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Innate immunity represents an early defense mechanism against invading pathogens. Whereas in mammals this is complemented by adaptive immunity, host defenses in many other animal species, including arthropods—the largest phylum among animals—relies exclusively on innate immunity. In arthropods, which are endowed with an unusually efficient antimicrobial system, host defense depends on coagulum formation by the hemolymph1. This pre-vents pathogen intrusion into the circulatory system and hinders pathogens from invading host cells. A similar cross-talk between innate immunity and coagulation has been postulated to persist in mammals2–6. Indeed, blood neutrophils, major effectors of mam-malian innate immunity, accumulate rapidly at sites of vessel injury in concert with aggregating platelets, even in large arteries exposed to high shear forces and without a pathogen challenge. Whereas neutrophil-related procoagulant mechanisms might contribute to large-vessel thrombosis in various types of cardiovascular dis-ease (for example, myocardial infarction7,8, sepsis9 and thrombo-cythemia10), it is largely undefined how neutrophils control fibrin generation in vivo and whether this activation of coagulation affects antimicrobial defenses. Upon activation, neutrophils release the serine proteases neutrophil elastase and cathepsin G to inactivate pathogens such as Gram-negative bacteria and fungi11–14. Here we show that these neutrophil serine proteases, in concert with exter-nalized nucleosomes, promote thrombus formation inside blood vessels. This thrombus formation helps retain bacteria inside liver
microvessels and thereby prevents the extravasation of pathogens, directly supporting antimicrobial defense.
RESULTSNeutrophil serine proteases promote coagulation in vivoThe neutrophil serine proteases neutrophil elastase and cathepsin G are able to degrade regulators of blood coagulation in vitro15–21, although it is unknown whether this translates into a role of neutrophil serine proteases in coagulation in vivo. To address this issue, we used intravital videomicroscopy22,23 to monitor fibrin generation within injured carotid arteries of mice deficient in neutrophil elastase and cathepsin G (Elane−/−;Ctsg−/− mice) or of wild-type (WT) mice. In WT mice, fibrin formation occurred within 5 min after vessel injury and reached a peak within 15 min (Fig. 1a,b). In line with previous reports24,25, fibrin formation in response to FeCl3-induced vessel injury required both tissue factor, the initiator of the extrinsic path-way, and factor XII, the trigger of the contact pathway: fibrin forma-tion was strongly reduced in WT mice pretreated with neutralizing antibody to mouse tissue factor or with the factor XII inhibitor Pro-Phe-Arg-chloromethylketone (PCK) (Supplementary Fig. 1a). When we induced vascular injury in Elane−/−;Ctsg−/− mice, fibrin formation 5 min after FeCl3 exposure was similar to that observed in control mice; however, it was markedly reduced 10–15 min after vessel injury and remained low throughout the entire course of the experiment (Fig. 1a,b). Likewise, the fibrin-covered area as well as
1Deutsches Herzzentrum, Technische Universität, Munich, Germany. 2Institut für Klinische Chemie, Ludwig-Maximilians-Universität, Munich, Germany. 3Hämostaseologie, Ludwig-Maximilians-Universität, Munich, Germany. 4Max-Planck-Institut für Infektionsbiologie, Berlin, Germany. 5Institut für Biochemie, Justus-Liebig-Universität, Giessen, Germany. 6These authors contributed equally to this work. Correspondence should be addressed to B.E. ([email protected]).
Received 11 February; accepted 21 June; published online 1 August 2010; doi:10.1038/nm.2184
Reciprocal coupling of coagulation and innate immunity via neutrophil serine proteasesSteffen Massberg1,6, Lenka Grahl2,6, Marie-Luise von Bruehl1,6, Davit Manukyan2,3,6, Susanne Pfeiler2, Christian Goosmann4, Volker Brinkmann4, Michael Lorenz1, Kiril Bidzhekov2, Avinash B Khandagale2, Ildiko Konrad1, Elisabeth Kennerknecht1, Katja Reges2, Stefan Holdenrieder2, Siegmund Braun1, Christoph Reinhardt2, Michael Spannagl3, Klaus T Preissner5 & Bernd Engelmann2
Blood neutrophils provide the first line of defense against pathogens but have also been implicated in thrombotic processes. This dual function of neutrophils could reflect an evolutionarily conserved association between blood coagulation and antimicrobial defense, although the molecular determinants and in vivo significance of this association remain unclear. Here we show that major microbicidal effectors of neutrophils, the serine proteases neutrophil elastase and cathepsin G, together with externalized nucleosomes, promote coagulation and intravascular thrombus growth in vivo. The serine proteases and extracellular nucleosomes enhance tissue factor– and factor XII–dependent coagulation in a process involving local proteolysis of the coagulation suppressor tissue factor pathway inhibitor. During systemic infection, activation of coagulation fosters compartmentalization of bacteria in liver microvessels and reduces bacterial invasion into tissue. In the absence of a pathogen challenge, neutrophil-derived serine proteases and nucleosomes can contribute to large-vessel thrombosis, the main trigger of myocardial infarction and stroke. The ability of coagulation to suppress pathogen dissemination indicates that microvessel thrombosis represents a physiological tool of host defense.
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the overall size of the thrombi were clearly reduced in Elane−/−;Ctsg−/− mice compared to WT mice (Fig. 1c,d). Consistent with inefficient thrombus formation, the time required for occlusion of the carotid artery was also prolonged in Elane−/−;Ctsg−/− mice compared to control mice (Fig. 1e). Even though vessel occlusion was delayed in neutrophil serine protease–deficient mice, it was not fully inhibited. However, the thrombi that formed in Elane−/−;Ctsg−/− mice were more fragile than WT thrombi, and blood flow was reestablished rapidly (<5 min) after initial occlusion (Fig. 1e). In contrast, clo-sure persisted for >10 min in WT mice (Fig. 1e). In mice lacking neutrophil elastase but expressing cathepsin G (Elane−/−;Ctsg+/+), fibrin formation was reduced to a similar degree as observed in Elane−/−;Ctsg−/− mice (Fig. 1f). Infusion of neutrophil elastase (50 μg, this amount being necessary to overwhelm degradation by endo-genous neutrophil elastase inhibitors) restored the coagulation defect in Elane−/−;Ctsg+/+mice (Fig. 1f), suggesting that intravascular fibrin formation in mice relies predominantly on neutrophil elastase. The contribution of neutrophil serine proteases to fibrin formation was not restricted to FeCl3-induced (chemical) vessel injury. Elane−/−;Ctsg−/− mice also showed diminished fibrin generation when we induced vascular injury by mechanical endothelial denudation (carotid liga-tion) (Fig. 1g). This method induces exposure of collagen26 and also initiates fibrin formation via both tissue factor23 and factor XII24, but, in contrast to FeCl3-induced injury, does not result in instantaneous vessel occlusion. Finally, to evaluate the relevance of neutrophil serine proteases for hemostasis, we measured tail bleeding times. Bleeding was prolonged twofold in Elane−/−;Ctsg−/− mice compared to WT mice
(Fig. 1h), indicating an impaired ability of the hemostatic system in neutrophil serine protease–deficient mice to prevent blood loss. Taken together, these data show that neutrophil serine proteases foster intra-luminal fibrin deposition and have a key role in thrombus develop-ment and stabilization.
Although neutrophil serine proteases can proteolytically modify cellular adhesion receptors27, the impaired coagulation and thrombus formation in the absence of neutrophil elastase and cathepsin G did not result from alterations in blood cell adhesion to the injured vessel wall. In fact, leukocyte recruitment was slightly elevated, and platelet adhesion was unchanged, in Elane−/−;Ctsg−/− mice (Supplementary Fig. 1b,c). We also did not observe intrinsic defects in platelet activation, as washed platelets from both WT and mutant mice showed similar shape changes and aggregation kinetics and comparable expression of P-selectin and GPIIb-IIIa (integrin αIIbβ3) (Supplementary Fig. 1d,e). Therefore, loss of neutrophil elastase and cathepsin G predominantly affects fibrin formation and fibrin-dependent thrombus stabilization, whereas leukocyte or platelet recruitment are not compromised.
Proteolysis of tissue factor pathway inhibitor (TFPI)To dissect the mechanisms underlying neutrophil serine protease–driven coagulation, we examined factor X activation by neutrophils in vitro. WT mouse neutrophils alone poorly supported factor X activation, whereas addition of activated WT platelets synergisti-cally increased it (Fig. 2a; in line with human blood cell data28). The potent procoagulant activity resulting from the addition of both
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Figure 1 Neutrophil serine proteases trigger fibrin formation. (a) Fibrin formation in WT and Elane−/−;Ctsg−/− mice in vivo after FeCl3-induced vessel injury. Representative images of fibrin formation (pseudocolored; before injury and 5 and 15 min after injury). Arrows indicate flow direction. Images representative of at least five mice per group. Scale bar, 50 μm. The schematic at left shows the injury site, with the segment highlighted where images were taken. ACE, ACI and ACC: external, internal, and common carotid artery, respectively. At right is the color scale for fibrin signal intensity. (b) Quantitative analyses of fibrin formation. n = 5 (WT) and 8 (Elane−/−;Ctsg−/−), *P < 0.05. (c) Fibrin-positive area 15 min after vessel injury. n = 4 (WT) and 8 (Elane−/−;Ctsg−/−), *P < 0.05. (d) Thrombus size 30 min after injury. n = 9 (WT) and 15 (Elane−/−;Ctsg−/−), *P < 0.05. (e) Thrombus growth kinetics. Left, time to vessel occlusion. *P < 0.05 compared to WT. Right, duration of occlusion. n = 4 (WT) or 6 (Elane−/−; Ctsg−/−), *P < 0.05. (f) Left, fibrin formation in WT, Elane−/−;Ctsg−/− and Elane−/−;Ctsg+/+ mice. *P < 0.05 (versus WT). Values for WT and Elane−/−; Ctsg−/− mice are the same data as in b. Right, fibrin formation after neutrophil elastase (NE) (50 μg in 100 μl PBS or vehicle alone) was infused into Elane−/−;Ctsg+/+ mice. n = 4 or 5 per group, *P < 0.05 compared to WT mice (left) or to absence of NE (right). (g) Fibrin generation after ligation-induced injury of the carotid artery, 15 min after injury. n = 3 per group, *P < 0.05. (h) In vivo tail bleeding time. n = 16 per group, *P < 0.05. Error bars represent s.e.m. in all cases.
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neutrophils and platelets required both tissue factor and factor IXa but not factor XII (Supplementary Fig. 2a), consistent with findings in humans showing that platelets and neutrophils trigger factor Xa formation in a tissue factor– (derived from platelets29 or neutro-phils30) and IXa28-dependent manner. This tissue factor and factor IXa dependence most likely results from the action of two pathways of factor Xa formation. In the first pathway, prothrombin (con-tained in the coagulation factor concentrate applied) is converted to thrombin by platelet-released factor Va; thrombin then triggers formation of factor XIa which, in turn, amplifies factor Xa forma-tion by activation of factor IX. In the second pathway, factor IX is directly activated by the TF-VIIa complex31.
Factor X activation resulting from the concerted action of neutrophils and platelets strictly required the expression of serine proteases by neutro-phils but not by platelets. Factor X activation was strongly reduced when Elane−/−;Ctsg−/− neutrophils were incubated with either mutant or WT platelets (Fig. 2a). In contrast, the procoagulant activity of WT neutro-phils was significantly increased by either WT or Elane−/−;Ctsg−/− plate-lets (Fig. 2a). We next tested the role of neutrophil serine proteases in coagulation using human cells. Factor Xa generation induced by the combination of human neutrophils and human platelets was inhibited by Ala-Ala-Pro-Val-chloromethylketone (CMK; a neutrophil elastase- specific inhibitor), by α1-antitrypsin (an inhibitor of both neutrophil elastase and cathepsin G) or by α1-antichymotrypsin (a cathepsin G inhibitor), but not by the matrix metalloprotease (MMP) inhibitor tissue inhibitor of metalloproteases-1 (TIMP-1) (Supplementary Fig. 2b). Thus, in both mice and humans, neutrophils and platelets cooperate to enhance coagulation in a process depending on neutrophil serine proteases.
Next, we analyzed the molecular mechanisms by which neutro-phil serine proteases promote coagulation. We hypothesized that
neutrophil serine proteases might stimulate coagulation via proteo-lytic inactivation of endogenous anticoagulants15–18. Among these, TFPI is a primary candidate, as it is rapidly secreted by activated platelets29,32. Platelet-released TFPI was considerably degraded in the presence of human neutrophils (which are devoid of TFPI), and this degradation was attenuated by the serine protease inhibitors CMK or α-1-antichymotrypsin (as analyzed in blood cell supernatants; data not shown and ref. 17). To assess whether neutrophil serine proteases affect TFPI’s anticoagulant function, we used neutrophil elastase to degrade it. Whereas intact TFPI strongly inhibited tissue factor acti-vity, TFPI proteolytically cleaved by neutrophil elastase was ineffec-tive (Supplementary Fig. 2c). Thus, degradation of TFPI impairs its anticoagulant activity, consistent with earlier results indicating that the anticoagulant efficacy of TFPI degradation products is decreased by about two orders of magnitude17. The inability of Elane−/−;Ctsg−/− neutrophils to efficiently cleave platelet-derived TFPI could explain their low procoagulant activity. The addition of antibody to TFPI substantially increased procoagulant activity in suspensions of Elane−/−;Ctsg−/−neutrophils and WT platelets to levels found using WT neutrophils and WT platelets (by 4.4 ± 0.2-fold versus control antibody; Fig. 2a). The effect of the TFPI-specific antibody was sig-nificantly attenuated when added to suspensions of WT neutrophils and WT platelets (2.0 ± 0.2-fold elevation versus control antibody, n = 3, P < 0.05 versus the increase seen with Elane−/−;Ctsg−/− neutro-phils and WT platelets). This difference in the effectiveness of the TFPI-specific antibody indicates that TFPI activity is less relevant in the case of combining WT neutrophils with WT platelets than in the case of combining Elane−/−;Ctsg−/−neutrophils with WT platelets, consistent with the notion that TFPI degradation is reduced when neutrophils lack serine proteases.
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Figure 2 TFPI degradation supports coagulation during thrombus development. (a) Factor Xa activity generated in suspensions of mouse neutrophils and platelets using coagulation factor concentrate. Neutrophils (N) and platelets (P) from WT or Elane−/−;Ctsg−/− (mut) mice were used, as indicated. Platelets were activated by collagen or ADP, as indicated. IgG, control antibody; anti-TFPI, antibody to mouse TFPI (50 μg ml−1). n = 3 or 4 per group, *P < 0.05 versus either N(WT) or P(WT), #P < 0.05 versus N(WT) + P(WT), §P < 0.05 versus N(mut) + P(mut), $P < 0.05 versus IgG control antibody. (b) Western blotting (left) and pull down experiments (right) for TFPI in plasma and platelets (Plat.) of uninjured mice. We used two sets of molecular weight (MW) markers. Representative of experiments on at least two mice per group. (c) Western blotting of TFPI in thrombi or heart from WT or Elane−/−;Ctsg−/− mice. Western blots, left; densitometry, right. The ratio between cleaved and intact TFPI was set as 100% in WT mice. n = 4 per group, *P < 0.05. (d) Factor Xa formation in suspensions of human neutrophils and platelets after addition of nTFPI and TFPI variants. Control, without TFPI. n = 5 per group, *P < 0.05 compared to control. (e) Factor Xa formation induced by recombinant tissue factor (left) or fibroblasts (right) after addition of nTFPI or the TFPI variant T87F L89A. Control, without TFPI. n = 4 per group, *P < 0.05 compared to control. (f) Fibrin deposition after injury of the carotid artery after infusion of nTFPI or T87F L89A (90 μg per kg body weight; ligation injury). n = 3 (WT) or 6 (Elane−/−;Ctsg−/−), *P < 0.05 compared to control. Error bars represent s.e.m. in all cases.
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We next investigated whether neutrophil serine proteases degrade TFPI during arterial thrombosis in vivo. Three isoforms of TFPI exist in mice (α, β and γ), with TFPI-β being the pre-dominant form33. TFPI-β lacks the Kunitz-3 domain; as a result, the heparin-releasable pool of TFPI in mice is small34. The three isoforms are indistinguishable by western blotting owing to their differential glycosylation pattern because the different lengths of the sugar chains obscure the differences in the molecular weights of the proteins. We found the main bands for TFPI in plasma, plate-lets, thrombi and heart to run at a molecular weight of 38–40 kDa (Fig. 2b,c). Pull-down experiments using microbead-coupled factor Xa (Fig. 2b) confirmed that the bands represented TFPI. Thrombus-associated TFPI was markedly degraded in WT mice (Fig. 2c). In contrast, TFPI cleavage was substantially attenuated in thrombi of Elane−/−;Ctsg−/− mice; in fact, in the absence of these serine proteases, the vast majority of TFPI persisted in its full-length form (Fig. 2c). The degradation product at 10–12 kDa observed in thrombi of WT mice, which is likely generated by neutrophil serine proteases because its abundance is reduced in Elane−/−;Ctsg−/− mice, probably represents the smaller degradation product induced by in vitro cleavage of TFPI (consisting mainly of the Kunitz-1 domain (Supplementary Fig. 2c). The expected larger cleavage product (consisting mainly of the Kunitz-2 and Kunitz-3 domains) is not detectable, indicating its subsequent degradation by additional proteases. Overall, these findings suggest that neutrophil serine proteases cleave TFPI within thrombi in vivo.
In contrast to thrombus-associated TFPI, the degradation status of plasma and platelet TFPI did not differ between WT and Elane−/−;Ctsg−/− mice, without or with vessel injury (Fig. 2b and Supplementary Fig. 3a), suggesting that TFPI degradation is restricted to the site of vascular injury and does not occur in the systemic circulation. Before injury, total TFPI concentrations in the plasma of WT and Elane−/−;Ctsg−/− mice were comparable (Supplementary Fig. 3b). Therefore, a difference in plasma TFPI concentrations does not account for the difference in the abundance of full-length TFPI in thrombi of the two types of mice. After vessel injury, plasma TFPI concentration was substantially reduced in WT mice (Supplementary Fig. 3b). Taken together, these results imply that TFPI, derived from the circulation and perhaps also from local sources such as endothelial cells and platelets, is recruited to the site of vessel injury and incorporated into thrombi, resulting in its degrada-tion by neutrophil serine proteases. In Elane−/−;Ctsg−/− mice, plasma TFPI concentrations were not substantially affected by vessel injury (Supplementary Fig. 3b), which may be due to the small thrombus size in these mice (Fig. 1d), possibly limiting TFPI incorporation into the growing thrombus.
Protease-resistant TFPI mutantsTo further substantiate the functional relevance of TFPI cleavage, we introduced mutations in human TFPI to increase its resistance to proteolysis. Alteration of the neutrophil elastase–specific cleavage site by substituting Thr87 with phenylalanine (T87F) prevented neutrophil elastase–induced TFPI degradation (Supplementary Fig. 3c). Additional substitution of Leu89 with alanine protected TFPI from cleavage by both neutrophil elastase and cathepsin G (Supplementary Fig. 3c). We also introduced an additional mutation at the cleavage site for MMPs (L21A) that prevented TFPI degradation by MMP-9 (Supplementary Fig. 3c). We then examined whether these TFPI variants affected coagulant activity induced by combining human neutrophils and platelets in vitro. All variants—that is, T87F, T87F
L89A and T87F L21A, but not native TFPI (nTFPI)—substantially inhibited the procoagulant activity of human neutrophils in the pres-ence of platelets (Fig. 2d). Although the neutrophil elastase– and cathepsin G–resistant T87F L89A variant was the strongest inhibitor, it was only slightly more effective than the T87F variant, suggest-ing that cathepsin G has a limited role in this system. To exclude the possibility that the intrinsic anticoagulant activity of the T87F L89A variant is affected, we analyzed its anticoagulant effects in the absence of neutrophil serine proteases. In such settings, T87F L89A inhibited procoagulant activity to an identical degree as nTFPI (Fig. 2e). Hence, the superior anticoagulant efficacy of T87F L89A is due to its enhanced resistance against proteolysis.
We next investigated the role of neutrophil-dependent TFPI clea-vage in triggering factor Xa generation by tissue factor–expressing THP-1 cells. The procoagulant activity of these THP-1 cells was strongly decreased by nTFPI in the absence of neutrophils, but not in their presence (Supplementary Fig. 4a,b). In contrast to nTFPI, the T87F-containing TFPI variants clearly decreased procoagulant activity in the presence of neutrophils (Supplementary Fig. 4b). Finally, we tested the effects of the TFPI variants on coagulation in human blood. Collagen-induced fibrin formation in whole blood was decreased by CMK (Supplementary Fig. 4c), indicating that this fibrin formation required neutrophil elastase. In this system, the T87F L89A variant but not nTFPI inhibited coagulation (Supplementary Fig. 4d). Collagen-triggered coagulation in whole blood involved tissue factor, as shown using a polyclonal antibody to tissue factor, which delayed coagulation by 59 ± 25 s compared to the coagulation time of 699 ± 46 s for isotype control IgG (mean values ± s.e.m., n = 7, P < 0.05; in line with the results in ref. 29). Coagulation in this system also involved factor XII, as shown using the factor XIIa inhibitor corn trypsin inhibitor (CTI), which delayed fibrin generation (Supplementary Fig. 4e). The protease-resistant TFPI variant T87F L89A thus inhibits coagula-tion despite the presence of neutrophil serine proteases in vitro.
We next infused nTFPI or the T87F L89A variant into mice (3 μg per mouse). This infusion increased total TFPI plasma concentration by 2.3-fold (Supplementary Fig. 5a). Control experiments confirmed that TFPI infused systemically was incorporated into the throm-bus (Supplementary Fig. 5b), again indicating that plasma TFPI is recruited to growing thrombi. Five minutes after ligation-induced vessel injury, T87F L89A inhibited coagulation with similar efficacy as nTFPI (Fig. 2f). This result indicates that the bioavailability of T87F L89A is comparable to that of nTFPI. However, the inhibitory effect of nTFPI gradually decreased thereafter and vanished after 30 min, whereas the anticoagulant effect of T87F L89A was sustained over 30 min, consistent with its superior protease resistance (Fig. 2f). In Elane−/−;Ctsg−/− mice, fibrin generation was not significantly lowered by infusion of either nTFPI or T87F L89A (Fig. 2f), indi-cating that the low amount of fibrin deposition in these mice is not further inhibited by exogenous TFPI, perhaps owing to the pres-ence of sufficient amounts of endogenous intact TFPI at the site of vessel injury. The superior anticoagulant efficiency of T87F L89A versus nTFPI was also evident after FeCl3-mediated vessel injury. As in the vessel ligation model, coagulation was significantly inhib-ited by T87F L89A but not by nTFPI 15 min after injury (P < 0.05; Supplementary Fig. 5c). Inhibition of coagulation by both TFPI species was less substantial after 5 min. Given that effects on fibrin formation in Elane−/−;Ctsg−/− mice and in T87F L89A–treated mice became clearly evident 10–15 min after injury, neutrophil serine proteases seem to predominantly regulate thrombus expansion as opposed to the initiation of thrombus formation.
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Procoagulant activity of externalized nucleosomesTo determine why serine protease–dependent degradation of TFPI is restricted to the thrombus compartment, we examined the cell surface sites supporting TFPI inactivation by neutrophils in vitro. TFPI colocalized with extracellular nucleosomes on the neutrophil surface (Fig. 3a). Neutrophil-derived externalized nucleosomes can form neutrophil extracellular traps (NETs) that capture pathogens35; however, the externalization of NETs is a slow process35,36. Given our previous observations indicating that neutrophils and platelets act in concert to promote coagulation (ref. 28 and Fig. 2a), we tested whether platelets could facilitate nucleosome release by neutrophils. Indeed, platelets activated by collagen or lipopolysaccharide substantially accelerated nucleo-some externalization by neutrophils (Supplementary Fig. 6a,b). Although the mechanisms underlying this effect are unclear, these results are in line with the ability of activated platelets to induce formation of NETs37. Externalized nucleosomes can be targeted with an antibody to the H2A-H2B–DNA complex35. This anti-body reduced TFPI binding to neutrophils (Fig. 3b). Cell surface TFPI colocalized with nucleosomes as well as with neutrophil elastase (Supplementary Fig. 6c). Disintegration of externalized nucleosomes with DNase abolished this colocalization of TFPI and neutrophil elastase (Supplementary Fig. 6c). Hence, externalized
nucleosomes enable the coassembly of neutrophil elastase and its substrate TFPI on the surface of activated neutrophils.
On the basis of these findings, we investigated the potential partici-pation of nucleosomes in neutrophil-induced coagulation. Purified reconstituted nucleosomes (NUCre) enhanced the procoagulant function of neutrophil elastase, as indicated by the ability of these nucleosomes to stimulate tissue factor activity in the presence of TFPI (Fig. 3c). In parallel, NUCre enhanced neutrophil elastase–dependent TFPI proteolysis (Fig. 3d,e). Nucleosomes prepared from cell nuclei ex vivo (NUCev) were similarly effective (Fig. 3d). Notably, the size of the major TFPI degradation product induced by neutrophil elastase in the presence of NUCre (Fig. 3e) corresponded to the TFPI frag-ment of 10–12 kDa found in vivo in thrombi of WT mice (Fig. 2c). After degradation of the NUCre with DNase, their stimulatory effect on neutrophil elastase–dependent TFPI proteolysis was attenuated (Fig. 3f). In addition, the H2A-H2B–DNA–specific antibody also attenuated TFPI degradation (Supplementary Fig. 6d).
To examine the role of nucleosomes in the procoagulant function of neutrophils, we treated neutrophils (previously incubated with acti-vated platelets to release their nucleosomes) with DNase to degrade exposed nucleosomes, removed the degraded nucleosomes, and then incubated the neutrophils with tissue factor–expressing THP-1 cells and TFPI. TFPI (2 nM) suppressed the tissue factor activity of these
Figure 3 Effect of externalized nucleosomes on TFPI function in vitro. (a) Visualization of TFPI and nucleosomes on the surface of activated neutrophils (human) with the use of TFPI–specific (green) and H2A-H2B–DNA–specific antibodies (red). Arrows, nucleosomes on neutrophil surface. Representative of three different experiments. Scale bar, 5 μm. (b) TFPI binding to the surface of activated neutrophils after addition of antibody to H2A-H2B–DNA or control antibody (control). Mean values of >150 cells, *P < 0.05. (c) Effect of nucleosomes on neutrophil elastase–dependent activity of recombinant tissue factor. Factor Xa generation by recombinant tissue factor was measured in the presence of TFPI (2 nM), neutrophil elastase (NE, 3 μg ml−1) or nucleosomes (reconstituted from isolated histones and isolated DNA obtained from Drosophila embryo nuclei; NUCre, 1 ng ml−1). n = 3–6 per group, *P < 0.05 versus control (no additions), #P < 0.05 versus TFPI plus NUCre,
§P < 0.05 versus TFPI plus neutrophil elastase. (d) Western blotting of isolated nTFPI (0.5 μg) using polyclonal TFPI–specific antibody after addition of neutrophil elastase (7 ng) with or without ex vivo–isolated nucleosomes (from nuclei of HeLa cells; NUCev, 5 pM) or NUCre (1 ng ml−1). Top, representative western blot, bottom, densitometry. Control, TFPI without additions (set as 100%), n = 3 per group. *P < 0.05 compared to control, #P < 0.05 compared to neutrophil elastase alone. (e) Western blotting of isolated nTFPI using TFPI Kunitz-1 domain–specific antibody after addition of neutrophil elastase with or without ex vivo–isolated nucleosomes (NUCre). (f) Western blotting of isolated nTFPI using TFPI Kunitz-1 domain–specific antibody after addition of neutrophil elastase with ex vivo–isolated nucleosomes (NUCre) in the presence of DNase I or CMK. Nucleosomes were pretreated with DNase I (10 U ml−1) or vehicle and then incubated with TFPI with or without neutrophil elastase and CMK (0.5 mM) under otherwise similar conditions to those used in d. (g) Effect of neutrophils on TFPI activity. Left, procoagulant activity of tissue factor–expressing THP-1 cells without (100%) or with TFPI (0.5 nM). Right, coagulant activity of neutrophils pretreated with DNase or clostripain and then incubated with THP-1 cells with or without TFPI. n = 3–6 per group, *P < 0.05 versus without additions. Error bars represent s.e.m. in all cases.
a
c
f g
b
d e
NE
NE
# §
–
* *
* *
** #
* #
* #
+ + +
+
+
+ ++
+
+
+ +
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+ +
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–
TFPI
MW (kDa)
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36
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NUCev
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))
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+ N
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THP-1 cells when neutrophil-derived nucleosomes had been previ-ously degraded and removed, but not in the presence of untreated neutrophils (Fig. 3g), further indicating that nucleosomes contribute to TFPI inactivation. In addition to promoting TFPI degradation, nucleosome-mediated recruitment of TFPI to neutrophils attenu-ated TFPI binding to THP-1 cells (Supplementary Fig. 6e). To iden-tify the molecular components of nucleosomes necessary for TFPI degradation, we selectively degraded the histone component of the nucleosomes with clostripain38. Clostripain treatment of neutrophils did not affect their ability to suppress the anticoagulant effects of TFPI (Fig. 3g), suggesting that histones are not needed for the ability of nucleosomes to degrade TFPI. Moreover, isolated DNA (used to reconstitute the nucleosomes in vitro), but not the isolated histone fractions, increased the procoagulant activity induced by neutrophil serine proteases (data not shown). These data suggest that the nucleic acid components of nucleosomes, acting through their polyanionic surfaces and probably independently of histones, support TFPI deg-radation and enhance coagulation. Because the assembly of several coagulation complexes requires anionic surfaces, additional mecha-nisms beyond the one described here—facilitation of the degradation of TFPI by neutrophil serine proteases—are likely to contribute to the procoagulant actions of externalized nucleosomes.
Nucleosomes enhance coagulation in vivo We next assessed the contribution of externalized nucleosomes to fibrin formation in mice. Examination of thrombi in FeCl3-injured vessel sections by confocal microscopy indicated that, in both WT and Elane−/−;Ctsg−/− mice, most thrombus-resident neutrophils were directly attached to or located in the vicinity of the damaged intima (Fig. 4a). In both types of mice, externalized nucleosomes could be detected on the surface of thrombus-resident neutrophils as well as adjacent to them (Fig. 4a). TFPI was also localized in the vicinity of
thrombus-associated neutrophils, but this association was decreased after injection of H2A-H2B–DNA–specific antibody (Supplementary Fig. 7), indicating that externalized nucleosomes contribute to the recruitment of TFPI to the neutrophil surface in vivo. In WT mice, administration of the H2A-H2B–DNA–specific antibody substantially suppressed coagulation at the site of vascular injury after both 15 and 25 min (Fig. 4b), consistent with the role of neutrophils in promoting thrombus growth. In contrast, fibrin formation in Elane−/−;Ctsg−/− mice was not significantly affected by H2A-H2B–DNA–specific anti-body administration (Fig. 4b). This result is consistent with the view that externalized nucleosomes predominantly support coagulation through effects on neutrophil serine proteases. Although the H2A-H2B–DNA–specific antibody did not alter platelet adhesion to the injured vessel site (Fig. 4c), it clearly delayed occlusion of the carotid artery in WT mice (Fig. 4d; in line with previous results39). Moreover, whereas the antibody also decreased the time to reestablishment of flow after initial occlusion in WT mice, indicating decreased thrombus stability (Fig. 4d), it did not affect vessel occlusion and the time to rees-tablishment of flow in Elane−/−;Ctsg−/− mice (Fig. 4d). Nucleosomes thus support coagulation and thrombus formation by compartmental-izing thrombus-associated TFPI and neutrophil-derived serine pro-teases, facilitating TFPI degradation at sites of vessel injury in vivo.
Neutrophil-dependent coagulation during systemic infectionWe next explored whether coagulation induced by neutrophil serine proteases might be an evolutionarily conserved strategy that sup-ports innate immunity responses to pathogen invaders. To address this, we infected mice systemically with Escherichia coli (E. coli). In WT mice, E. coli infection triggered fibrin deposition inside ves-sels of the hepatic microvasculature (after 6 h), resulting in partial or even complete occlusion by fibrin clots (Fig. 5a). We found that 94 ± 4% of the vessels occluded by fibrin were sinusoids (mean values
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Figure 4 Externalized nucleosomes promote fibrin formation in vivo. (a) Visualization of extracellular nucleosomes, neutrophils and DNA in thrombi after FeCl3-induced vessel injury (20 min) using antibody to H2A-H2B–DNA (red), myeloperoxidase (MPO)-specific antibody (green) and Hoechst 33342 (blue). Arrows indicate nucleosomes on the surface and in the vicinity of thrombus-associated neutrophils. Differential interference contrast images (left) show thrombus segments and adjacent vessel walls. Boxes indicate thrombus segments labeled. Scale bar, 25 μm. Representative of at least three experiments per group. (b) Effect of H2A-H2B–DNA–specific antibody on fibrin generation in WT mice. After injection of H2A-H2B–DNA–specific antibody (or control IgG), fibrin formation at the injury site in the carotid artery was determined 15 and 25 min after injury with FeCl3. n = 4 per group, *P < 0.05 versus value obtained at same time point after infusion of isotype IgG (set as 100%). (c) Effect of H2A-H2B–DNA–specific antibody on platelet deposition. Firm platelet adhesion was determined by measuring the attachment of 2′,7′-dichlorofluorescein (DCF)-labeled platelets to the site of vessel injury in WT mice. n = 4 per group. (d) Effect of H2A-H2B–DNA–specific antibody on vessel occlusion. Time to vessel occlusion (left) and duration of occlusion (right) were determined after FeCl3 injury in the absence or presence of H2A-2B–DNA–specific antibody in WT and Elane−/−;Ctsg−/− mice. n = 3–6 per group, *P < 0.05. Error bars represent s.e.m. in all cases.
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± s.e.m., n = 133), as indicated by staining for stabilin-2, a marker for sinusoidal endothelial cells40. Intravascular fibrin colocalized with aggregates of activated neutrophils (Supplementary Fig. 8a), known to lodge primarily in liver microvessels during systemic infection41, and also in part with Kupffer cells (Supplementary Fig. 8b). We also frequently detected GFP-labeled E. coli in association with the fibrin clots (Supplementary Fig. 8c). In most cases, the bacteria accumu-lated in the periphery of the clots.
Fibrin deposition in liver sinusoids of Elane−/−;Ctsg−/− mice infected systemically with E. coli was markedly reduced compared to WT mice (Fig. 5a,b). In parallel, the number of microvessels occluded by fibrin was decreased (Fig. 5b). Despite this, the total number of neutrophils in microvessels of Elane−/−;Ctsg−/− mice was similar to that in WT mice (Fig. 5c; in agreement with previous results10,11); likewise, blood concentrations of the inflammatory mediators tumor necrosis factor-α (TNF-α) and monocyte chemotactic protein-1 (MCP-1) were unaltered (Fig. 5d), excluding the possibility that the reduced fibrin accumulation in Elane−/−;Ctsg−/− mice was due to changes in neutrophil accumulation or in the inflammatory response. Intravascular E. coli colocalized with Kupffer cells in both types of mice (Supplementary Fig. 8d), in line with the ability of these cells to bind and phagocytose bacteria and suggesting that Kupffer cells bind bacteria in both strains of mice. Tissue factor and factor XII both contribute to activation of coagulation during systemic infec-tion, as indicated by the ability of both mouse tissue factor-specific antibody and the contact pathway inhibitor PCK (both injected before infection) to suppress fibrin deposition in liver microvessels in WT mice (Fig. 5e). As in our previous observations in a model of arterial thrombosis, externalized nucleosomes were important for micro-vascular fibrin generation in E. coli–infected WT mice, as infusion of H2A-H2B–DNA–specific antibody decreased fibrin deposition and reduced the number of occluded microvessels (Fig. 5f). Hence, similar to their effects on large-vessel thrombosis, neutrophil serine proteases and externalized nucleosomes crucially support intravascular coagu-lation during systemic E. coli infection.
Neutrophil-driven coagulation prevents extravasation of E. coliTo address whether coagulation driven by neutrophil serine proteases and externalized nucleosomes might control bacterial spreading, we assessed bacterial distribution inside and outside the hepatic micro-vasculature. In WT mice, we detected E. coli mostly in hepatic blood vessels with diameters <30 μm that stained positive for stabilin-2; E. coli was essentially absent from larger vessels or from perivascular hepatic tissue (Fig. 6a,b). Significantly more E. coli were present in the perivascular tissue parenchyma in Elane−/−;Ctsg−/− mice compared to WT mice, and the tissue-to-vessel ratio of bacterial distribution was strongly increased in Elane−/−;Ctsg−/− mice (Fig. 6a,b). This increase in bacterial accumulation in the perivascular tissue of Elane−/−;Ctsg−/− mice was not associated with manifest liver damage, as serum con-centrations of aspartate aminotransferase and alanine aminotrans-ferase were barely elevated (Supplementary Fig. 9a). Nonetheless, systemic blood counts of E. coli were low in both strains of mice (<400 colony-forming units (CFU) ml−1 in blood). We also evaluated the role of serine protease–triggered coagulation in bacterial spreading in other tissues. CFU measurements indicated that the numbers of E. coli were higher in lung, liver and spleen (the preferential tissue localization sites of E. coli at this time point42) of Elane−/−;Ctsg−/− compared to those in WT mice10,11 (Supplementary Fig. 9b). In addi-tion to the reduced fibrin deposition in the liver microcirculation of Elane−/−;Ctsg−/− mice after systemic E. coli infection, fibrin deposition was also reduced in the splenic microcirculation (Supplementary Fig. 9c). This reduction was accompanied by enhanced pathogen accumulation in the perivascular tissue of the spleen (Supplementary Fig. 9c) indicating that changes in fibrin deposition and bacterial distribution in Elane−/−;Ctsg−/− mice are not restricted to the liver.
We next tested whether decreased intraluminal fibrin deposi-tion was responsible for the increased bacterial spreading in the absence of neutrophil serine proteases. Treatment of WT mice with the thrombin inhibitor hirudin markedly reduced fibrin deposition and the number of occluded vessels (Fig. 6c). This treatment also impaired intravascular entrapment of E. coli, resulting in a substantial
Figure 5 Effect of neutrophil serine proteases on fibrin deposition during systemic infection. (a) Fibrin deposition in liver sinusoids after systemic infection. Analyses were performed 6 h after infection of WT mice with E. coli. Fibrin and liver sinusoids were visualized with fibrin-specific (red) and stabilin-2–specific antibodies (green; arrowheads). Arrows, hepatocyte nuclei. Insets show images obtained with similarly labeled respective control antibodies. Scale bars, 10 μm; insets, 20 μm. Representative images of four mice per group. (b) Quantitative analysis of fibrin deposition (left) and fibrin-occluded microvessels (right) in liver sinusoids after systemic infection. Analysis of >1,200 microvessels in four mice per group, *P < 0.05. (c) Total numbers of intravascular neutrophils after systemic infection with E. coli. n = 4 or 5 per group. (d) Quantification of TNF-α and MCP-1 in plasma obtained from systemic blood of WT and Elane−/−;Ctsg−/− mice 6 h after E. coli infection. n = 4 per group. (e) Fibrin deposition in liver microvessels was measured in WT mice which had been injected with mouse tissue factor–specific antibody or the contact pathway inhibitor PCK immediately before injection of bacteria. n = 3–5 per group. *P < 0.05 versus control (IgG and vehicle for treatment with tissue factor–specific antibody and PCK, respectively); values for the controls were set as 100%. (f) Effect of nucleosomes on coagulation during systemic infection. Fibrin deposition (left) and fibrin-occluded microvessels (right) in WT mice were measured after infusion of H2A-H2B–DNA–specific antibody or of isotype IgG (control). Analysis of >900 vessels per group. n = 3 or 4 per group, *P < 0.05. Error bars represent s.e.m. in all cases.
Anti–stabilin-2 Anti-fibrin Merge
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*
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Fib
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)M
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increase in tissue invasion and in the tissue-to-vessel ratio of bacterial distribution (Fig. 6d). Conversely, stimulation of coagulation by injec-tion of recombinant factor VIIa (rVIIa) led to a significant increase in fibrin deposition and in the number of occluded vessels (Fig. 6c), and did not affect bacterial distribution (Fig. 6d). Hirudin injection into Elane−/−;Ctsg−/− mice slightly diminished fibrin deposition and vessel occlusion and increased pathogen invasion into the tissue (Fig. 6e). Injection of rVIIa into these mice enhanced fibrin deposition and the number of occluded vessels (Fig. 6d), reaching the level of coagulation activation observed in WT mice. The tissue-to-vessel ratio of E. coli in these mice was also similar to the level seen in WT mice (Fig. 6e versus Fig. 6b), suggesting that the defect in intravascular bacterial trapping in Elane−/−;Ctsg−/− mice can be compensated for by activa-tion of coagulation.
We next investigated whether, similar to our observations in the case of arterial thrombosis, TFPI proteolysis and externalized nucleosomes participate in the activation of coagulation in liver
microvessels after systemic E. coli infection. Administration of either nTFPI or the T87F L89A variant immediately before injection of E. coli reduced fibrin deposition in liver microvessels; however, the decrease induced by T87F L89A was substantially greater than that induced by nTFPI (Fig. 6f). In parallel, both TFPI species promoted tissue invasion of E. coli; however, the effect of T87F L89A was more pronounced, as indicated by an increase in the E. coli vessel-to-tissue ratio (Fig. 6f). Moreover, infusion of H2A-H2B–DNA–specific anti-body, which suppresses fibrin formation (Fig. 5f), strongly shifted bacterial localization toward the tissue (Supplementary Fig. 9d). These results indicate that neutrophil serine protease–induced TFPI cleavage not only supports coagulation after vascular injury but also during systemic infection, contributing to the retention of bacteria inside microvessels.
To test whether neutrophil serine protease–dependent TFPI cleav-age affects the viability of E. coli, we determined overall bacterial counts in liver and spleen after injection of either nTFPI or T87F
WTWT
Elane–/–;Ctsg–/–
Elane–/–;Ctsg–/–
Elane–/–;Ctsg–/–
Elane–/–;Ctsg–/–
Elane–/–;Ctsg–/–
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3
0
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Co rVllaHir
CoCoCo Co
Co
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CoCo CoCo Co
nTFPI
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nTFPInTFPI nTFPInTFPI nTFPI
T87FL89A
T87FL89A
T87FL89A
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T87FL89A
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CorVllarVllarVlla rVllaHirHirHir
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Figure 6 Effect of fibrin formation on bacterial tissue invasion. (a) Localization of GFP-labeled E. coli (pseudocolored blue) in liver sinusoids (detected with labeled stabilin-2–specific antibody, green) 6 h after injection into WT and Elane−/−;Ctsg−/− mice. Arrowheads, positions of bacteria. Scale bars, 5 μm. Representative images of four mice per group. (b) Bacterial distribution in the microcirculation of the liver. Left, bacterial numbers in the microvessel lumen and surrounding tissue. >2,000 vessels per group, n = 4 per group. *P < 0.05 versus WT vessel, #P < 0.05 versus Elane−/−;Ctsg−/− vessel, §P < 0.05 versus WT tissue. Right, same data as in the left graph, plotted as tissue-to-vessel ratio. Broken line, threshold above which the number of bacteria in tissue exceeds that in the vessel. *P < 0.05. (c) Fibrin deposition and vessel occlusion in liver sinusoids 6 h after infection of WT mice with E. coli in the presence of hirudin and rVIIa. >1,100 vessels per group, n = 4 per group. Co, control (without hirudin or rVIIa). Hir, hirudin. *P < 0.05 versus control. (d) Left, bacterial distribution in the microcirculation of the liver in WT mice in the presence of hirudin and rVIIa. >1,200 vessels per group. Right, same data as in the left graph, plotted as tissue-to-vessel ratio. n = 4 per group, *P < 0.05 versus vessel (same condition), #P < 0.05 versus tissue (control or rVIIa addition), §P < 0.05 versus control or rVIIa. (e) Bacterial distribution in the microcirculation of the liver in Elane−/−; Ctsg−/− mice in the presence of hirudin and rVIIa. Control values are the same values as in Figure 5b and panel b. >1,100 vessels per group, n = 4 per group, *P < 0.05 versus control and versus hirudin (left, middle) or rVIIa (right). (f) Fibrin deposition and bacterial distribution in the microcirculation of the liver in WT mice after infusion of nTFPI or T87F L89A. >1,000 vessels per group, n = 4 per group, *P < 0.05 versus control, #P < 0.05 versus nTFPI. Control, absence of TFPI. (g) Bacterial CFUs in liver and spleen 6 h after systemic infection of WT mice with E. coli in the presence of nTFPI or T87F L89A. n = 4 mice per group. *P < 0.05 versus control, #P < 0.05 versus nTFPI. Control, absence of TFPI. (h) Fibrin deposition (left) and bacterial distribution in liver microcirculation (middle left), as well as bacterial CFUs in liver (middle right) and in spleen (right) 6 h after systemic infection of Elane−/−;Ctsg−/− mice with E. coli in the presence of nTFPI or T87F L89A. Control values in left, middle left and middle right graphs are the same values as in Figure 5b and panel b. n = 3 mice per group. *P < 0.05 versus control. Error bars represent s.e.m. in all cases.
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L89A. Both TFPI species augmented CFU counts in liver and spleen of WT mice, but bacterial survival was substantially higher after injec-tion of T87F L89A compared to nTFPI (Fig. 6g). In Elane−/−;Ctsg−/− mice, in which degradation of endogenous TFPI is largely abrogated, injection of either of the TFPI species only slightly decreased fibrin deposition (Fig. 6h). This was accompanied by a further shift in the bacterial distribution toward the extravascular compartment of the liver and increased bacterial counts in liver and spleen (Fig. 6h). The observation that nTFPI and T87F L89A had similar effects in Elane−/−;Ctsg−/− mice (Fig. 6h) suggests that the differential effects of nTFPI and T87F L89A in WT mice on fibrin deposition and bacterial survival (Fig. 6f,g) are due to the increased resistance of the T87F L89A variant to neutrophil serine protease–mediated degrada-tion. Hence, activation of coagulation induced by neutrophil serine proteases helps restrict bacterial survival during systemic infection. Overall, these findings suggest that intravascular coagulation supports pathogen retention in liver microvessels and suppresses bacterial tissue invasion, most likely at the expense of tissue perfusion.
DISCUSSIONWe report here that neutrophil serine proteases, major components of the mammalian antimicrobial machinery, not only kill micro-organisms but also trigger blood coagulation. This stabilizes nas-cent thrombi and helps reduce blood loss after vessel injury. During systemic infection, neutrophil serine protease–driven coagulation supports immobilization of bacteria within the microvasculature, thereby attenuating the spreading of pathogens into the extravascular tissue. These findings suggest that thrombosis represents a physio-logical tool of innate antimicrobial defense. Hence, the ability of coagulation to support host defense, which is readily apparent in arthropods1, is conserved in mammals. Activation of intravascular coagulation probably constitutes a major strategy of the general intra-vascular host defense system43. Neutrophil serine proteases thus con-tribute to establishing a bidirectional communication between the immune response and blood coagulation, enhancing the efficacy of two major host protection systems, hemostasis and innate immunity (Supplementary Fig. 10).
We identified externalized nucleosomes as a platform for intravas-cular fibrin generation. Externalized nucleosomes capture not only pathogens35 but also substrates of neutrophil serine proteases such as TFPI (Supplementary Fig. 10), which originate from multiple sources including activated or disrupted endothelial cells, activated platelets and plasma. Externalized nucleosomes, which are known to be released from activated neutrophils but could additionally also be liberated from damaged vessel wall cells at a site of injury, facilitate coassembly of neutrophil elastase and the anticoagulant TFPI, sup-porting TFPI inactivation and unleashing suppression of factor Xa, thereby fostering fibrin formation. By inactivating TFPI, neutrophil serine proteases primarily affect the extrinsic pathway of coagula-tion initiated by tissue factor originating from vessel wall cells and from intravascular sources28. Apart from tissue factor, activation of coagulation also requires the factor XII–induced contact path-way under various conditions of thrombus formation. The partial overlap of the inhibition profiles seen here with tissue factor and factor XII blockade in vivo suggests that the extrinsic and contact pathways might be connected in a sequential manner. Indeed, plate-lets activated by thrombin previously generated via the extrinsic pathway might activate factor XII; TFPI might thus act upstream of contact pathway activation. Moreover, neutrophil serine proteases and externalized nucleosomes might directly stimulate the contact
pathway; for example, by inactivation of its antagonist C1 inhibitor44 or via activation of factor XII45. The latter possibility is particularly attractive, as factor XII activation is promoted by nucleic acids46.
The reduced TFPI cleavage observed in neutrophil serine protease–deficient mice is likely to contribute to the prolonged bleeding times observed here in these mice. Because bleeding times are only moderately prolonged in “low tissue factor” mice47 and are unaffected by factor XII deficiency24, the ability of TFPI to inhibit factor IXa generation48 might, apart from its ability to inhibit factor Xa generation, contribute to the enhanced bleeding in neutrophil serine protease-deficient mice. The key role of neutrophil elastase for fibrin formation in vivo may be unexpected considering the broad substrate affinity of neutrophil serine proteases. However, the procoagulant function of these proteases depends on a temporally defined establishment of a nucleosome scaffold to entrap its sub-strates. In addition, the rapid extrusion of the nucleosomes from neutrophils requires interactions with activated platelets that are specifically enabled during intraluminal thrombus development. Hence, rather specific spatiotemporal conditions are required to sup-port the procoagulant activity of neutrophil elastase, consistent with the proposal that individual functions of multisubstrate enzymes are specified by a distinct spatiotemporal context49.
Our findings show that neutrophil serine proteases, which activate coagulation during systemic infection, also trigger occlusion of large vessels in the absence of a pathogen challenge, resulting in arterial thrombosis, a leading cause of death worldwide50. The procoagulant activity of these proteases primarily reflects their ability to stabilize intraluminal thrombus development by counteracting endogenous anticoagulants that impede intraluminal coagulation under physio-logical conditions. Neutrophil-mediated prothrombotic mechanisms may therefore contribute to pathology in cardiovascular diseases such as myocardial infarction, as well as in infectious disease and other diseases7–10,51. Consequently, the use of protease-resistant TFPI mutants and suppression of coagulation as induced by extra-cellular nucleosomes provide a rationale for the development of new therapies targeting disease-specific thrombotic complications.
METHODSMethods and any associated references are available in the online version of the paper at http://www.nature.com/naturemedicine/.
Note: Supplementary information is available on the Nature Medicine website.
ACKnowLEDGMEnTSWe acknowledge support of Deutsche Forschungsgemeinschaft and Wilhelm Sander-Stiftung to B.E. and S.M. We are grateful to W. Bode, J. Roes, Peter Lohse, Pia Lohse and A. Moseman for helpful suggestions and comments. We thank G. Längst (University of Regensburg) and K. Rippe (University of Heidelberg) for preparation of isolated nucleosomes, M. Monestier (Temple University), H. Wardemann and R. Hurwitz (Max-Planck-Institut für Infektionsbiologie) for providing H2A-H2B–DNA–specific antibody, DNA-specific antibody and human neutrophil elastase–specific antibody, K. Schledzewski (University of Heidelberg) for supplying the stabilin-2–positive antibody, L. Petersen (NovoNordisk) for donating rVIIa and D. Kirchhofer (Genentech) for providing mouse tissue factor–specific antibody. L.G. was supported by Deutsche Forschungsgemeinschaft-Forschergruppe 440 and by Rudolf-Marx-Stipendium from Gesellschaft für Thrombose-und Hämostaseforschung. K.B. and C.R. were participants of Deutsche Forschungsgemeinschaft-Graduiertenkolleg 438.
AUTHoR ConTRIBUTIonSS.M. (together with B.E.) designed and supervised the study, analyzed data and contributed to writing the manuscript, L.G. conducted in vitro and in vivo experiments and analyzed data, M.-L.v.B. conducted in vivo experiments and analyzed data, D.M. conducted in vitro experiments and analyzed data, S.P. performed histochemical analyses, C.G. and V.B. performed morphological
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analyses, M.L. performed protein analyses, K.B. prepared TFPI mutants, A.B.K. performed protein analyses, I.K. and E.K. conducted part of the in vivo experiments, K.R., S.H. and S.B. performed analyses on in vitro and in vivo experiments, C.R. performed protein analyses, M.S. and K.T.P. contributed to design the study and B.E. designed the study, supervised the work, analyzed data and wrote the manuscript.
CoMPETInG FInAnCIAL InTERESTSThe authors declare no competing financial interests.
Published online at http://www.nature.com/naturemedicine/. Reprints and permissions information is available online at http://npg.nature.com/reprintsandpermissions/.
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ONLINE METHODSCells and mice. We prepared human platelets as described28 and isolated human neutrophils using microbeads28 or Ficoll gradients. To remove red blood cells, suspensions of isolated neutrophils were then applied to a Histopaque gradient or to hypotonic lysis. We obtained informed consent from all blood donors; the study was approved by the Ethics Committee of the Medical Faculty of Ludwig-Maximilians-University. We isolated mouse platelets as previously described26 and prepared mouse neutrophils from bone marrow of WT and Elane−/−;Ctsg−/− mice by a Percoll density gradient as previously described52. We cultivated human skin fibroblasts (PromoCell, cat. no. C-12352) and THP-1 cells (Abnova, cat. no. L007V2) in DMEM plus 10% FBS containing 1% of a penicillin-streptomycin suspension. Elane−/−;Ctsg−/− and Elane−/−;Ctsg+/+ mice were generated as previously described12. WT mice were of the same strain background (SV129S1, Charles River). This mouse strain has lower platelet counts compared to C57BL/6 mice53. Mice from the various groups were age- and sex-matched, and their carotid artery dia-meters and vessel wall thicknesses were comparable. The experiments were performed in accordance with the Animal Facility guidelines of the Medical Faculty of Technical University Munich. All procedures performed on mice were approved by the local legislation on protection of animals (Regierung von Oberbayern, Munich).
Preparation of nucleosomes. Preparation of NUCev from nuclei of HeLa cells was performed as previously described54. Histones and DNA were isolated sepa-rately from Drosophila embryo nuclei and then reconstituted to nucleosomes (NUCre) as previously described55.
TFPI mutants. We performed site-directed mutagenesis of human TFPI (isoform TFPI-α) with the GENE Taylor kit (Invitrogen). The proteins were expressed in E. coli and isolated from bacterial supernatants.
Confocal microscopy. We exposed thrombus sections excised from mouse carotid arteries after injury with FeCl3 to H2A-H2B–DNA–specific anti-body56. In order to label the H2A-H2B–DNA–specific antibody, it was directly coupled to the fluorophore Atto 550 (Atto-Tec). On identical sec-tions, neutrophils were detected using an MPO-specific antibody (DAKO, cat. no. A0398) which was visualized with secondary donkey antibody to rabbit IgG coupled to Cy5 (Jackson ImmunoResearch, cat. no. 711-175-152). We stained DNA with bis-benzimide 33342 (Hoechst 33342) or with human monoclonal DNA-specific antibody detected with Cy2-labeled don-key antibody to human IgG (Jackson ImmunoResearch, cat. no. 702-225-149). We analyzed the specimens with a confocal microscope (SP5, Leica) or with a Leica DMR Microscope equipped with a Nuance Multispectral Imaging system.
Western blots and TFPI pull-down. We analyzed TFPI degradation in vitro in supernatants of activated human platelets and neutrophils. We performed western blotting with a polyclonal goat antibody to human TFPI (Santa Cruz, cat. no. sc18713), or, where indicated, a monoclonal TFPI Kunitz-1 domain–specific antibody (American Diagnostica, cat. no. 4903). To determine TFPI degradation in vivo, we analyzed thrombus sections excised from mouse carotid arteries after injury with FeCl3 using a rabbit antibody to mouse TFPI (American Diagnostica, cat. no. 4913). For pull-down of TFPI, we precipitated the anticoagulant from mouse tissues with bovine factor Xa (NEB, cat. no. P8010L) coupled to beads (Roti-MagBeads COOH, Roth, cat. no. HP58.1).
Coagulation assays. We determined procoagulant activities in the presence of factor VIIa (Novo Nordisk, NovoSeven) and factor X23 (Calbiochem, Merck 233282) or using a coagulation factor concentrate28 (Aventis Pharma, Beriplex P/N 500, cat. no. 6975286). Whole-blood coagulation was measured by throm-belastography (roTEG, Dynabyte)28.
In vivo thrombosis and fibrin formation. We exposed the carotid arteries of anaesthetized mice to FeCl3 or subjected them to ligation as previously described26. We visualized fibrin formation by high-speed intravital microscopy23 with an Alexa488-labeled antibody to the fibrin IIβ chain (Accurate Chemical, cat. no. NYBT2G1). After FeCl3 exposure, the time interval was registered until complete arrest of blood flow in the center of the vessel. We considered the sub-sequent time interval until reappearance of flow as the duration of occlusion.
Tail bleeding. We cut tails of isoflurane-anesthetized mice 5–6 mm from the tip with a sterile scalpel and placed the remaining tail on a filter paper to measure the time until bleeding stopped.
Systemic infection with Escherichia coli. We injected GFP-labeled E. coli57 (3.2 × 108 CFU) into the jugular vein of WT and Elane−/−;Ctsg−/− mice. After 6 h, we isolated livers and spleens and prepared sections to visualize fibrin, bacteria, Kupffer cells and neutrophils. Liver and spleen sinusoids were detected by stain-ing with Alexa546-labeled antibody to mouse stabilin-2 (ref. 58). We counted the number of GFP-positive E. coli in the lumen and in the perivascular tissue of stabilin-2–positive vessels. Neutrophil numbers were determined in liver sec-tions after staining with H&E. We counted CFUs in blood, liver, spleen, lung and kidneys and measured plasma levels of MCP-1 and TNF-α by ELISA (R&D Systems, MJE00 and MTA00).
Statistical analyses. All data shown are means ± s.e.m. The results were analyzed using the Mann-Whitney rank-sum test and one-way analysis of variance fol-lowed by the Student-Newman-Keuls test where appropriate. We considered P < 0.05 as statistically significant.
Additional methods. Detailed methodology is described in the Supplementary Methods.
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56. Losman, M.J., Fasy, T.M., Novick, K.E. & Monestier, M. Monoclonal autoantibodies to subnucleosomes from a MRL/Mp(-)+/+ mouse. Oligoclonality of the antibody response and recognition of a determinant composed of histones H2A, H2B, and DNA. J. Immunol. 148, 1561–1569 (1992).
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58. Schledzewski, K. et al. Lymphatic endothelium-specific hyaluronan receptor LYVE-1 is expressed by stabilin-1+, F4/80+, CD11b+ macrophages in malignant tumours and wound healing tissue in vivo and in bone marrow cultures in vitro: implications for the assessment of lymphangiogenesis. J. Pathol. 209, 67–77 (2006).
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