-acetate (PMA) at 100 nmol/L, PAR1 AP (TFLLR-NH2) at 30 µmol/L, PAR2 AP (SLIGRL-NH2) at 30 µmol/L, and Ionomycin at 1 µmol/L, except where otherwise specified.
Chemical inhibitors were used at concentrations that maintained HUVEC viability >85% of mock-treated controls as assessed by the XTT assay of mitochondrial activity.25
ELISA Assay of vWF
Supernatant from mock- or agonist-stimulated HUVECs was removed into separate tubes and kept on ice until analysis. The vWF concentration was determined by ELISA. Calibration was performed with a vWF standard (Human reference plasma, Precision Biologics). Regulated vWF release was calculated by subtracting the vWF concentration in unstimulated wells from the stimulated wells. The effect of the inhibitors was determined by comparing the mean vWF release to its control group. The mean difference of the treatment among several experiments, as noted in the figure legends, was calculated and tested for statistical significance (P<0.05) by ANOVA using SPSS (SPSS).
Endothelial Monolayer Permeability
HUVECs at passage 3 were plated onto fibronectin-coated polycarbonate membranes (5-um pore size Transwell, Costar) at 6x104 cells/cm2 in M199/20% FBS. The monolayers were cultured for 5 days with media changes every 2 days and were not disturbed for 24 hours before the experiment. Horse radish peroxidase (HRP, MW 44kDa, Sigma) was added to the upper chamber at a final concentration of 1 µg/ml, then the specific peptide agonist was added at the concentration indicated in the figure legends. Where indicated, the monolayers were pretreated as described in the figure legend. Medium was harvested from the lower chamber after 20 minutes, and the HRP activity was determined colorimetrically by absorbance at 490 nm of the o-phenyl diamine reaction product. The mean HRP concentration from triplicate experimental wells was compared with the HRP concentration in wells treated with ionomycin 10 µmol/L according to the formula: (experimental-media control)/(ionomyin-media control)x100=% maximum permeability.
The difference of the means between treatment and controls among several experiments was calculated and tested for statistical significance (P<0.05) by ANOVA using SPSS.
Affinity Precipitation of Rho-GTP
The RBD-GST fusion protein was prepared and isolated using glutathione beads (Amersham, Baie d’Urfe, PQ), then used to affinity precipitate GTP-Rho as described previously.26 Rho was detected with anti-Rho mAb 26C4 then the blots were developed using chemiluminescence (Supersignal, Pierce). Images were acquired from film using a CCD camera and the density of the bands were quantitated using the Fluor-S Max software package (BioRad).
Transient Transfection of Endothelial Cells
Dominant-negative myc-tagged mutant Rac117N and Cdc4217N cDNA were generously provided by Dr Gary Bokoch (Scripps) then subcloned into pCSD7 (a gift of John Elliott, University of Alberta, Edmonton, Canada) for transfection of HUVECs. The cDNAs were sequenced to confirm the presence of the mutation before use. Briefly, HUVECs were plated at confluence on Fn matrix in C-6 plates in growth medium 1 day before transfection. The cells were washed with OptiMEM (Life Technologies), then 0.2 µg DNA/well was introduced into the cells using the Effectene kit (Qiagen) according to the instructions of the manufacturer. After 3 to 4 hours, the wells were washed and the media was replaced with M199/10% FBS and ECGS. Pilot experiments indicated that expression of the myc-tagged mutant GTPase was optimal at approximately 24 hours after transfection; therefore, the cells were used in the experiments as indicated at this time. Endothelial cell expression of the myc-tagged protein was confirmed in each experiment.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Weibel-Palade Body Exocytosis Is Stimulated by Activating Peptides Specific for PAR2
First, we sought to examine the effect of selective PAR2 activation compared with PAR1 activation on endothelial cell exocytosis. Cells were stimulated with PAR1 (TFLLR-NH2), or PAR2 (SLIGRL-NH2) agonist peptide, then the media was assayed for vWF release. The amount of agonist peptide-stimulated vWF release was compared with vWF release in parallel wells maximally stimulated with PMA. The response to PMA stimulation among different HUVEC cell lines was variable. Hence, to facilitate comparison among independent experiments, the effect of PAR agonist peptide was normalized to PMA-induced vWF release. Concentration-effect curves were generated for both PAR1 and PAR2 agonist peptides (Figure 1). The maximum release of vWF occurred at a concentration range of 40 to 60 µmol/L for either peptide. In this concentration range (ie, 50 µmol/L), TFLLR-NH2 selectively activates PAR1 and SLIGRL-NH2 selectively activates PAR2.22 No further increase of WP body exocytosis occurred with higher concentrations (up to 150 µmol/L, data not shown). After adjusting for constitutive release, PAR1 or PAR2 agonist peptide stimulation at 50 µmol/L led to vWF release of 1.1±0.1 (mean±SEM) U/mL and 1.1±0.1 U/mL of vWF, respectively. These values corresponded to 21%±2% (mean±SEM) of PMA-induced vWF release. Further experiments used the agonist peptide at a concentration (30 µmol/L) that approximates the EC50 for the response elicited by the peptides. These experiments confirm that either PAR2 or PAR1 can independently stimulate endothelial exocytosis.
|
PAR2 Activation, Unlike PAR1, Fails to Increase Monolayer Permeability
We next examined if selective PAR2 stimulation altered endothelial monolayer integrity. Like the PAR1 agonist, the PAR2 agonist peptide induced actin stress fiber formation in confluent HUVECs (data not shown). Notwithstanding, unlike PAR1, PAR2 stimulation failed to increase monolayer permeability (Figure 2). Ionomycin stimulation elicited a marked increase in transfer of HRP across the monolayer (18±6 versus 68±7 ng/cm2 per hour; mean±SEM) and served as a comparator to which the effects of the PAR agonists were normalized. These data indicate that although PAR2, like PAR1 stimulation, is able to induce bundled f-actin stress fiber formation, unlike PAR1, PAR2 fails to induce pore formation.
|
Effect of Cytochalasin D and Jasplakinolide on PAR2 Agonist Peptide-Stimulated Exocytosis
Both endothelial cell exocytosis and monolayer permeability changes are thought to require rearrangement of the f-actin cytoskeleton, hence we sought to determine if PAR2 stimulated f-actin reorganization. Cytochalasin D and jasplakinolide have been previously shown to depolymerize or stabilize the f-actin cytoskeleton, respectively.27,28 To determine if the endothelial cytoskeleton was reorganized in PAR2-stimulated regulated exocytosis, HUVECs were pretreated with either agent, then treated or not with peptide agonist. Neither agent changed constitutive vWF release. However, cytochalasin D pretreatment led to a modest augmentation of both PAR2- and PAR1-regulated WP body exocytosis (Figure 3). These data indicate that an intact cytoskeleton is not required for PAR-stimulated WP body exocytosis.
|
Jasplakinolide was used to test the requirements of each agonist for remodeling of the intact cytoskeleton to elicit vWF release (Figure 3). HUVECs pretreated with jasplakinolide then stimulated with PAR1 agonist peptide, TFLLR-NH2, released 54±15% of vWF compared with mock-pretreated cells (P=0.03, n=6 experiments). Similarly, jasplakinolide inhibited vWF release on stimulation with PAR2 agonist peptide, SLIGRL-NH2, to 43±16% of mock-pretreatment release (P=0.05, n=5 experiments). These data indicate that PAR2-mediated exocytosis requires active remodeling of the endothelial f-actin cytoskeleton.
PAR2 Exocytosis Requires Intracellular Calcium
The initiation of a calcium flux has previously been identified to play a critical role in thrombin-stimulated WP body exocytosis and monolayer permeability.29,30 To determine if PAR2-stimulated exocytosis was similarly dependent on calcium-dependent signaling, HUVEC monolayers were pretreated with BAPTA-AM, then stimulated with the PAR agonist peptides. As noted in Figure 4A, PAR2-stimulated vWF release was markedly inhibited under these conditions. This indicates that WP body exocytosis, stimulated by either PAR1 or PAR2, is dependent on elevated intracellular calcium concentration.
|
Our data suggest that both cytoskeletal reorganization and elevated intracellular calcium are required for efficient PAR-stimulated exocytosis. However, high concentrations of jasplakinolide may inhibit sustained intracellular calcium levels after stimulation via G protein-coupled receptors.31 Because a calcium flux is sufficient to induce WP body exocytosis,29 we sought to determine if calcium ionophore-induced vWF release was affected by jasplakinolide. First, HUVECs were stimulated with ionomycin to define a concentration-effect curve (data not shown). Ionomycin (1 µmol/L) induced vWF release comparable to that of the PAR-activating peptides at 30 to 50 µmol/L. Jasplakinolide pretreatment diminished ionomycin-induced vWF release to 52±13% (Figure 4B; P=0.02, n=5 experiments) of mock-pretreated controls. This observation indicates that the effect of jasplakinolide lies downstream of the generation of a calcium flux and suggests that PAR2 stimulation results in a calcium-dependent reorganization of the endothelial cytoskeleton.
Involvement of Rho-GTPase Signaling After PAR2 Stimulation
The Rho family of small GTP-binding proteins regulate the f-actin cytoskeleton and have been implicated in thrombin-stimulated increased endothelial monolayer permeability,16,32 and in exocytosis in a variety of other cell types.33,34 Rho kinase, a Rho-GTP-dependent serine/threonine kinase, acts to phosphorylate and inactivate myosin light chain phosphatase. Myosin light chain phosphorylation is also implicated in both thrombin-stimulated permeability and exocytosis.16,29 We therefore sought to determine if a Rho-GTPase-dependent signaling pathway was recruited after PAR2 stimulation.
We pretreated confluent HUVEC monolayers with TcdB toxin to selectively inhibit the small GTPases, Rho, Rac, and Cdc42. PAR2- and PAR1-mediated exocytosis were markedly inhibited (Figure 5A). This strongly suggests that Rho-GTPases serve a critically important function to effect PAR-stimulated exocytosis in endothelial cells.
|
To determine if RhoA activity elicited by PAR2 stimulation was involved in exocytosis, we tested the effect of specific inhibitors of the RhoA signaling pathway. First, we inhibited Rho activity with C3 exotoxin.24 C3 pretreatment did not inhibit vWF release using the PAR2 agonist peptide to stimulate endothelial cell exocytosis (Figure 5A). We similarly observed that inhibition of Rho-kinase with either Y-27632 or H-7 (data not shown) failed to inhibit, and tended to augment, PAR-stimulated vWF release. In contrast, PAR1 stimulation of monolayer permeability was markedly attenuated by inhibition of Rho kinase with Y-27632 (Figure 2). These data indicate that RhoA-GTPase signaling is not required for PAR2-stimulated exocytosis and suggest that other Rho-GTPase family members account for the TcdB-mediated inhibition.
To identify the Rho-GTPase family member involved in PAR-stimulated exocytosis, we expressed dominant-negative mutant Rac117N or Cdc4217N in the endothelial cells. Both PAR2- and PAR1-stimulated exocytosis was inhibited in HUVEC-expressing Cdc4217N but not Rac117N (Figure 5B). Taken together these observations indicate that stimulation of either endothelial PAR receptor recruits a signal that is dependent on Cdc42 activity to elicit exocytosis.
PAR2 Stimulation Elicits Weak RhoA Activity
PAR1-stimulated monolayer permeability depends on Rho kinase activity (Figure 2), hence we directly assessed RhoA activation after PAR2 stimulation to determine if the failure of PAR2 to elicit pore formation correlated with RhoA-GTP. As shown in Figure 6, RhoA-GTP was robustly activated by PAR1 stimulation, but only weakly by PAR2 stimulation at receptor-selective concentrations of the agonist peptides. To determine if the weak RhoA-GTP stimulation by PAR2 could account for the failure of pore formation through failing to increase cell contractility, we examined myosin light chain phosphorylation. Both PAR2 and PAR1 agonist peptides induced phosphorylation of myosin light chain, with a much greater effect of PAR1 (Figure 7). Interestingly, stimulation of MLC phosphorylation by either agonist peptide was inhibited by pretreatment of the HUVECs with Y27632 (data not shown). These observations indicate that the failure of PAR2 stimulation to elicit monolayer pore formation correlates with weak signaling through RhoA.
|
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
The major observations of this article are as follows: first, PAR2-stimulated exocytosis is dependent on both remodeling the actin cytoskeleton and eliciting an intracellular calcium flux. PAR2-stimulated exocytosis does not require either myosin light chain phosphorylation, or RhoA-GTPase-dependent Rho kinase activity. However, both PAR2 and PAR1-stimulated exocytosis is sensitive to TcdB inhibition of the Rho family of GTPases. Cdc42 signaling appears to be the principal Rho-GTPase required for endothelial Weibel-Palade body exocytosis. Interestingly, unlike PAR1, PAR2 failed to elicit increased endothelial monolayer permeability, despite generating stress fibers and myosin light chain phosphorylation to enable cell contractility. Activity of RhoA-GTP and myosin light chain phosphorylation are required for PAR1-mediated monolayer permeability.
We studied selective agonists of both the PAR2 and PAR1 G protein-coupled receptors and observed that each stimulated similar amounts of vWF release from the endothelial cell storage pool. The use of a selective activator of PAR2 indicates that the PAR2 system can have as great an impact on Weibel-Palade body exocytosis and hence lumenal display of P-selectin as the thrombin-regulated PAR1 system. Interestingly, like PAR1, PAR2-mediated exocytosis required f-actin reorganization, and involves a calcium-dependent rather than a cAMP-dependent second messenger. A similar requirement for f-actin reorganization was observed using ionomycin to mimic the calcium flux initiated by the PAR receptors, indicating that the requirement for f-actin remodeling likely occurred downstream of the PAR-stimulated calcium signal. These observations are in agreement with previous work that inferred activation of the PAR2 receptor using trypsin to stimulate Weibel-Palade body exocytosis19 and that detected a calcium flux in endothelial cells after trypsin stimulation.35
Coupling of PAR2 to endothelial cytoskeletal-associated proteins has not been examined previously. However, regulated exocytosis by mast cells or pancreatic acinar cells can be facilitated by dissolution of the cortical actin barrier and is thought to be a prerequisite for vesicle fusion to the plasma membrane.36,37 The endothelial PAR2 stimulated calcium signal is required for exocytosis and may regulate disassembly of cortical f-actin,38 for example by augmenting the activity of actin-severing proteins.39 The sensitivity of PAR2 stimulated exocytosis to the actin-stabilizing effect of jasplakinolide is consistent with this model.
Regulation of f-actin reorganization occurs under the influence of the Rho family of monomeric GTP-binding proteins.40 For example, Rac1 is linked to PAR signaling pathways that regulate stress fiber induction and cortical actin remodeling.32 In addition, GTP-
S-induced exocytosis in permeabilized mast cells is inhibited by C3 exotoxin, which specifically ADP-ribosylates and inactivates Rho.33 Rac and Cdc42 activity are also implicated in regulated exocytosis in basophilic leukemic cells and pancreatic ß-cells.34,41,42 Our observation that pretreatment of the HUVECs with TcdB, to inactivate Rho, Rac, and Cdc42, markedly inhibits PAR-stimulated vWF release, clearly implicating Rho-GTPases in both the PAR2 and PAR1 signaling pathways that direct Weibel-Palade body exocytosis.
Rho-GTP rapidly increases after stimulation of HUVECs with either the PAR2 or PAR1 agonist. To determine the participation of Rho directly in PAR-stimulated WP body exocytosis, we used the C3 exotoxin to selectively modify and abrogate interaction of Rho with downstream effector molecules. We found no consistent inhibitory effect of C3 exotoxin when used at concentrations known to markedly ADP-ribosylate endothelial Rho proteins.24 Moreover, inhibition of Rho kinase with Y27632 also fails to block PAR-stimulated exocytosis. This result indicates that neither RhoA nor Rho kinase activity is required for WP body exocytosis after stimulation of either PAR, and confirm recent data examining the role of RhoA in thrombin-stimulated exocytosis.43 In contrast, we observed that expression of the Cdc4217N mutant significantly decreased PAR-stimulated Weibel-Palade body exocytosis. Because the transfection efficiency in these experiments was approximately 30% to 50%, diminished Cdc42 activity likely accounts for much of the inhibition observed with TcdB treatment. Taken together these experiments implicate an obligatory role for Cdc42 activity in the cascade of events leading to endothelial exocytosis downstream of either proteinase activated receptor. However, we cannot exclude that Rho-GTPase family members function partially redundantly in this system.
Interestingly, despite the ability to mobilize intracellular calcium and elicit f-actin remodeling and myosin light chain phosphorylation, PAR2 signaling, in contrast with PAR1, fails to increase endothelial monolayer permeability. Our observations confirm and extend those of Compton et al20 who observed no endothelial monolayer permeability change after trypsin stimulation. We observe that PAR1-mediated monolayer permeability required Rho-GTP and Rho kinase activity for this event and demonstrate that PAR2 recruited Rho-GTP, albeit only weakly. These results indicate that unlike PAR1, PAR2 fails to sufficiently activate the RhoA, and possibly other, signal transduction pathways necessary for monolayer pore formation.
In summary, we describe a role for cytoskeleton remodeling and the Rho family of GTP-binding proteins in both PAR1 and PAR2 signaling to endothelial Weibel-Palade body exocytosis. The signal involves Cdc42 GTP-binding protein acting independently of Rho kinase or Rac1. Our data are consistent with a model of Cdc42 GTP-binding protein-dependent cortical cytoskeletal remodeling. However, in comparison with PAR1, PAR2 elicits less RhoA-GTP and MLC phosphorylation and does not elicit increased monolayer permeability.
|
Acknowledgments |
|---|
A.G.M. is the recipient of a Clinician Investigator award from the Alberta Heritage Foundation for Medical Research. This work was supported in part by an operating grant from the Kidney Foundation of Canada and Aventis/Canadian Blood Services Research Fund to A.G.M.
|
Footnotes |
|---|
*Both authors contributed equally to this study.
Received February 14, 2002; revision received December 20, 2002; accepted January 10, 2003.
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
1. Golino P, Ragni M, Cirillo P, Avvedimento VE, Feliciello A, Esposito N, Scognamiglio A, Trimarco B, Iaccarino G, Condorelli M, Chiariello M, Ambrosio G. Effects of tissue factor induced by oxygen free radicals on coronary flow during reperfusion. Nat Med. 1996; 2: 35–40.[CrossRef][Medline] [Order article via Infotrieve]
2. Erlich JH, Boyle EM, Labriola J, Kovacich JC, Santucci RA, Fearns C, Morgan EN, Yun W, Luther T, Kojikawa O, Martin TR, Pohlman TH, Verrier ED, Mackman N. Inhibition of the tissue factor-thrombin pathway limits infarct size after myocardial ischemia-reperfusion injury by reducing inflammation. Am J Pathol. 2000; 157: 1849–1862.
3. Ejiri S, Eguchi Y, Kishida A, Ishigami F, Kurumi Y, Tani T, Kodama M. Cellular distribution of thrombomodulin as an early marker for warm ischemic liver injury in porcine liver transplantation: protective effect of prostaglandin I2 analogue and tauroursodeoxycholic acid. Transplantation. 2001; 71: 721–726.[CrossRef][Medline] [Order article via Infotrieve]
4. Palazzo AJ, Jones SP, Girod WG, Anderson DC, Granger DN, Lefer DJ. Myocardial ischemia-reperfusion injury in CD18- and ICAM-1-deficient mice. Am J Physiol. 1998; 275: H2300–H2307.[Medline] [Order article via Infotrieve]
5. Yadav SS, Howell DN, Steeber DA, Harland RC, Tedder TF, Clavien PA. P-Selectin mediates reperfusion injury through neutrophil and platelet sequestration in the warm ischemic mouse liver. Hepatology. 1999; 29: 1494–1502.[CrossRef][Medline] [Order article via Infotrieve]
6. Singbartl K, Green SA, Ley K. Blocking P-selectin protects from ischemia/reperfusion-induced acute renal failure. FASEB J. 2000; 14: 48–54.
7. MacFarlane SR, Seatter MJ, Kanke T, Hunter GD, Plevin R. Proteinase-activated receptors. Pharmacol Rev. 2001; 53: 245–282.
8. Mirza H, Yatsula V, Bahou WF. The proteinase activated receptor-2 (PAR-2) mediates mitogenic responses in human vascular endothelial cells. J Clin Invest. 1996; 97: 1705–1714.[Medline] [Order article via Infotrieve]
9. Hamilton JR, Frauman AG, Cocks TM. Increased expression of protease-activated receptor-2 (PAR2) and PAR4 in human coronary artery by inflammatory stimuli unveils endothelium-dependent relaxations to PAR2 and PAR4 agonists. Circ Res. 2001; 89: 92–98.
10. Fox MT, Harriott P, Walker B, Stone SR. Identification of potential activators of proteinase-activated receptor-2. FEBS Lett. 1997; 417: 267–269.[CrossRef][Medline] [Order article via Infotrieve]
11. Riewald M, Ruf W. Mechanistic coupling of protease signaling and initiation of coagulation by tissue factor. Proc Natl Acad Sci U S A. 2001; 98: 7742–7747.
12. Robinson SD, Frenette PS, Rayburn H, Cummiskey M, Ullman-Cullere MD, Wagner DD, Hynes RO. Multiple, targeted deficiencies in selectins reveal a predominant role for P-selectin in leukocyte recruitment. Proc Natl Acad Sci U S A. 1999; 96: 11452–11457.
13. Hattori R, Hamilton KK, Fugate RD, McEver RP, Sims PJ. Stimulated secretion of endothelial von Willebrand factor is accompanied by rapid redistribution to the cell surface of the intracellular granule membrane protein GMP-140. J Biol Chem. 1989; 264: 7768–7771.
14. Wolff B, Burns AR, Middleton J, Rot A. Endothelial cell "memory" of inflammatory stimulation: human venular endothelial cells store interleukin 8 in Weibel-Palade bodies. J Exp Med. 1998; 188: 1757–1762.
15. Cirino G, Cicala C, Bucci MR, Sorrentino L, Maraganore JM, Stone SR. Thrombin functions as an inflammatory mediator through activation of its receptor. J Exp Med. 1996; 183: 821–827.
16. van Nieuw Amerongen GP, van Delft S, Vermeer MA, Collard JG, van Hinsbergh VW. Activation of RhoA by thrombin in endothelial hyperpermeability: role of Rho kinase and protein tyrosine kinases. Circ Res. 2000; 87: 335–340.
17. Vergnolle N, Hollenberg MD, Sharkey KA, Wallace JL. Characterization of the inflammatory response to proteinase-activated receptor-2 (PAR2)-activating peptides in the rat paw. Br J Pharmacol. 1999; 127: 1083–1090.[CrossRef][Medline] [Order article via Infotrieve]
18. Lindner JR, Kahn ML, Coughlan SR, Sambrano GR, Schauble E, Bernstein D, Foy D, Hafezi-Moghadam A, Ley K. Delayed onset of inflammation in protease-activated receptor-2-deficient mice. J Immunol. 2000; 165: 6505–6510.
19. Storck J, Kusters B, Vahland M, Morys-Wortmann C, Zimmermann ER. Trypsin induced von Willebrand factor release from human endothelial cells in mediated by PAR-2 activation. Thromb Res. 1996; 84: 463–473.[CrossRef][Medline] [Order article via Infotrieve]
20. Compton SJ, Cairns JA, Holgate ST, Walls AF. Human mast cell tryptase stimulates the release of an IL-8-dependent neutrophil chemotactic activity from human umbilical vein endothelial cells (HUVEC). Clin Exp Immunol. 2000; 121: 31–36.[CrossRef][Medline] [Order article via Infotrieve]
21. Lafleur MA, Hollenberg MD, Atkinson SJ, Knauper V, Murphy G, Edwards DR. Activation of pro-(matrix metalloproteinase-2) (pro-MMP-2) by thrombin is membrane-type-MMP-dependent in human umbilical vein endothelial cells and generates a distinct 63 kDa active species. Biochem J. 2001; 357: 107–115.[CrossRef][Medline] [Order article via Infotrieve]
22. Kawabata A, Saifeddine M, Al-Ani B, Leblond L, Hollenberg MD. Evaluation of proteinase-activated receptor-1 (PAR1) agonists and antagonists using a cultured cell receptor desensitization assay: activation of PAR2 by PAR1-targeted ligands. J Pharmacol Exp Ther. 1999; 288: 358–370.
23. Gimbrone MA. Culture of vascular endothelium. Prog Hemost Thromb. 1976; 3: 1–28.[Medline] [Order article via Infotrieve]
24. Essler M, Amano M, Kruse HJ, Kaibuchi K, Weber PC, Aepfelbacher M. Thrombin inactivates myosin light chain phosphatase via Rho and its target Rho kinase in human endothelial cells. J Biol Chem. 1998; 273: 21867–21874.
25. Murray AG, Khodadoust MM, Pober JS, Bothwell AL. Porcine aortic endothelial cells activate human T cells: direct presentation of MHC antigens and costimulation by ligands for human CD2 and CD28. Immunity. 1994; 1: 57–63.[CrossRef][Medline] [Order article via Infotrieve]
26. Ren XD, Kiosses WB, Schwartz MA. Regulation of the small GTP-binding protein Rho by cell adhesion and the cytoskeleton. EMBO J. 1999; 18: 578–85.[CrossRef][Medline] [Order article via Infotrieve]
27. Copper JA. Effects of cytochalasin and phalloidin on actin. J Cell Biol. 1987; 105: 1473–1478.
28. Bubb MR, Senderowicz AM, Sausville EA, Duncan KL, Korn ED. Jasplakinolide, a cytotoxic natural product, induces actin polymerization and competitively inhibits the binding of phalloidin to F-actin. J Biol Chem. 1994; 269: 14869–14871.
29. Birch KA, Pober JS, Zavoico GB, Means AR, Ewenstein BM. Calcium/calmodulin transduces thrombin-stimulated secretion. J Cell Biol. 1992; 118: 1501–1510.
30. Draijer R, Atsma DE, Van der Laarse A, van Hinsbergh VW. cGMP and nitric oxide modulate thrombin-induced endothelial permeability. Circ Res. 1995; 76: 199–208.
31. Patterson RL, van Rossum DB, Gill DL. Store-operated Ca2+ entry: evidence for a secretion-like coupling model. Cell. 1999; 98: 487–499.[CrossRef][Medline] [Order article via Infotrieve]
32. Wojciak-Stothard B, Potempa S, Eichholtz T, Ridley A. Rho and Rac but not Cdc42 regulate endothelial cell permeability. J Cell Sci. 2001; 114: 1343–1355.[Abstract]
33. Norman JC, Price LS, Ridley AJ, Koffer A. The small GTP-binding proteins, Rac and Rho, regulate cytoskeletal organization and exocytosis in mast cells by parallel pathways. Mol Biol Cell. 1996; 7: 1429–1442.[Abstract]
34. Kowluru A, Li G, Rabaglia ME, Segu VB, Hofmann F, Aktories K, Metz SA. Evidence for differential roles of the Rho subfamily of GTP-binding proteins in glucose- and calcium-induced insulin secretion from pancreatic beta cells. Biochem Pharmacol. 1997; 54: 1097–1108.[CrossRef][Medline] [Order article via Infotrieve]
35. Vergnolle N, Macnaughton WK, Al-Ani B, Saifeddine M, Wallace JL, Hollenberg MD. Proteinase-activated receptor 2 (PAR2)-activating peptides: identification of a receptor distinct from PAR2 that regulates intestinal transport. Proc Natl Acad Sci U S A. 1998; 95: 7766–7771.
36. Nakata T, Hirokawa N. Organization of cortical cytoskeleton of cultured chromaffin cells and involvement in secretion as revealed by quick-freeze, deep-etching, and double-label immunoelectron microscopy. J Neurosci. 1992; 12: 2186–2197.[Abstract]
37. Muallem S, Kwiatkowska K, Xu X, Yin HL. Actin filament disassembly is a sufficient final trigger for exocytosis in nonexcitable cells. J Cell Biol. 1995; 128: 589–598.
38. Sullivan R, Price LS, Koffer A. Rho controls cortical F-actin disassembly in addition to, but independently of, secretion in mast cells. J Biol Chem. 1999; 274: 38140–38146.
39. Trifaro J, Rose SD, Lejen T, Elzagallaai A. Two pathways control chromaffin cell cortical F-actin dynamics during exocytosis. Biochimie. 2000; 82: 339–352.[Medline] [Order article via Infotrieve]
40. Hall A. Rho GTPases and the actin cytoskeleton. Cell. 1998; 279: 509–514.
41. Prepens U, Just I, von Eichel-Streiber C, Aktories K. Inhibition of Fc epsilon-RI-mediated activation of rat basophilic leukemia cells by Clostridium difficile toxin B (monoglucosyltransferase). J Biol Chem. 1996; 271: 7324–7329.
42. Djouder N, Prepens U, Aktories K, Cavalie A. Inhibition of calcium release-activated calcium current by Rac/Cdc42-inactivating clostridial cytotoxins in RBL cells. J Biol Chem. 2000; 275: 18732–18738.
43. Vischer UM, Barth H, Wollheim CB. Regulated von Willebrand factor secretion is associated with agonist-specific patterns of cytoskeletal remodeling in cultured endothelial cells. Arterioscler Thromb Vasc Biol. 2000; 20: 883–891.
This article has been cited by other articles:
 |
|
 |
|
  J. Disse, N. Vitale, M.-F. Bader, and V. Gerke Phospholipase D1 is specifically required for regulated secretion of von Willebrand factor from endothelial cells Blood, January 22, 2009; 113(4): 973 - 980. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. Gavard and J. S. Gutkind Protein Kinase C-related Kinase and ROCK Are Required for Thrombin-induced Endothelial Cell Permeability Downstream from G{alpha}12/13 and G{alpha}11/q J. Biol. Chem., October 31, 2008; 283(44): 29888 - 29896. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. Agarwal, L. Covic, L. M. Sevigny, N. C. Kaneider, K. Lazarides, G. Azabdaftari, S. Sharifi, and A. Kuliopulos Targeting a metalloprotease-PAR1 signaling system with cell-penetrating pepducins inhibits angiogenesis, ascites, and progression of ovarian cancer Mol. Cancer Ther., September 1, 2008; 7(9): 2746 - 2757. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 E. Ispanovic, D. Serio, and T. L. Haas Cdc42 and RhoA have opposing roles in regulating membrane type 1-matrix metalloproteinase localization and matrix metalloproteinase-2 activation Am J Physiol Cell Physiol, September 1, 2008; 295(3): C600 - C610. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 H. Uusitalo-Jarvinen, T. Kurokawa, B. M. Mueller, P. Andrade-Gordon, M. Friedlander, and W. Ruf Role of Protease Activated Receptor 1 and 2 Signaling in Hypoxia-Induced Angiogenesis Arterioscler. Thromb. Vasc. Biol., June 1, 2007; 27(6): 1456 - 1462. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 G. Su, M. Hodnett, N. Wu, A. Atakilit, C. Kosinski, M. Godzich, X. Z. Huang, J. K. Kim, J. A. Frank, M. A. Matthay, et al. Integrin {alpha}vbeta5 Regulates Lung Vascular Permeability and Pulmonary Endothelial Barrier Function Am. J. Respir. Cell Mol. Biol., March 1, 2007; 36(3): 377 - 386. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. H. Cleator, W. Q. Zhu, D. E. Vaughan, and H. E. Hamm Differential regulation of endothelial exocytosis of P-selectin and von Willebrand factor by protease-activated receptors and cAMP Blood, April 1, 2006; 107(7): 2736 - 2744. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. Suzuki, Z.-G. Jin, D. F. Meoli, T. Matoba, and B. C. Berk Cyclophilin A Is Secreted by a Vesicular Pathway in Vascular Smooth Muscle Cells Circ. Res., March 31, 2006; 98(6): 811 - 817. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. Mehta and A. B. Malik Signaling Mechanisms Regulating Endothelial Permeability Physiol Rev, January 1, 2006; 86(1): 279 - 367. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
E. A. Lidington, R. Steinberg, A. R. Kinderlerer, R. C. Landis, M. Ohba, A. Samarel, D. O. Haskard, and J. C. Mason A role for proteinase-activated receptor 2 and PKC-{epsilon} in thrombin-mediated induction of decay-accelerating factor on human endothelial cells Am J Physiol Cell Physiol, December 1, 2005; 289(6): C1437 - C1447. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. A. Khan, C. Bianchi, E. Araujo, P. Voisine, S.-H. Xu, J. Feng, J. Li, and F. W. Sellke Aprotinin Preserves Cellular Junctions and Reduces Myocardial Edema After Regional Ischemia and Cardioplegic Arrest Circulation, August 30, 2005; 112(9_suppl): I-196 - I-201. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 X. Su, E. Camerer, J. R. Hamilton, S. R. Coughlin, and M. A. Matthay Protease-Activated Receptor-2 Activation Induces Acute Lung Inflammation by Neuropeptide-Dependent Mechanisms J. Immunol., August 15, 2005; 175(4): 2598 - 2605. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. N. McLaughlin, L. Shen, M. Holinstat, J. D. Brooks, E. DiBenedetto, and H. E. Hamm Functional Selectivity of G Protein Signaling by Agonist Peptides and Thrombin for the Protease-activated Receptor-1 J. Biol. Chem., July 1, 2005; 280(26): 25048 - 25059. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 N Vergnolle CLINICAL RELEVANCE OF PROTEINASE ACTIVATED RECEPTORS (PARS) IN THE GUT Gut, June 1, 2005; 54(6): 867 - 874. [Full Text] [PDF]
|
|
|
|
 |
|
 J.-P. Fortin, G. E. Rivard, A. Adam, and F. Marceau Studies on rabbit natural and recombinant tissue factors: intracellular retention and regulation of surface expression in cultured cells Am J Physiol Heart Circ Physiol, May 1, 2005; 288(5): H2192 - H2202. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. Feistritzer and M. Riewald Endothelial barrier protection by activated protein C through PAR1-dependent sphingosine 1-phosphate receptor-1 crossactivation Blood, April 15, 2005; 105(8): 3178 - 3184. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Steinhoff, J. Buddenkotte, V. Shpacovitch, A. Rattenholl, C. Moormann, N. Vergnolle, T. A. Luger, and M. D. Hollenberg Proteinase-Activated Receptors: Transducers of Proteinase-Mediated Signaling in Inflammation and Immune Response Endocr. Rev., February 1, 2005; 26(1): 1 - 43. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. Bhatia, K. Matsushita, M. Yamakuchi, C. N. Morrell, W. Cao, and C. J. Lowenstein Ceramide Triggers Weibel-Palade Body Exocytosis Circ. Res., August 6, 2004; 95(3): 319 - 324. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 X. Shi, B. Gangadharan, L. F. Brass, W. Ruf, and B. M. Mueller Protease-Activated Receptors (PAR1 and PAR2) Contribute to Tumor Cell Motility and Metastasis Mol. Cancer Res., July 1, 2004; 2(7): 395 - 402. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Ahamed and W. Ruf Protease-activated Receptor 2-dependent Phosphorylation of the Tissue Factor Cytoplasmic Domain J. Biol. Chem., May 28, 2004; 279(22): 23038 - 23044. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
G. M. Hjortoe, L. C. Petersen, T. Albrektsen, B. B. Sorensen, P. L. Norby, S. K. Mandal, U. R. Pendurthi, and L. V. M. Rao Tissue factor-factor VIIa-specific up-regulation of IL-8 expression in MDA-MB-231 cells is mediated by PAR-2 and results in increased cell migration Blood, April 15, 2004; 103(8): 3029 - 3037. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
B. H. Rauch, E. Millette, R. D. Kenagy, G. Daum, and A. W. Clowes Thrombin- and Factor Xa-Induced DNA Synthesis Is Mediated by Transactivation of Fibroblast Growth Factor Receptor-1 in Human Vascular Smooth Muscle Cells Circ. Res., February 20, 2004; 94(3): 340 - 345. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Minami, A. Sugiyama, S.-Q. Wu, R. Abid, T. Kodama, and W. C. Aird Thrombin and Phenotypic Modulation of the Endothelium Arterioscler. Thromb. Vasc. Biol., January 1, 2004; 24(1): 41 - 53. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
P. E. Szmitko, C.-H. Wang, R. D. Weisel, G. A. Jeffries, T. J. Anderson, and S. Verma Biomarkers of Vascular Disease Linking Inflammation to Endothelial Activation: Part II Circulation, October 28, 2003; 108(17): 2041 - 2048. [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||