Selective factor VIII activation by the tissue factor–factor VIIa–factor Xa complex

Blood clotting in response to tissue injury is key for hemostasis¹  and innate immunity²  but can cause vascular thrombosis, leading to serious diseases.^1,3  In the current coagulation scheme (Figure 1A), the extrinsic pathway initiation complex of tissue factor (TF) with active factor VIIa (FVIIa) promotes a cascade of proteolytic reactions yielding FXa; this combines with FVa in the prothrombinase complex, converting prothrombin to thrombin. This initially generated thrombin activates the FVIII and FV cofactors in feedback reactions that amplify coagulation,¹  but active cofactor generation prior to significant thrombin production is also possible, and indeed, FXa is now viewed as the relevant FV activator during coagulation initiation.⁴  Whether FXa^5,-7  or TF-FVIIa^8,9  contribute to coagulation initiation through direct FVIII activation remains unclear.

Extrinsic coagulation initiation is controlled by the TF pathway inhibitor (TFPI), which by inactivating FVIIa and FXa within a quaternary complex with TF^10,11  attenuates thrombosis.¹²  Moreover, a positively charged TFPIα carboxyl terminal region interacts with a specific acidic sequence in partially processed FV interfering with FXa formation of active prothrombinase.^13,14  These mechanisms reducing direct thrombin generation are compensated for by TF-FVIIa activating the intrinsic pathway FIX¹⁵  in a kinetically favored reaction in the presence of physiologic plasma inhibitors.¹⁶  Alternatively, FIXa is generated by FXIa activated in a thrombin feedback loop,¹⁷  also promoting vascular inflammation,¹⁸  or by contact phase FXIIa.¹⁹  In mouse models,²⁰  FXIIa contributes to amplified thrombin generation in experimental thrombosis but, consistent with human data,²¹  has no role in hemostasis.

The current coagulation paradigm, with the expanded function of TFPI tightly controlling both TF-dependent initiation and prothrombinase generation, cannot readily explain how initially produced thrombin can be the origin of FVIIIa cofactor for FIXa produced by the contact phase pathway or by TF-FVIIa itself. Here, we outline a novel function of the extrinsic coagulation initiation complex whereby nascent FXa associated with TF-FVIIa directly activates FVIII, resisting inhibitor control by TFPI. Such a mechanism may be relevant for the function of TF in hemostasis and provides new perspectives for interpreting the distinct roles of coagulation reactions in physiologic and pathologic thrombus formation.

TF-coated glass coverslips were perfused with venous blood at a wall shear rate of 300 s⁻¹ (see the supplemental Materials and methods, available on the Blood Web site). A Zeiss Axiovert 135M/LSM 410 microscope with a Plan-Apochromat ×40/1.40-NA oil immersion objective (Carl Zeiss AG, Oberkochen, Germany) was used to visualize platelets/leukocytes and fibrin stained with mepacrine and a specific antibody, respectively. Image analysis to calculate volumes was performed as has been described.²² 

TG in platelet-rich plasma (PRP) or reconstituted PRP was evaluated as has been described.²³  Platelets in PRP were adjusted to 180 ⋅ 10³ per microliter with homologous platelet-poor plasma (PPP), and corn trypsin inhibitor (CTI) was added at 30-50 μg/mL. Reconstituted PRP was prepared with washed platelets resuspended at 180 ⋅ 10³ per microliter into PPP. TG was initiated by adding recombinant TF (rTF), FIXa, or both at defined concentrations with 18 mM CaCl₂ into microtiter-plate wells containing 360 µM benzyloxycarbonyl-glycyl-glycyl-l-arginine coupled to fluorogenic 7-amido-4-methylcoumarin (Bachem Americas, Torrance, CA) as thrombin substrate. The rate of fluorescence intensity increase (measured at 355/460 nm excitation/emission) as a function of time (dF/dt) was calculated with Turbo Delphi 2006 (Borland Software Corporation, Austin, TX) and converted to thrombin-equivalent concentration (nM) using a calibration curve. TG was also measured with a discontinuous 2-stage assay with a detection limit of 5 pM. In this assay, reactions incubated for up to 11 min at 37°C were terminated with 20 mM EDTA, and generated thrombin was measured using the sensitive thrombin substrate H-D-CHA-Ala-Arg-AMC⋅2AcOH (Pefafluor TH, Pentapharm, Basel, Switzerland) at 50 µM.

Coagulation products were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis under reducing conditions; only reactions containing anti-FVIIa monoclonal antibody (MoAb) were processed under nonreducing conditions to avoid confounding effects from immunoglobulin G (IgG) heavy chains with a molecular mass comparable to FVIII A1 domain. Proteins transferred to polyvinylidene fluoride membranes were probed with anti-FVIII MoAb C5 (0.5 µg/mL)²⁴  or anti-FV AHV-5146 (1 µg/mL). FVIIIa and FVa were quantified by infrared detection with the Odyssey infrared imager (Li-COR, Lincoln, NE), calibrated with known FVIIIa and FVa quantities.

Reactions in 50 mM Tris-buffered saline, pH 7.4, with 0.1% bovine serum albumin included 0.7 nM FVIII, 3 nM FV, 135 nM FX, and 1 µM prothrombin without or with 4 μM dansylarginine N-(3-ethyl-1,5-pentanediyl)amide (DAPA). TFPIα and other inhibitors were added as indicated. Reactions were initiated by rTF (50 or 400 pM) with FVIIa (200 or 500 pM), FIXa (2 or 10 nM), or both added with 2.5 mM CaCl₂ and incubated at 37°C for the indicated times. After quenching the reaction with 10 mM EDTA, generated FXa was measured with S-2765 (180 µM). FVIIIa procoagulant activity was measured as FIXa-dependent FXa generation; FVIIIa procoagulant activity generated by the nascent TF-FVIIa-FXa complex was calculated by subtracting the amount of FXa produced in reactions initiated by FVIIa and FIXa individually from that produced in reactions initiated by FVIIa/FIXa combined. When indicated, 200 nM lepirudin was used to inactivate possible thrombin contamination.

Animal procedures complied with the Guide for Care and Use of Laboratory Animals and were approved by the Animal Care and Use Committee of The Scripps Research Institute. Vascular injury was induced in C57BL/6J mice by 1 0.8-μL drop of 7% (0.26 M) or 8% (0.30 M) FeCl₃⋅6H₂O applied on the carotid artery for 3 min; or a 0.4-μL 4% (0.15 M) drop for 1 min on the femoral vein, followed by rinsing. Antibodies were administered by bolus injection into the catheterized jugular vein. FVIII and FVIIIa were administered by a bolus injection (1.4 pmol), followed by maintenance with continuous infusion at the rate of 0.47 pmol/min for 15 min. Time to first occlusion after injury and flow index were quantified as has been described.²²  Additional details for experimental procedures are described in the supplemental Methods.

Studies involving human subjects were approved by appointed institutional review boards. Human volunteers gave informed consent to participate in the studies before blood collection, and experiments were performed in accordance with the Declaration of Helsinki.

Group variances were evaluated with Levene’s median and Bartlett’s tests; differences were evaluated with 1-way analysis of variance (ANOVA) or the Kruskal-Wallis test, followed by Tukey’s and Dunn’s tests, respectively, for multiple comparisons. Data were transformed as y = log₁₀y when necessary to obtain homoscedasticity. Software packages used were GraphPad Prism (version 7; GraphPad Software, La Jolla, CA) and XLSTAT (Addinsoft, Paris, France).

As was previously shown, contact phase FXII^20,25,26  and TF^22,27  contribute to experimental thrombosis in the ferric chloride-induced carotid artery occlusion model, but how this happens remains unclear. We found that MoAbs blocking the TF function²²  or FXI activation by FXIIa²⁵  independently reduced occlusion after a vessel wall lesion caused by 7% (0.26 M) FeCl₃ (Figure 2A). After a lesion by 8% (0.3 M) FeCl₃ the same MoAb concentrations were ineffective, but a higher dose of the anti-FXI MoAb, not of the anti-TF, still prevented thrombosis (Figure 2A). Thus, even after the more severe lesion, thrombogenesis required FIXa generation by FXIIa-FXIa.^20,25  Remarkably, under the latter conditions, combining individually inactive low doses of anti-TF and anti-FXI MoAbs reduced arterial occlusion significantly (Figure 2A), confirming a role for the TF pathway in this intrinsic coagulation-dependent model of thrombus formation. To explain such findings, we hypothesized that TF might contribute to activate FVIII, the essential cofactor for the intrinsic tenase protease, FIXa. As visualized in the femoral vein, FVIIIa, but not FVIII, prevented inhibition of FeCl₃-induced fibrin deposition by the low-dose anti-TF/anti-FXI MoAb combination but not by high-dose anti-FXI alone (Figure 2B-C). These results ruled out the hypothesis that FIXa generated by TF-FVIIa or alternative pathways used exogenously provided FVIIIa to trigger thrombosis, supporting the idea that TF-FVIIa contributes to FVIII activation in vivo.

To challenge this concept, we measured TG in PRP. In reactions containing an inhibitory polyclonal antibody blocking TFPI and CTI preventing FXI activation by FXIIa,²⁸  we defined the concentrations of FIXa or relipidated rTF yielding comparable TG (Figure 2D, left). At the same concentration but without TFPI blockade (Figure 2D, right), rTF produced little thrombin and late in the reaction, but enhanced TG with FIXa added concurrently. This amplification of FIXa-triggered TG required FVIII at <10% plasma concentration (supplemental Figure 1) but not FIX (Figure 2E), excluding additional TF-dependent FIXa generation.^16,17  Because FVIII concentrations sufficient to prevent severe spontaneous bleeding in FVIII-deficient patients²⁹  could support synergistic TG amplification, the conditions of this assay appear to be relevant for assessing hemostatic competence in PRP.

We explored the mechanism of TF-induced priming of intrinsic coagulation using a sensitive 2-stage TG assay. In FVII-deficient reconstituted PRP, 0.15 pM rTF with FVIIa wild-type (WT) produced little thrombin (<20 pM in 11 min), similar to that generated by the active site mutant FVIIa S195A (iFVIIa)³⁰  or FIXa alone (Figure 3A). However, combining FIXa with rTF and FVIIa WT, but not inactive iFVIIa, amplified TG ∼5-20 times in 5-11 min (Figure 3A). Thus, the additive quantities of thrombin produced in this assay by TF-FVIIa and FIXa separately were far less than those yielded by the 2 combined, demonstrating a synergistic interaction linking TF-initiated and intrinsic coagulation upstream of thrombin generation.

To identify the reaction that, apart from thrombin, could yield FVIIIa for FIXa-dependent coagulation, we considered TF-FVIIa^8,9  and FXa^5,-7  as potential FVIII activators. First, we determined that TF-FVIIa without FX (or TF-FX without FVIIa) generated no or minimal FVIIIa activity but that TF-FVIIa with FX (generating FXa) produced substantial amounts of FVIIIa with all reactants at physiologically relevant concentrations (Figure 3B). To distinguish between functions of FXa released from or still associated with TF-FVIIa, we formed a stable TF-iFVIIa-FXa complex with the nematode anticoagulant protein (NAP) c2.^31,32  Use of iFVIIa excluded FVIIa catalytic activity, whereas FXa is known to retain catalytic function in the formed NAPc2 complex.³²  This complex failed to induce TG in FVII-deficient reconstituted PRP but markedly enhanced FIXa-induced TG (Figure 3C), suggesting that FXa associated with TF-FVIIa can activate FVIII. As was expected, a similar complex formed with TFPIα, which inhibits FXa, was inactive alone and did not support FIXa-dependent TG in PRP (Figure 3C). Surprisingly, the NAPc2-stabilized TF-iFVIIa-FXa complex could activate purified FVIII but not the homologous cofactor FV (Figure 3D). Specific inhibition with tick anticoagulant peptide (TAP) confirmed that FXa in the stabilized TF-iFVIIa-FXa complex activated FVIII (Figure 3D), implying that nascent FXa still associated with TF-FVIIa can exert the same function (Figure 3E).

We next evaluated whether free FXa, which triggers coagulation by forming the prothrombinase complex, could be a physiologic FVIII activator akin to its function in generating the prothrombinase cofactor, FVa.⁴  On the same procoagulant membrane used in the experiments above, increasing concentrations of FVa markedly stimulated prothrombin conversion by a low concentration of FXa (Figure 3F). In contrast, generation of FVIIIa activity (Figure 3G) or proteolytic cleavage of FVIII (Figure 3H) by the same FXa concentration was progressively inhibited by adding FVa in the same concentration range, indicating that FXa bound to FVa cannot efficiently activate FVIII. Thus, nascent FXa associated with TF-FVIIa preferentially activates FVIII, triggering the intrinsic coagulation pathway. Subsequent FXa transfer from TF-FVIIa into the prothrombinase complex with FVa marks the transition to direct TF-induced coagulation.

To assess the relative roles of the nascent TF-FVIIa-FXa complex and thrombin in generating FVIIIa, we studied procofactor activation on rTF-bearing phospholipid vesicles mixed with purified FX, prothrombin, FVIII, and FV. Activation of FVIII after FVIIa addition was partially inhibited by blocking thrombin with DAPA or by replacing normal prothrombin with the inactive S195A mutant,³³  but ∼15% FVIIIa was still detectable under both conditions (Figure 4A). Importantly, although thrombin could generate more FVIIIa, as was expected from efficient FVIII cleavage in the solution phase, the amount of VIII activated by the TF-initiated reaction in the absence of active thrombin was sufficient for full function of membrane assembled FVIIIa-FIXa intrinsic tenase complex (Figure 4B; supplemental Figure 2A). In agreement with the results in PRP containing endogenous coagulation inhibitors (see Figure 2D), TFPIα that markedly suppressed direct FXa generation by TF-FVIIa (supplemental Figure 2B) had limited effect on the formation of functional FVIIIa-FIXa complex in TF-initiated reactions (Figure 4C). The latter was also resistant to inhibition by TFPIα with the cofactor protein S^34,35 ; partial inhibition by protein S alone likely resulted from competition for limited procoagulant surfaces (supplemental Figure 2C). Consistent with the observed thrombin-independent FVIII activation by nascent FXa in PRP, FVIII activation by TF-FVIIa-FXa was not influenced by von Willebrand factor (VWF) binding FVIII (supplemental Figure 2D).

To demonstrate directly that rTF supports FVIII activation in a physiological plasma milieu independently of thrombin feedback reactions, we used hirugen (63-O-sulfo-tyr-hirudin[55-65]) to block thrombin exosite I required for cofactor activation.^36,37  Hirugen dose-dependently inhibited FVIII activation by thrombin (supplemental Figure 3A) but not FXa generated by TF-FVIIa-FIXa (supplemental Figure 3B). In PRP with CTI and anti-FXI MoAb to block feedback TG amplification through increased FIXa generation by FXIa,³⁸  2 μM of hirugen blocked FIXa-dependent TG (Figure 4D), demonstrating that this reaction required thrombin feedback activation of FVIII. In contrast, the same hirugen concentration failed to inhibit TG by FIXa combined with rTF, even though no TG could be detected when FIXa and rTF were added separately (Figure 4E). Thus, the TF pathway activates FVIII in plasma when thrombin feedback loops are inhibited. Experiments with mouse microvesicles³⁹  generated from WT or human TF knock-in⁴⁰  macrophages showed that thrombin-independent FVIIIa generation occurred also on a natural procoagulant surface with human or mouse TF-FVIIa (supplemental Figure 3C).

To elucidate further how FXa generation by TF-FVIIa distinctly contributes to intrinsic pathway activation as opposed to direct extrinsic pathway TG, we studied 2 FVIIa mutants, T99Y and E154A, that cleave FX but display very low substrate turnover because of impaired FXa release.^30,41  In a phospholipid-free assay or with phospholipid-reconstituted rTF, the FVIIa mutants produced an initial burst but, in contrast to FVIIa WT, could not sustain FXa generation (Figure 5A). Remarkably, TF complexes with both mutants were comparable to FVIIa WT in supporting FXa-dependent FVIII activation (supplemental Figure 4A), and importantly, TFPIα at supraphysiological concentrations (10 nM) did not appreciably influence this pathway of FVIIIa generation (Figure 5B). In marked contrast to FVIII, the FVIIa mutants failed to induce FV activation, and TFPIα inhibited FVa generation induced by FVIIa WT (Figure 5C).

Both FVIIa mutants with impaired FXa turnover formed a functional FVIIIa-FIXa intrinsic tenase complex when FIXa was available, but only the exosite mutant E154A produced FVIIIa-IXa activity when zymogen FIX was present instead (Figure 5D), in agreement with the fact that FVIIa T99Y,³⁰  unlike E154A, cannot activate FIX (supplemental Figure 4B). Thus, complementing the ability to generate FIXa,^15,16  direct activation of the antihemophilic FVIII cofactor by nascent FXa product of TF-FVIIa enables intrinsic pathway coagulation before TFPIα inhibitory control. We tested these conclusions in FVII-deficient reconstituted PRP in which, besides endogenous plasma coagulation inhibitors, platelets are an additional source of TFPIα.¹¹  Under these conditions, FVIIa WT, but not E154A or T99Y mutants, induced TG in the presence of a neutralizing anti-FVIII MoAb (Figure 5E), confirming that the mutants could not directly generate thrombin in a plasma milieu. Without FVIII inhibition, TG by FVIIa E154A was only slightly slower than that by WT, whereas TG by FVIIa T99Y was clearly decreased. FXIa inhibition reduced markedly TG by FVIIa T99Y but less by WT or E154A (Figure 5F; supplemental Figure 4C), in agreement with the latter generating FIXa as well as FVIIIa.

To prove directly that thrombin-independent FVIII activation occurred in these reactions, we first verified that the thrombin exosite blocker, hirugen, abolished FIXa-initiated TG in FVII-deficient plasma (Figure 5G). In the presence of the same hirugen concentration, TG by mutant FVIIa E154A, not by FVIIa WT, was entirely FVIII dependent, and without FVIII inhibition, TG induced by FVIIa WT and E154A was of similar magnitude, but TG by the latter was clearly delayed (Figure 5H). To explain the delay, we reasoned that impaired direct FXa generation by the mutant FVIIa could reduce FVa cofactor generation for initial prothrombinase assembly. Indeed, adding FVa normalized the delay in FVIIIa-dependent TG by FVIIa E154A (Figure 5I). Accordingly, adding FVa to normal PRP accelerated TF-initiated TG but not more than did blocking TFPI function (supplemental Figure 4D). These data indicate that prothrombinase activity is regulated by TFPI control of FXa generation^13,14  that contributes to FV activation⁴  and reinforce the concept that FVIII activation during TF-initiated coagulation generates FVIIIa-FIXa intrinsic tenase activity independently of thrombin feedback reactions and escaping TFPI control.

We then screened a library of MoAbs to FVIIa to identify a proof of principle inhibitor that could recapitulate the shift in functional properties (loss of efficient FXa and FIXa generation but not of FVIII activation) seen with the FVIIa mutant T99Y. In contrast to the fully inhibitory MoAb 3G12, antibody 12C7 had no effect on FVIII activation (Figure 6A). Moreover, it had no significant effect on the generation of intrinsic tenase activity when FIXa was present, but it markedly inhibited when FIX was supplied instead (Figure 6B). MoAb 12C7 attenuated TG in PRP but, as seen with FVIIa T99Y, rendered TG FXIa dependent (Figure 6C). Thus, results with mutant FVIIa molecules and inhibitory antibodies concordantly show that the TF-FVIIa complex can initiate intrinsic and extrinsic coagulation pathways in distinct reactions.

To evaluate whether TF-FVIIa initiates thrombus formation in flowing blood ex vivo when direct initial thrombin generation is limited, we surface-immobilized rTF at a low concentration sufficient for FVIII-dependent fibrin formation at a wall shear rate of 300 s⁻¹. Under these conditions established with WT FVIIa, FVIIa T99Y were less thrombogenic in spite of supporting more platelet adhesion than were inactive FVIIa S195A (Figure 7A-B). Addition of 10 pM FIXa to blood containing FVIIa T99Y, but not containing inactive FVIIa S195A, restored FVIII-dependent fibrin formation (Figure 7A-B). In contrast, mutant FVIIa E154A, which is as defective as T99Y in direct TG (Figure 5E), supported FVIIIa-dependent thrombus formation similar to FVIIa WT when added without FIXa to FVII-deficient reconstituted blood (Figure 7A,C). This confirmed that the nascent FXa product of TF-FVIIa can directly generate FVIIIa functioning in the intrinsic tenase complex with the potential to enhance hemostasis in low-TF environments with limited direct TF-dependent thrombin generation activating feedback loops.

Our findings delineate a novel function of the extrinsic TF-FVIIa complex, namely, providing selective feed-forward activation of the FVIII antihemophilic cofactor independently of thrombin feedback loops (Figure 1B). This specific reaction of nascent FXa escapes control by physiologic coagulation inhibitors in PRP or TFPIα in purified systems. Together with generation of the FIXa antihemophilic protease by TF-FVIIa,^15,16  direct FVIII activation by TF-FVIIa-nascent FXa completes a pathway to FVIIIa-FIXa intrinsic tenase activity fully integrated within TF-initiated coagulation preceding direct activation of the common coagulation pathway.

Nascent TF-FVIIa-FXa generates FVIIIa, facilitating the formation of intrinsic tenase but without providing FVa for prothrombinase activity. Generating the latter requires FXa undocking from TF-FVIIa, thus exposing free FXa to inhibitory control. Therefore, the newly identified TF-FVIIa-FXa function allows for accumulation of active prohemostatic antihemophilic FVIIIa cofactor without increasing prothrombotic FVa. This may be of relevance for targeted FXa anticoagulants that, with comparable antithrombotic potency, have a lesser impact on hemostasis than do vitamin K antagonists.^42,-44  Of note, such a mechanism is independent of thrombin feedback reactions and FXI activity and may thus support hemostasis during treatment with thrombin inhibitors or recently validated strategies targeting FXI.⁴⁵ 

Selectivity for cofactor activation indicates distinct functional properties of FXa in complex with or released from TF-FVIIa. Although coagulation cofactor-enzyme complexes are typically geared toward efficient substrate turnover for rapid thrombin generation, throughout evolution the TF initiation complex appears to have preserved mechanisms favoring its stability. FX interacts with TF-FVIIa through an extended interface that is minimally affected by zymogen to enzyme transition.^46,-48  In this interface, FVIIa residue E154, conserved across species, transmits conformational changes from the substrate-occupied active site of the protease⁴¹  and may thereby regulate subsequent product release. Elimination of this conformational switch was sufficient to segregate macromolecular substrate FX activation and product FXa turnover by TF-FVIIa. Thus, mutant FVIIa E154A helped inform on direct FVIII activation by nascent FXa associated with TF-FVIIa as well as the contribution of this novel pathway to thrombin generation and thrombogenesis in platelet-rich plasma and whole blood under flow.

Stability of the TF coagulation initiation complex likely represents the evolutionary advantage of preserving key signaling roles of TF-FVIIa-FXa that link coagulation activation and innate immunity.³²  In line with efficient FVIII activation, FVIIa T99Y is fully functional in mediating TF-FVIIa-FXa activation of protease activated receptor (PAR) 2.^30,32  Moreover, as seen for FVIIIa generation, resistance to functional inhibition by TFPIα is also an important feature of PAR signaling induced by TF-FVIIa-FXa in endothelial cells.⁴⁹  This signaling complex is additionally stabilized by recruitment of the FXa-binding partner, endothelial protein C receptor (EPCR), in mouse and man.^50,51  A key innate immune signaling role for the TF-FVIIa-FXa-EPCR complex recently emerged in dendritic cells,⁵²  in which it is essential for toll-like receptor 4 induction of interferon-regulated genes. Negative regulation of this pathway by the alternative EPCR ligand, activated protein C, utilizes the canonical anticoagulant cofactor functions of FV and protein S.⁵³  Thus, these and other nontraditional functions of the coagulation system⁵⁴  likely use the same mechanistic features that simultaneously serve diverse roles in immunity, hemostasis, and injury repair.

Our findings provide the biochemical bases for defining distinct roles of TF in supporting hemostasis or contributing to thrombosis. One can envision application of mutants of FVIIa or the described antibody reagents to define the partial functions of TF in triggering thrombogenesis directly or provide activation of the intrinsic pathway of relevance for hemostasis. Ultimately, this should lead to the possibility of selectively assessing the effects of recent and future new anticoagulants on the dual roles of TF in hemostasis and thrombosis. The novel concepts on coagulation presented here may also have implications for the development and evaluation of hemostatic agents, providing protection from severe bleeding complications while avoiding adverse thrombotic complications in patients with underlying vascular pathologies.

The authors thank Hiroshi Deguchi and Thomas Siegemund for technical advice on TG assays.

This work was supported by grants from the National Institutes of Health, National Heart, Lung, and Blood Institute (HL-117722 [Z.M.R.], HL-31950 [W.R. and Z.M.R.], and UM1 HL120877 [W.R.]), the MERU Foundation, Italy, and the Roon Research Center for Arteriosclerosis and Thrombosis at The Scripps Research Institute (Z.M.R.); and the Humboldt Foundation, Germany (W.R.).

Contribution: Y.K. planned research, performed and interpreted coagulation studies, and contributed to writing the manuscript; G.L.M., A.Z., P.M., and J.N.O. performed mouse thrombosis and ex vivo thrombogenesis studies; A.S.R. prepared and characterized macrophage-derived microparticles; A.J.G., S.K., A.G., H.Ø., and L.C.P. provided essential reagents and reviewed the manuscript; and W.R. and Z.M.R. conceived and planned the study, interpreted results, and wrote the manuscript.

Conflict-of-interest disclosure: Z.M.R. is founder, President, and CEO of MERU-VasImmune, Inc., and has equity interest in Sedicidodici, s.r.l., 2 companies that are developing coagulation assays based on technology related to work presented in this article. Y.K. is a part-time employee of MERU-VasImmune. Oregon Health & Science University and A.G. may have a financial interest in the results of this study. H.Ø. and L.C.P. are employees of Novo Nordisk. The remaining authors declare no competing financial interests.

Correspondence: Zaverio M. Ruggeri, Department of Molecular Medicine, The Scripps Research Institute, MEM-175, 10550 N. Torrey Pines Rd, La Jolla, CA 92037; e-mail: ruggeri@scripps.edu; and Wolfram Ruf, Department of Immunology and Microbiology, The Scripps Research Institute, 10550 N. Torrey Pines Rd, La Jolla, CA 92037; e-mail: ruf@scripps.edu.