Stimulated platelets generate bioactive TF

Since stimulated platelets use their splicing machinery to produce mature TF mRNA, we asked if TF-dependent procoagulant activity increased in activated cells. Freshly isolated platelets, at cell numbers (2 × 10⁹ total) that approximate those found in 5–10 ml of whole blood, did not possess significant levels (P < 0.05) of procoagulant activity (Fig. 2, A and B). In contrast, procoagulant activity was markedly increased as early as 5 min after platelets adhered to fibrinogen in the presence of thrombin (Fig. 2 A) and continued to accumulate in platelets and platelet-derived microparticles in a time-dependent fashion (Fig. 2 B). Platelets activated in suspension with ADP, collagen, or thrombin also accumulated TF-dependent procoagulant activity (unpublished data). We consistently observed that activated platelets possessed higher procoagulant activity than quiescent platelets in samples from different donors, but the magnitude was variable among subjects, ranging from a 2.8- to a 15.3-fold increase over baseline (mean increase over baseline 7.7 ± 2.0). In contrast to activated platelets, monocytes (5 × 10⁶) stimulated with fibrinogen and thrombin did not generate appreciable procoagulant activity, although they did respond to lipopolysaccharide (unpublished data).

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Figure 2.

Platelet activation increases TF-dependent procoagulant activity. (A and B) A timecourse (A, 0–60 min; B, 0–4 h) of TF-dependent procoagulant activity in platelets that have adhered to fibrinogen in the presence of thrombin. In B, each line represents procoagulant activity from platelet membranes (Plts), platelet-derived microparticles (Mp), or platelet membranes together with microparticles from the same sample (Plts + Mp). The data are displayed as pM of TF per 2 × 10⁹ platelets and represent the mean ± SEM of three independent experiments. (C) Immunolocalization of TF in freshly isolated platelets (left) and platelets that have adhered and spread on immobilized fibrinogen in the presence of thrombin for 2 h (right). The green stain in both panels represents actin. The red stain in the right panel shows immunolocalization of TF on the surface of activated platelets (see arrow). This experiment is representative of two independent studies. (D and E) The bars (n = 4) in these panels show TF-dependent procoagulant activity in freshly isolated platelets or platelets activated as in B for 4 h in the presence or absence of factor VIIa (D) or a neutralizing antibody directed against TF (E). A single asterisk (*) indicates a statistically significant difference (P < 0.05) in TF-dependent procoagulant activity between freshly isolated (baseline) and activated platelets; the double asterisk (**) represents a significant difference (P < 0.05) between activated platelets under untreated or treated conditions. (F) These panels show clot formation in plasma that is incubated with membranes isolated from quiescent platelets or platelets activated with fibrinogen and thrombin for 5 min (left) or 2 h (right). The white bars represent activated platelets that were incubated with a neutralizing antibody directed against tissue factor (Anti-TF). The data represent the mean ± SEM of six independent experiments.

Next, we analyzed the protein by immunocytochemistry and observed TF on the surfaces of activated platelets that had adhered and spread on immobilized fibrinogen in the presence of thrombin but not on quiescent platelets (Fig. 2 C). Staining was detected on the surface of all the platelets, consistent with the detection of TF mRNA in every cell (Fig. 1 E, top right).

Deletion of factor VIIa from the activity assay or incubation of platelets with a neutralizing anti-TF antibody significantly (P < 0.05) reduced procoagulant activity in stimulated platelets (Fig. 2, D and E). This activity assay, however, evaluates factor Xa generation in the presence of supraphysiologic levels of exogenous factor VIIa (7). Therefore, we determined if platelet-derived TF decreased plasma clotting times. We found that human plasma incubated with platelets for 5 min or 2 h formed clots at an accelerated rate compared with freshly isolated platelets, which by themselves had no appreciable effect on plasma clot formation ( Fig. 2 F). Clotting was significantly delayed in the presence of an inhibitory anti-TF antibody (Fig. 2 F), indicating that TF generated by activated platelets is capable of accelerating in vitro clot formation in humans. Collectively, the studies depicted in Fig. 2 demonstrate that bioactive TF protein accumulates in platelets adherent to fibrinogen in the presence of thrombin. The role of platelet-derived TF in the propagation and stabilization of platelet clots in vitro and in vivo will require further studies using both human and murine thrombosis models.

Clk1 interfaces with the splicing machinery in platelets

The intracellular signaling pathways that control TF pre-mRNA splicing and activity in platelets are not known (18). In nucleated cells, serine-arginine (SR)–rich proteins regulate splicing, and we recently found that human platelets contain the SR family member SF2/ASF (18). Thus, we considered that platelets possess critical upstream kinases that regulate SF2/ASF activity and focused on Clk family members because one of them, Clk1, contains an N-terminal region enriched in arginine-serine dipeptides (RS) that interacts with SF2/ASF (21). We found Clk1 protein in the cytoplasm of mature megakaryocytes (unpublished data), in proplatelets that extend from the megakaryocytes (Fig. 3 A, arrows), and in quiescent circulating platelets from human blood (Fig. 3 B, arrows). In activated platelets, Clk1 was distributed to the tips of F-actin stress cables (Fig. 3 B). Intracellular redistribution of Clk1 resembles the accumulation of vinculin in focal adhesion complexes of platelets that are spread and activated on immobilized fibrinogen (unpublished data and reference 22). Preliminary screens for other family members (Clk2, 3, and 4) were negative (unpublished data), suggesting that Clk1 is the primary Clk in mature, circulating platelets.

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Figure 3.

Platelet Clk1 induces SF2/ASF phosphorylation. (A) Megakaryocytes with proplatelet extensions were stained with anti-Clk1 (top) or control IgG (bottom). The right panels are overlays where anti-Clk1 or IgG are in red, wheat germ agglutinin is in green, and topro-3, which stains nuclei, is in blue. (B) Platelets were left quiescent (top) or activated by adherence to immobilized fibrinogen in the presence of thrombin for 1 h (middle and bottom). The cells were stained with anti-Clk1 (top and middle) or control IgG (bottom). The right panels are overlays in which anti-Clk1 or IgG is in red, polymerized actin is in green, and colocalization of the two markers is identified by yellow staining. A and B are representative images of multiple cells taken from three and four independent experiments, respectively. (C and D) Platelets were activated as in B, lysed, and immune complex kinase assays for Clk1 activity were performed with recombinant SF2/ASF as the substrate. C shows a time course of Clk1-dependent SF2/ASF phosphorylation in activated platelet lysates. D shows Clk1-dependent SF2/ASF phosphorylation in freshly isolated platelet lysates or platelets that were pretreated with vehicle or Tg003 (Clk Inh) (lanes 2–5). The bars for C and D represent fold increases in SF2/ASF phosphorylation over baseline as estimated by densitometry. The gels in C and D are representative of three independent experiments.

In nucleated cells, Clk1 directly phosphorylates SF2/ASF and alters the intracellular localization patterns of SR proteins (21). Therefore, we captured endogenous platelet Clk1 by immunoprecipitation and determined if it regulates SF2/ASF phosphorylation. Clk1 from activated platelets markedly increased SF2/ASF phosphorylation (Fig. 3 C). Increased SF2/ASF phosphorylation was not seen when control IgG was used as the immunoprecipitating reagent (Fig. S4 A, available at http://www.jem.org/cgi/content/full/jem.20061302/DC1).

A benzothiazole compound, Tg003, suppresses Clk1-catalyzed phosphorylation and thereby inhibits SF2/ASF-dependent splicing of in vitro–transcribed pre-mRNAs in immortalized cell lines (23). In stimulated platelets Tg003, but not its vehicle, suppressed Clk1-dependent SF2/ASF phosphorylation (Fig. 3 D), consistent with previous characterization of the inhibitor in other cells (23).

Clk1 controls the expression of TF in activated platelets

We next asked if interruption of signaling from Clk1 to SF2/ASF modulates activation-dependent splicing and found that Tg003 prevented processing of TF pre-mRNA in activated platelets (Fig. 4 A). This Clk1-dependent splicing inhibitor also blocked the expression of TF protein in activated platelets and platelet-derived microparticles (Fig. 4 B). Consistent with its effect on protein, Tg003 significantly (P < 0.05) reduced TF-dependent procoagulant activity in stimulated platelets (Fig. 4 C) and delayed the onset of plasma clot formation (Fig. 4 D). Puromycin, an inhibitor of mRNA translation, also significantly (P < 0.05) reduced TF-dependent procoagulant activity demonstrating the increases were caused by de novo protein synthesis (Fig. S5, available at http://www.jem.org/cgi/content/full/jem.20061302/DC1).

Although Tg003 blocked SF2/ASF phosphorylation (Fig. 3 C) and the expression of TF protein in activated platelets (Fig. 4), it had no effect on other platelet functional responses that included cellular adherence and spreading, actin polymerization, organization of β-tubulin, or the redistribution of Clk1 to focal adhesion complexes (Fig. S6, available at http://www.jem.org/cgi/content/full/jem.20061302/DC1). These data suggest that the Clk1 signaling pathway primarily interfaces with the splicing machinery in platelets.

mRNA detection systems.

For most of the studies, primers that targeted sequences in exon four (5′-CTCGGACAGCCAACAATTCAG-3′) and five (5′-CGGGCTGTCTGTACTCTTCC-3′), and thus spanned intron four, were used to determine endogenous splicing of TF pre-mRNA in platelets. Indirect in situ hybridization or direct in situ PCR was used to detect TF pre-mRNA in megakaryocytes and platelets as previously described (18).

Protein detection systems.

Detailed strategies for protein detection by flow cytometry, Western blot analysis, and immunocytochemistry have been previously published (18, 29).

Measurement of TF-dependent procoagulant activity.

TF-dependent procoagulant activity was calculated with an Actichrome TF assay (American Diagnostica) as previously described (13).

Plasma clotting.

Platelets were left quiescent or activated in the presence or absence of Tg003. Platelet membranes were isolated and added to human plasma (37°C), and clotting was initiated with CaCl₂ as previously described (7).

Immune complex kinase assay for Clk1 activity.

Clk1 activity in platelets was determined using an immune complex kinase assay. An antibody against Clk1 was used for immunoprecipitation of the protein. Nonimmune rabbit IgG was used as a control, and in select experiments recombinant SF2/ASF was removed from the assays to screen for nonspecific incorporation of radiolabeled phosphate (Fig. S4, A and B). Kinase assays were performed by addition of recombinant SF2/ASF (Protein One) in the presence of γ-[³²P]ATP (MP Biomedicals). At the end of this incubation period, the agarose beads and immune complexes were removed by centrifugation, and the unbound sample, which contained SF2/ASF, was resolved by SDS-PAGE.

Statistical analyses.

ANOVA was conducted to identify differences that existed among multiple experimental groups. If significant differences were found, a Student-Newman-Keuls post-hoc procedure was used to determine the location of the difference. For all of the analyses, P < 0.05 was considered statistically significant.

We thank the Labor and Delivery staff of Cottonwood Hospital for collecting cord blood for stem cell differentiation studies and Donnie Benson and Jessica Phibbs for sample pick-up and delivery. We also thank the University of Utah Fluorescence Microscopy Core for use of its facilities and, especially, Chris Rodesch and Don Gentry for their technical assistance. Finally, we thank Diana Lim for preparation of the figures and consultation regarding effective display of the images.

This work was supported by the National Institutes of Health (HL-66277 to A.S. Weyrich, HL-44525 to G.A. Zimmerman, HL-75507 to L.W. Kraiss, and HL-48872 to N. Mackman) and an Established Investigator Award from the American Heart Association (A.S. Weyrich). H. Schwertz was supported by a Deutsche Forschungs Gemeinschaft grant (SCHW 1167/1-2) and then a Western Affiliate American Heart post-doctoral fellowship. M.M. Denis was supported by a physician-scientist training award from the American Diabetes Association (ADA PID2206052).

The authors have no conflicting financial interests.