FXa-Induced Responses in Vascular Wall Cells are PAR-Mediated and Inhibited by ZK-807834

Thrombosis Research 103 (2001) 281 – 297

REGULAR ARTICLE

FXa-Induced Responses in Vascular Wall Cells are PAR-Mediated and Inhibited by ZK-807834 Kirk McLean, Sabine Schirm, Anthony Johns, John Morser and David R. Light Berlex Biosciences, Richmond, CA 94804, USA (Received 5 December 2000 by Editor J. S. Bennett; revised/accepted 8 June 2001)

Abstract During thrombosis, vascular wall cells are exposed to clotting factors, including the procoagulant proteases thrombin and factor Xa (FXa), both known to induce cell signaling. FXa shows dose-dependent induction of intracellular Ca2 + transients in vascular wall cells that is active-site-dependent, Gla-domain-independent, and enhanced by FXa assembly into the prothrombinase complex. FXa signaling is independent of prothrombin activation as shown by the lack of inhibition by argatroban, hirudin and the sulfated C-terminal peptide of hirudin (Hir54 – 65(SO3 )). This peptide binds to both proexosite I in prothrombin and exosite I in thrombin. In contrast, signaling is completely blocked by the FXa inhibitor ZK-807834 (CI1031). No inhibition is observed by peptides which block interaction of FXa with effector cell protease 1 receptor (EPR-1), indicating that this receptor does not mediate signaling in the cells assayed. Receptor desensitization studies with thrombin or peptide agonists (PAR-1 or PAR-2) and experiments with PAR-1-blocking antibodies indicate that signaling by FXa is mediated by both PAR-1 and PAR-2. Potential pathophysiological responses to FXa include increased cell proliferation, increased production of the proinflammatory cytokine IL-6 and increased production of prothrombotic tissue factor. These cellular Corresponding author: Kirk McLean, Berlex Biosciences, Richmond, CA 94804, USA. Tel: +1 (510) 669 4135; Fax: +1 (510) 669 4246; E-mail: .

responses, which may complicate vascular disease, are inhibited by ZK-807834. D 2001 Elsevier Science Ltd. All rights reserved.

T

he serine protease factor Xa (FXa) is the convergence point of the extrinsic and intrinsic arms of the coagulation cascade. Factor X circulates as an inactive zymogen until activated by proteolytic cleavage between Arg52 and Ile53. Activation of FX to FXa occurs through multiple pathways. The FVIIa–tissue factor complex (extrinsic pathway) and the FIXa – FVIIIa complex (intrinsic pathway) are the principal modes of activation in blood coagulation. In addition, FX bound to CD11b/CD18 (MAC-1) on monocytes can be activated by cathepsin G [1]. During haemostasis, FXa associates with FVa and negatively charged phospholipids on platelets to form the prothrombinase complex and activate prothrombin to thrombin — the enzyme that converts fibrinogen to fibrin. In addition to their essential roles in the coagulation cascade, both FXa and thrombin function as signaling molecules. FXa in the prothrombinase complex is a major source of prothrombotic activity in thrombus [2,3]. The presence of persistent mural thrombus in vascular disease [4,5] provides a source for chronic exposure of cells in vascular tissue to FXa. We then analyzed the structural requirement for this activity and probed the mechanism (s) of induction. Cellular responses with a potential to effect pathophysiological outcome are described. Signaling by thrombin leads to well-characterized responses, including platelet aggrega-

0049-3848/01/$ – see front matter D 2001 Elsevier Science Ltd. All rights reserved. PII S0049-3848(01)00330-9

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tion, proinflammatory cytokine release from smooth muscle cells [6], endothelium-dependent relaxation of human coronary arteries [7], smooth muscle cell proliferation [8], smooth muscle cell procollagen synthesis [9], induction of Ca2 + transients in MDCK cells [10] and induction of apoptosis in motor neurons [11]. These responses have been attributed to activation of the thrombin receptor, also referred to as protease-activated receptor-1 (PAR-1). Thrombin cleavage of PAR-1 between Arg41 and Ser42 releases an N-terminal peptide, generating a new N-terminus that is the tethered ligand SFLLRN. The tethered ligand interacts with residues in the second extracellular loop of PAR-1, leading to activation. Synthetic peptide analogs of the tethered ligand are able to activate PAR-1 independent of receptor cleavage. PAR-1 expression has been detected on platelets [12], endothelial cells [12,13], smooth muscle cells [9], gingival fibroblasts [14], monocytes [15], T cells [16], neurons [17] and various tumor cells [18]. PAR-1 is one member of a family of receptors known as the protease-activated receptors (PARs), which to date includes PAR1, PAR2, PAR3 and PAR4. Thrombin can activate PAR1, PAR3 and PAR4 [19,20]. PAR2 can be activated by trypsin and is reported to mediate activation of coronary artery smooth muscle cells [21]. Functional interactions are reported for the PAR receptors on cell surfaces. Examples include the transactivation of PAR2 by PAR1 on human endothelial cells [22] and the cofactor activity of PAR3 for activation of PAR4 on mouse platelets [23]. Several of the responses elicited by FXa are reminiscent of those attributed to thrombin, but the mechanism of FXa signaling is less well understood. FXa induces Ca2 + responses, adhesion molecule expression (ICAM-1, VCAM-1) and production of proinflammatory cytokines (MCP-1, IL-6, IL-8) in endothelial cells [24,25]. FXa also acts as a potent mitogen for smooth muscle cells [26], stimulates lymphocyte proliferation [27] and is implicated in VEGF production by fibroblasts [28]. The receptor system involved in these responses is controversial. PAR2 activation has been shown both by FXa, produced in situ by the factor VIIa–tissue factor complex and by the factor VIIa – tissue factor

complex itself [29]. However, it is suggested that some effects of FXa are mediated by FXa binding to effector cell protease 1 receptor (EPR-1) [25,27]. EPR-1 is a 337-amino acid protein containing an 81-amino acid serine-rich cytoplasmic domain, a single transmembrane domain and a 230-amino acid extracellular domain. The FXa sequence from Leu83 to Leu88 has been shown to interact with the EPR-1 and inhibit [30] binding of FXa to EPR-1 and subsequent Ca2 + signals [31] in human umbilical vein endothelial cells. Recently, a survey of EST databases failed to detect expression of this gene and polymerase chain reaction and Southern blot hybridization analyses using EPR-1-specific probes failed to detect EPR-1 in either the mouse or human genomes [32]. In order to determine the potential effects of FXa exposure to vascular tissue, we determined the ability of FXa to induce Ca2 + transients in the principle types of vascular wall cells.

1. Materials and Methods 1.1. Reagents FXa was from Enzyme Research Laboratories, South Bond, IN or Haematological Technologies, Essex Junction, Vermont. FXa-beta, des-Gla FXa, FXa–EGR and a-thrombin were from Haematological Technologies, Essex Junction, Vermont. Recombinant hirudin was from American Diagnostica, Greenwich, CT. Argatroban was a gift from Mitsubishi (Japan). Tick anticoagulant peptide or TAP, peptides and ZK-807834 [33] were made at Berlex Biosciences (Richmond, CA). Cell culture media was from Clonetics, San Diego, CA. HEPES buffer and Hank’s salts were from Gibco, Rockville, MD BRL. BSA was from Sigma, St. Louis, MO. ATAP2 and WEDE15 antibodies [34] were a generous gift from Dr. Lawrence Brass. 1.2. Cell Culture Human coronary artery smooth muscle cells (HCASMCs), human aortic smooth muscle cells (HASMCs), human adventitial fibroblasts (HAOAFs) and human aortic endothelial cells (HAECs) were purchased at passage 3 from Clonetics, San Diego, CA and used from passage

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4 through 8. HCASMC and HASMC were carried in SMGM. HAOAF and HAEC were carried in FGM and EGM-2 media, respectively.

factor were assayed by ELISA 24 h after stimulation of the cells (R&D Systems, Minneapolis, MN and American Diagnostica, respectively).

1.3. Measurement of Ca2+ Transients

1.7. Thymidine Incorporation

Cells were plated in black-bottom, 96-well tissue culture plates (Costar, Acton, MD) at a density of 5000 cells/well and allowed to grow until confluent. For SMC, the medium was changed to serum-free SMBM the night before the experiment. The cells were loaded with the Ca2 + -sensitive dye Fluo-3 (Molecular Probes, Eugene, OR) for 60–90 min at 37C. For HAEC, HCASMC and HASMC, 42 ml of a 50:50 mix of Fluo-3 (2 mM, DMSO)/20% pluronic acid was diluted into 1  Hank’s salts, 20 mM HEPES, 0.1% BSA (HHB). For HAOAF, fibroblast basal medium was used instead of Hank’s salts. After loading the dye, the cells were washed four times with HHB in a Denley plate washer and exposed to inhibitors and agonists in HHB. The Ca2 + transients were measured in a fluorometric Imaging Plate Reader (FLIPR; Molecular Devices Corporation, Sunnyvale, CA). Readings were taken every second for the first 60 s then every 3 s for the next 2 min.

For proliferation assays, cells were growtharrested for 2 days in serum-free medium in the absence of growth factors and then incubated with 6-3H-thymidine (5 Ci/mmol; AmershamPharmacia, Piscataway, NJ) at a final concentration of 2 mM thymidine (Sigma, St. Louis, MO) for another 48 h. The cells were washed with PBS and lysed in 0.5 M NaOH. Thymidine incorporation was determined by liquid scintillation counting in EcoLume (ICN, Costa Mesa, CA).

1.4. Desensitization Studies For desensitization studies, cells were exposed to FXa (347 nM), a-thrombin (1 nM) or PAR peptides (100 mM) during the loading period. For treatment with peptides, 100 mM peptide was added a second time after the washing step. 1.5. PAR-1 Antibody Inhibition The PAR-1 antibodies ATAP2 and WEDE15 (20 mg/ml each) were added to the cells during the loading period. The cells were washed four times with HHB and then cells were exposed to antibody again 5 min prior to addition of agonist.

2. Results 2.1. FXa Induces Ca2+ Transients in Vascular Wall Cells The ability of human FXa to act as a signaling molecule was assessed in human aortic adventitial fibroblasts (HAOAFs), HASMCs, HAECs and HCASMCs using a fluorescence-based analysis of intracellular Ca2 + transients. The Ca2 + measurements were made using the FLIPR instrument from Molecular Devices Corporation, Sunnyvale, CA that measures Ca2 + transients on attached cells. FXa induced Ca2 + transients dose-dependently in all four cell types (Fig. 1). We observed signaling at concentrations as low as 22 nM FXa. In all cell lines tested, FXa was a weaker agonist than a-thrombin and twoorders-of-magnitude-greater levels of FXa were required to induce an equivalent response. In HCASMC and HAECs, we found that Ca2 + transients induced by FXa were slightly greater than those induced by equal molar concentrations of g-thrombin (data not shown).

1.6. IL-6 and Tissue Factor Induction

2.2. FXa Structural Requirements for Induction of Ca2+ Transients

Cells were plated at a density of 25,000 cells/well in 24-well plates coated with 2% gelatin (Sigma, St. Louis, MO) and allowed to grow to confluency. The release of IL-6 and the expression of tissue

We determined the active site and Gla domain dependence for the FXa-induced Ca2 + transients. The FXa-induced response was active-sitedependent as shown by an inability of FXa–EGR

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Fig. 1. Ca2 + transients induced by FXa. (A) HAOAF, (B) HCASMC, (C) HASMC, (D) HAEC. Data are expressed as maximum peak height. Representative experiment shown for each.

(active-site-inhibited FXa) to induce a Ca2 + response in HAOAF or HCASMC. In agreement with this observation, the FXa active site inhibitor TAP (1 mM) blocked 90 ± 2% (average ± S.D., n = 4) of the Ca2 + transients induced in HAOAF by FXa (173 nM). The Gla domain plays an important role in Ca2 + -dependent association of FXa with phospholipids. We purchased Gla domainless FXa (des-Gla FXa) from Haematological Technologies, Essex Junction, Vermont. N-terminal sequencing verified that the product had the Gla domain removed. Equal molar amounts of des-Gla FXa induced Ca2 + transients in HAOAF (90 ± 19% of control FXa, n = 4) and HCASMC (78 ± 15% of control FXa, n = 4), indicating that the Gla domain of FXa is not required for signaling. In both HAOAF (data not shown)

and HASMC (Fig. 2), we found that EGTA completely inhibited FXa-induced Ca2 + transients, but had no effect on thrombin-induced Ca2 + transients. FXaa cleaves itself at Arg290, releasing a 19amino acid C-terminal peptide to generate FXab. We compared signaling by FXaa and FXab in HASMC. The ED50 values (concentration of FXa necessary to reach 50% of the maximum signal) were found to be 114 and 157 nM, respectively, a statistically insignificant difference. The response of HASMC to FXa from different sources (Enzyme Research Laboratories, South Bond, IN and Haematological Technologies, Essex Junction, Vermont) was identical with ED50 values of 121 and 114 nM, respectively. Neither the source of FXa nor deletions at the

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Fig. 2. EGTA inhibits FXa, but not thrombin-induced Ca2 + transients in HASMC. Concentrations of 347 nM FXa (square) and 500 pM thrombin (triangle) were used as agonists. Increasing amounts of EGTA were premixed with the agonists prior to addition. A representative experiment is shown. Data are expressed as percent uninhibited control. Average ± S.D. (n = 2).

C-terminus affected the signaling ability of FXa with these cells. 2.3. FXa-Induced Ca2+ Transients are Independent of Thrombin Generation In the presence of Ca2 + and phospholipids, FXa converts prothrombin to thrombin (factor Va accelerates this reaction) [35]. Any prothrombin present in these experiments could be converted to thrombin that would activate cells. All experiments were carried out in serum-free medium to eliminate this source of prothrombin. To verify that signaling was not due to residual prothrombin being activated to thrombin, inhibition by the thrombin inhibitors, hirudin and argatroban, was compared to inhibition by ZK807834, a potent (0.11 nM), selective, FXa inhibitor [33]. Concentrations of hirudin and argatroban that completely blocked thrombin-induced Ca2 + transients in HAOAF (Fig. 3A), HASMC (data not shown) and HAEC (only hirudin tested; data not shown) did not significantly affect FXa and bradykinin-induced Ca2 + transients. ZK-807834 completely abolished signaling caused by FXa, whereas the effect on bradyki-

nin-induced signaling was insignificant. A modest 20% reduction in thrombin-dependent signaling by 1 mM ZK-807834 was expected due to the weak inhibition of thrombin by ZK807834 (Ki = 2 mM). In order to eliminate the possibility that activation of residual prothrombin on the cell surface could result in formation of a form of thrombin protected from inhibition by the active site inhibitors hirudin and argatroban, additional experiments were performed with the sulfated C-terminal peptide of hirudin (Hir54 – 65(SO3 )). Hir54 – 65(SO3 ) is a potent binder of both the exosite I in thrombin and the proexosite I in prothrombin and thus, treatment with this peptide will inhibit any prothrombin present prior to activation. Saturating concentrations (100 mM) of Hir54 – 65(SO3 ) that completely inhibited thrombin signaling in HAOAF did not effect FXa signaling (Fig. 3B). These data verify that FXa signaling is independent of thrombin generation. Dose–response curves were generated (n = 5) and IC50 values determined for ZK-807834 with 347 nM FXa, in HAOAF (IC50, 163 ± 10 nM), HAEC (IC 50 , 179 ± 11 nM), and HCASMCs (IC50, 174 ± 7.4 nM). The concentration of FXa

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Fig. 3. Neither prothrombin nor thrombin inhibitors inhibit FXa-induced Ca2 + transients in HAOAF. (A) FXa (347 nM), thrombin (1 nM) and bradykinin (100 nM) were used as agonist. The agonists were premixed with buffer as control (open), 48 U/ml hirudin (striped), 50 nM argatroban (solid) and 1000 nM ZK-807834 (stippled) prior to addition. Results are expressed as percent uninhibited control. Data are average ± S.D. (n = 4). (B) Hir54 – 65(SO3 ), a peptide capable of binding to prothrombin prior to activation, effectively blocks signaling induced by thrombin but not by FXa. FXa (175 nM) and thrombin (1 nM) were used as agonist. The agonists were premixed with buffer as control (open), 100 mM Hir54 – 65(SO3 ) (solid) or 500 nM ZK-807834 (stippled) prior to addition. Results are expressed as percent uninhibited control. Data are average ± S.D. (n = 10).

used was three orders of magnitude above the Ki of this inhibitor for FXa. The IC50 values generated from these data are consistent with the stoichiometric titration of the agonist FXa used in these experiments. Sufficient concentrations of the active site inhibitor, ZK-807834, completely inhibit FXa-induced Ca2 + transients in these cells.

2.4. Thrombin or FXa Pretreatment Attenuates Thrombin or FXa-Induced Ca2+ Transients Proteolytically active FXa is necessary for signaling, therefore, FXa may interact with a member of the PAR family of receptors to trigger cellular responses. Pretreatment of cells containing the thrombin receptor PAR-1 with thrombin leads to

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activation and subsequent desensitization of this PAR and other thrombin receptors. Pretreated, desensitized cells will not respond to a second exposure to thrombin or other agonists that utilize the same receptor. Pretreatment of HASMC cells with either thrombin (1 nM) or FXa (347 nM) during the loading period did desensitize the cells and inhibited subsequent activation by FXa by 44 ± 3% and 64 ± 2%, respectively (Fig. 4A). In the same experiment, thrombin pretreatment inhibited subsequent activation by 1 nM thrombin by 93 ± 3%. FXa pretreatment partially desensitized the cells to subsequent treatment by thrombin (34 ± 1% inhibition). Similar results were obtained with HAEC (Fig. 4B). For HAOAF cells, thrombin desensitization led to a 90 ± 3%

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inhibition of FXa-induced Ca2 + transients. The greater inhibition seen with HAOAF is consistent with a greater proportion of the signal proceeding through PAR-1 in these cells (see PAR-1 antibody results below). Cross-desensitization indicated that part of the response to FXa and thrombin was through the same receptor, and this receptor was likely to be PAR-1. 2.5. PAR-1 and PAR-2 Activation Peptides Induce Ca2+ Transients and Desensitize Cells Pretreatment with thrombin desensitized cells to FXa, indicating that the PAR-1 receptor may mediate FXa signaling. Only partial desensitization resulted from thrombin pretreatment, there-

Fig. 4. Thrombin or FXa pretreatment desensitizes cells to subsequent FXa activation. Buffer (open), 347 nM FXa (striped) or 1 nM thrombin (solid) were present during the 1-h loading period. Cells were washed and then exposed to FXa (347 nM) or thrombin (1 nM). Both HASMC (Panel A) and HAEC (Panel B) were tested. Data are expressed as percent nonpretreated control. Data are average ± S.D. (n = 2).

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fore, the contribution of PAR-2 (not cleaved by thrombin) to FXa signaling was determined. Agonist peptides have been developed corresponding to the tethered ligands of the thrombin-sensitive receptor PAR-1 and the thrombininsensitive receptor PAR-2 [36]. The PAR-1 activation peptide (PAR-1 AP, SFLLRN) and PAR-2 activation peptide (PAR-2 AP, SLIGKV) at 100 mM caused Ca2 + transients in HAOAF,

HCASMC and HASMC (Fig. 5A – C). The responses induced by the PAR-1 AP were equal to or slightly greater than the responses elicited by 1 nM a-thrombin. These experiments verified that for these cells, activation of PAR-1 and PAR-2 induces Ca2 + transients. As observed for other G-protein-coupled receptors, activation of the PAR receptors is followed by a period of desensitization [37]. Before treatment with FXa, thrombin or bradykinin, vascular cells were desensitized by pretreatment with PAR-1 or PAR-2 activation peptides (100 mM). Upon returning to baseline, cells were challenged with FXa, thrombin or bradykinin. Prior desensitization with the PAR-1 peptide led to a > 90% inhibition of FXa and thrombininduced Ca2 + transients in HAOAF, HCASMC, HASMC and HAEC (Table 1). In HAOAF cells, PAR-2 peptide desensitization inhibited FXainduced Ca 2 + transients by approximately 38 ± 11%, while having little effect on the thrombin- (8 ± 18%, n = 12) and bradykinin- (11 ± 18%, n = 12) induced Ca2 + transients. In HCASMC (and HUVEC, data not shown), desensitization with the PAR-2 AP led to a larger nonspecific downregulation of the Ca2 + response as measured by the inhibition of thrombin- (29 ± 14%, n = 12) and bradykinin- (35 ± 18%, n = 12) induced Ca2 + transients. In these cells, FXa-induced Ca2 + transients were inhibited to a greater degree, following PAR-2 AP desensitization (85 ± 18%, n = 12), consistent with a PAR-2-mediated signaling component for FXa. 2.6. PAR-1 Antibodies Inhibit Ca2+ Transients Induced by FXa or Thrombin

Fig. 5. PAR peptides stimulate Ca 2 + transients. HAOAF (Panel A), HCASMC (Panel B) and HASMC (Panel C) were exposed to 100 mM of either SFLLRN (PAR-1 AP) or SLIGKV (PAR-2 AP). Data are expressed are maximum peak height. Representative experiments shown for each. Data are average ± S.D. (n = 4).

The PAR-1 activation peptide activates both PAR-1 and PAR-2 receptors [38] and the PAR2 AP has some nonspecific negative effect on the induction of Ca2 + transients in the SMC and HAEC. To specifically address the issue of the involvement of PAR-1 or PAR-2 in cell activation by FXa PAR-1-directed monoclonals, ATAP2 and WEDE15 (20 mg/ml each), which inhibit activation of PAR1 by thrombin [34], were used. For HAOAF, HCASMC, HUVEC and HAEC, the PAR-1 antibodies completely inhibited Ca2 + transients induced by 1 nM athrombin, while a control monoclonal antibody

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Table 1. Attenuation of FXa-induced Ca2 + transients following PAR peptide desensitization and PAR-1 antibody pretreatment Cell line

PAR-1 AP

PAR-2 AP

HAOAF HCASMC HAEC HUVEC

90 ± 9 92 ± 2 95 ± 2 n.d.

38 ± 11 85 ± 18 64 ± 3 85 ± 5

PAR-1 antibodies 68 ± 20 35 ± 22 53 ± 7 34 ± 10

PAR-1 antibodies/PAR-2 AP 97 ± 8 100 ± 3 100 ± 3 99 ± 1

The PAR-1 antibodies ATAP2/WEDE15 were used at 20 mg/ml each to inhibit PAR-1-mediated Ca2 + transients. Agonist peptides for PAR-1 and PAR-2 were used at 100 mM. Both antibodies or agonist peptides were present during Flou-3 loading and again added following the wash step and just prior to addition of agonists. Data are expressed as percent inhibition versus nonpretreated control. Data are average ± S.D. (n = 4 for HAEC and HUVEC; n = 12 for HAOAF and HCASMC).

was without effect. The extent to which the antibodies-inhibited activation by FXa (347 nM) varied with the cell line being tested (Table 1). For HAOAF cells, inhibition was 68 ± 20%, while for HCASMC, inhibition was 35 ± 22% — a statistically significant difference ( P < .001, n = 12). The ability of PAR-1 antibodies to inhibit FXa-induced Ca2 + transients in HUVEC and HAEC was also determined. The average inhibition was 35 ± 10% and 53 ± 7%, respectively (n = 4). Following desensitization of these cells with the PAR-2 activation peptide, coupled with the addition of the PAR-1 antibodies prior to addition of FXa, it was possible to completely inhibit (>95%) FXa-induced Ca2 + transients in HAOAF, HCASMC, HAEC, and HUVEC. 2.7. EPR-1 Peptides Do Not Affect FXa-Induced Ca2+ Transients The effector cell protease receptor-1, EPR-1, has been proposed to act as a receptor for FXa on leukocytes, platelets, smooth muscle cells and endothelial cells [26,39 – 41]. The peptide LFTRKL(G) is based on the FXa sequence from Leu83 to Leu88 and has been reported to inhibit FXa binding to EPR-1 in HUVEC and HASMC [26,41]. The peptide SPGKPGNQNSKNEPP corresponds to the sequence on EPR-1 that binds to FXa. This peptide contains a region of homology to a sequence on factor Va (NQNSKN on EPR-1, NQNSSN on FVa) and also inhibits binding of FXa to EPR-1 [42]. These peptides were tested at concentrations up to 500 mM to assess the role of EPR-1 in mediating FXa-induced Ca2 + transients in HAOAF and HASMC. Neither peptide affected FXa-induced Ca2 + transients, consistent with the reported ineffectiveness of an EPR-1

peptide in FXa-induced cytokine production in HUVEC cells [24]. 2.8. Effects of Prothrombinase Assembly Assembly of FXa into the prothrombinase complex accelerates its rate of prothrombin cleavage 500,000-fold [43]. To determine if FXa-induced Ca2 + transients were increased in the presence of the prothrombinase complex components, factor Va and phospholipids were added to FXa in the absence of prothrombin. The response of HAOAF to FXa (13 nM) was enhanced fivefold when FXa was assembled in the prothrombinase complex. The increased signal was not due to thrombin generation because hirudin, added at a concentration that completely inhibited signaling by 1 nM thrombin, had no effect on the prothrombinase signal. In contrast, 1 mM ZK-807834 completely inhibited signaling. Neither factor Va nor phospholipids, alone or in combination, had any effect on Ca2 + transients in the absence of FXa. 2.9. FXa Induces the Release of Proinflammatory Factors FXa induces proinflammatory responses [24,25]. Because FXa induced Ca2 + transients in the vascular cells and enhanced production of the proinflammatory cytokine IL-6 in HUVEC [24], we investigated IL-6 production in HCASMC and HAOAF cells. Cells in 24-well dishes were grown to confluence, growth-arrested for 24 h and triplicates incubated with increasing concentrations of FXa under serum-free condition. As a control, parallel cell cultures were stimulated with TNF-a and IFN-g or left without a stimulus.

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FXa induced the release of IL-6 into the culture medium from both HAOAF (data not shown) and HCASMC (Fig. 6A) in a dose-dependent fashion. When 173 nM FXa was mixed with increasing concentrations of the active site inhib-

Fig. 7. ZK-807834 inhibits FXa induced proliferation of HCASMC. Cells were treated with growth arrest media (GA; no agonist) or Fxa (347 nM) in the presence of increasing concentrations of ZK-807834. Labelling with [3H]thymidine was carried out for 48 h after stimulation of cells. Incorporation of thymidine was determined by liquid scintillation counting. A representative experiment is shown. Data are average ± S.D. (n = 3).

itor ZK-807834 (Fig. 6B), the FXa-induced release of IL-6 was completely inhibited (IC50, 98 ± 25 nM). Induction of IL-6 by FXa was not affected by hirudin at 40 U/ml (Fig. 6C), which fully inhibits thrombin-induced IL-6 release in agreement with the Ca2 + transient experiments. Thus, thrombinindependent signaling by FXa leads to a biological outcome. 2.10. FXa Induces Tissue Factor Expression in HCASMC FXa has been shown to increase tissue factor expression in HUVEC cells [44]. To determine if a similar response occurs in HCASMC, we anaFig. 6. FXa induces the release of IL-6 from HCASMC. (Panel A) Increasing concentrations of FXa were added to HCASMC. After 24 h, the media was removed and assayed for IL-6 content. (Panel B) A constant amount of FXa (173 nM) was mixed with increasing concentrations of ZK-807834 before adding to the cells. After 24 h, the media was removed and assayed for IL-6 content. (Panel C) Cells were treated with FXa (173 nM) or thrombin (10 nM). These conditions were used ± 40 U/ml hirudin ( Hir). After 24 h, the media was removed and assayed for IL-6 content. For all panels, GA is growth arrest media without agonist. Units are picograms per milliliter of IL-6. A representative experiment is shown. Data are average ± S.D. (n = 3).

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lyzed cells for elevated content of tissue factor after treatment with 173 nM FXa. Similar to the observed effect of induced IL-6 release, the amount of cell-associated tissue factor increased from 583 ± 30 to 1175 ± 98 pg/ml (average ± S.D., n = 3) after exposure to FXa and the increase was inhibited by ZK-807834 in a dose-dependent manner (IC50, 120 ± 55 nM, n = 3). 2.11. FXa Induces DNA Synthesis in HCASMC Finally, we determined that FXa signaling affects DNA synthesis in HCASMC. Treatment of confluent cells in serum-free medium with 173 and 347 nM FXa increases thymidine incorporation by 2.15- and 2.29-fold, respectively. Treatment with 10 nM thrombin resulted in a similar increase (data not shown). The effect of FXa (347 nM) on DNA synthesis was inhibited by ZK-807834 (Fig. 7), but not hirudin (data not shown).

3. Discussion The potential effect of FXa exposure to vascular tissue was determined by testing the ability of FXa to induce Ca2 + transients in the principle types of vascular wall cells. A dose-dependent induction of Ca2 + transients by FXa in the primary vascular wall cell lines HAEC, HAOAF, HASMC and HCASMC was seen. Intracellular Ca2 + signals were observed at concentrations as low as 22 nM FXa. In HCASMC and HAEC, the magnitude of the FXa-induced Ca2 + response was at least as great as with equimolar concentrations of g-thrombin. In mechanistic experiments presented here and in similar studies [31], higher concentrations of FXa are utilized (100–300 nM) in order to maximize the signal. With respect to the physiological relevance of FXa signaling, similar levels could be achieved at focal sites of FX activation if all of the 140 nM FX in a localized volume of plasma are activated. In patients with disseminated intravascular coagulation, circulating levels of FX activation peptide increase from normal levels of 0.07 nM to as high as 0.55 nM [45]. Such large increases in circulating FX activation peptide may result from robust local activation of FX.

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The structural requirements for this activity were analyzed by comparing FXai, des-gla FXab, FXab, FXaa and the prothrombinase complex as Ca2 + transient agonists. Signaling by free FXa depends on the presence of an intact active site and external Ca2 + , but not the FXa Gla domain. EGTA inhibited Ca2 + transients induced by FXa, but not thrombin, indicating that FXa requires external Ca2 + to trigger internal Ca2 + transients. In addition to Ca2 + binding sites in the Gla domain, FXa contains high-affinity Ca2 + binding sites in the EGF and protease domains that affect proteolytic activity and may be required by FXa to activate receptors [46,47]. The protease activity of a truncated FXa analog lacking the Gla domain is still Ca2 + -dependent [46,47]. Reduced catalytic efficiency of FXa in the absence of Ca2 + may attenuate signaling in the presence of EGTA. Alternately, the interaction between FXa and a receptor or cofactor could be Ca2 + -dependent or FXa could trigger the influx of extracellular Ca2 + . Assembly of FXa into the prothrombinase complex resulted in Ca2 + transients at lower FXa concentrations than FXa alone. The fivefold increase in sensitivity to FXa is much less than the 500,000-fold increase in the ability of the prothrombinase complex to convert prothrombin to thrombin [43]. ZK-807834 (CI-1031), a potent and selective FXa inhibitor [33], inhibited FXa-induced Ca2 + signals. Due to the high affinity of ZK-807834 for FXa, inhibition of FXa-induced Ca2 + transients by ZK-807834 requires concentrations of inhibitor that stoichiometrically titrate the total amount of FXa used as agonist. Complete inhibition by ZK-807834 confirms that signaling and other cellular responses to FXa depend on a functional FXa active site. Thrombin inhibitors hirudin and argatroban failed to inhibit FXa signaling. During review, the possibility was suggested that activation of residual prothrombin on the cell surface could result in formation of a form of thrombin protected from inhibition by the active site inhibitors hirudin and argatroban. In order to eliminate this possibility, additional experiments were performed with the sulfated C-terminal peptide of hirudin (Hir54 – 65(SO3 )) in order to inhibit prothrombin prior to activation. Hir54 – 65(SO3 ) is a potent binder of both the exosite I in thrombin (25 nM)

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and the proexosite I in prothrombin (1.6–3 mM) and both forms will be saturated by 100 mM levels of the peptide [48,49]. In agreement with results for hirudin and argatroban, Hir54 – 65(SO3 ) fully inhibits signaling by thrombin, but not FXa. Thus, Ca2 + transients are directly induced by FXa and not dependent on thrombin generation. Experiments to probe the signaling mechanism utilized thrombin and PAR peptide desensitization, direct inhibition by PAR-1 antibodies and EPR-1 peptide competition studies. Pretreatment with thrombin desensitized cells to FXa signaling, indicating cross-reactivity between thrombin and FXa for a common receptor. Signaling by PAR peptides shows that PAR-1 and PAR-2 are expressed in human endothelial cells, smooth muscle cells and adventitial fibroblasts. Desensitization by PAR activation peptides implicates either PAR-1 or PAR-2 or both act as target receptors for FXa-induced Ca 2 + transients. PAR-1 antibodies, ATAP2 and WEDE15 verified that a component of the FXa-induced signaling occurs through PAR-1. Complete inhibition of FXa-induced Ca2 + transients resulted from the combination of PAR-2 AP desensitization and PAR-1 antibody pretreatment. These experiments point to PAR-1 and PAR-2 as the receptors involved in FXa-induced Ca2 + transients. PAR-1 activation by FXa suggests a potential role for signaling by FXa in diseases where thrombin is absent or fully inhibited. PAR-2 (not activated by thrombin) activation by FXa increases the spectrum of PAR receptors triggered by coagulation factors. PAR-2 is expressed on human neutrophils [50], neurons [51], lung fibroblasts [52] and vascular smooth muscle cells [53]. Reports of the effects of PAR-2 activation include induction of leukocyte rolling and adhesion [54], induction of inflammation in a rat hind paw mode [55], stimulation of a proinflammatory response via a neurogenic mechanism [51] and stimulation of proliferation in human lung fibroblasts [52] or vascular smooth muscle cells [53]. FXa triggered biological responses including IL-6 production, tissue factor production and proliferation in HCASMCs. These FXa-induced responses were blocked by ZK-807834, but not by the thrombin inhibitor hirudin. FXa induces proinflammatory responses in vitro and in vivo including increases in cytokine production, che-

motactic protein production, adhesion molecule expression and tissue factor expression in human endothelial cells [24,44,56]. Injection of FXa leads to edema formation in a rat foot paw model of acute inflammation [57]. PAR-1 activation has proinflammatory effects and is implicated in the release of chemotactic proteins in HUVEC [58]. The PAR-1-activating peptide induces edema in the rat paw model through release of bioactive amines from mast cells [59] and increases IL-6 gene expression and secretion in gingival fibroblasts [14]. FXa causes a dose-dependent induction of IL-6 and tissue factor expression in HCASMC and HAOAF. IL-6 is a potent inflammatory cytokine and increased levels correlate with poor prognosis in unstable angina [60–62]. Increased tissue factor expression increases the thrombogenic potential of the cell surface, leading to the generation of additional FXa and propagates thrombosis and associated inflammation induced by FXa and thrombin. During sepsis, LPS increases the expression of tissue factor and CD11b/CD18 on monocytes and macrophages. Increased expression of tissue factor and CD11b/CD18 in sepsis leads to activation of FX to FXa and triggers consumptive coagulopathy and DIC. In a baboon model of Escherichia coli-induced septic shock, DEGR-FXa inhibits the development of DIC; however, it has no effect on lethality [63]. DEGR–FXa competes with active FXa, inhibiting formation of an active prothrombinase complex and therefore generation of thrombin. Thrombin inhibitors, heparin and hirudin fail to impact lethality in this model [64]. In contrast, DEGR–FVIIa, TFPI and neutralizing tissue factor antibodies all lead to decreased coagulation as well as reduced mortality [65–67]. A potential explanation (Fig. 8) for the differential ability of selective anticoagulants to affect mortality is that DEGR–FXa inhibits thrombin generation, but not the ongoing generation of active FXa. Although FXa is effectively competed by DEGR–FXa from participation in prothrombinase assembly, and subsequent coagulation, active FXa may cleave and activate PAR receptors, leading to increased levels of proinflammatory mediators. Although thrombin inhibitors are effective inhibitors of coagulation and thrombin signaling through PARs, they cannot inhibit FXa signaling. FXa inhibitors are effective anticoagu-

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Fig. 8. Scheme depicting potential of coagulation inhibitors to prevent FXa and thrombin-induced signaling.

lants [68–70] and the present work shows they can eliminate PAR signaling by FXa. FXa inhibitors will also attenuate thrombin-dependent signaling by inhibiting the generation of thrombin (Fig. 8). Thus, FXa active site inhibitors may be superior to thrombin inhibitors in sepsis and related syndromes. Proliferation of SMC is one factor implicated in vessel narrowing during atherosclerosis and restenosis. We show that FXa induces smooth muscle cell proliferation. The induction of proliferation is not dependent on thrombin and is prevented by ZK-807834. Thrombin induces smooth muscle cell proliferation and induces procollagen synthesis through a PAR-1-mediated mechanism [9]. PAR-2 activation peptide induces proliferation in human vascular smooth muscle cells [53]. Proliferation and induction of collagen synthesis in smooth muscle cells contribute to vessel narrowing associated with atherosclerosis and restenosis following angioplasty. Restenosis occurs in approximately 30% of patients following angioplasty [71]. Coagulation inhibitors have been shown to reduce restenosis and neointimal hyperplasia in a rabbit injury model [72]. Inhibitors of early steps in the coagulation cascade had the greatest beneficial effect. Specific inhibition of FXa by TAP and recombinant antistasin has been shown to reduce restenosis after balloon angioplasty in rabbits [68]. Thus, while FXa inhibition provides acute anticoagulant benefit, long-term inhibition of FXa-dependent cell signaling may help limit restenosis. As therapeutic targets, FXa and prothrombinase are normally perceived to provide benefit in acute thrombosis and FXa inhibition is effective

against both venous and arterial thrombosis [69,70,73]. The present studies suggest additional benefits of both acute and longer-term FXa inhibition. Inhibition of tissue factor expression on vascular cells in vitro suggests that FXa inhibition may help to block the second wave of tissue factor induction measured in vascular injury models [3]. It will be important to determine the relative contribution to tissue factor generation from cells in the injured vessel wall versus that from infiltrating monocytes and whether FXa can signal in monocyte, neutrophil or platelets. In addition, the ability of FXa to induce cell proliferation suggests that a FXa inhibitor may modulate vascular remodeling following injury. Several vascular cell lines respond to FXa, resulting in PAR-1/PAR-2-mediated Ca2 + transients, prothrombotic, proinflammatory and proliferative effects that could contribute to disease. Selective FXa inhibition is an important mechanism deserving further testing in animal models of vascular disease. We thank J.-H. Lin, R. Pagila and C. Adams for help with TAP production; S. Biancalana for synthesis of peptides; Galina Rumennik for help with prothrombinase; Meina Liang for help using FLIPR; and Dr. William Dole for helpful discussions and support.

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