Activated protein C inhibits neutrophil extracellular trapformation in vitro and activation in vivoReceived for publication, November 21, 2016, and in revised form, April 11, 2017 Published, Papers in Press, April 13, 2017, DOI 10.1074/jbc.M116.768309

X Laura D. Healy‡1, Cristina Puy§, José A. Fernández¶, Annachiara Mitrugno§, Ravi S. Keshari�, Nyiawung A. Taku§,Tiffany T. Chu§, Xiao Xu¶, András Gruber§, Florea Lupu�, John H. Griffin¶, and Owen J. T. McCarty‡§2

From the Departments of ‡Cell, Developmental & Cancer Biology and §Biomedical Engineering, Oregon Health & ScienceUniversity, Portland, Oregon 97230, the ¶Department of Molecular Medicine, The Scripps Research Institute, La Jolla, California92037, and the �Cardiovascular Biology Research Program, Oklahoma Medical Research Foundation,Oklahoma City, Oklahoma 73104

Edited by John M. Denu

Activated protein C (APC) is a multifunctional serine prote-ase with anticoagulant, cytoprotective, and anti-inflammatoryactivities. In addition to the cytoprotective effects of APC onendothelial cells, podocytes, and neurons, APC cleaves anddetoxifies extracellular histones, a major component of neutro-phil extracellular traps (NETs). NETs promote pathogen clear-ance but also can lead to thrombosis; the pathways that nega-tively regulate NETosis are largely unknown. Thus, we studiedwhether APC is capable of directly inhibiting NETosis via recep-tor-mediated cell signaling mechanisms. Here, by quantifyingextracellular DNA or myeloperoxidase, we demonstratethat APC binds human leukocytes and prevents activatedplatelet supernatant or phorbol 12-myristate 13-acetate (PMA)from inducing NETosis. Of note, APC proteolytic activitywas required for inhibiting NETosis. Moreover, antibodiesagainst the neutrophil receptors endothelial protein C re-ceptor (EPCR), protease-activated receptor 3 (PAR3), andmacrophage-1 antigen (Mac-1) blocked APC inhibition ofNETosis. Select mutations in the Gla and protease domains ofrecombinant APC caused a loss of NETosis. Interestingly,pretreatment of neutrophils with APC prior to induction ofNETosis inhibited platelet adhesion to NETs. Lastly, in a non-human primate model of Escherichia coli-induced sepsis,pretreatment of animals with APC abrogated release ofmyeloperoxidase from neutrophils, a marker of neutrophilactivation. These findings suggest that the anti-inflammatoryfunction of APC at therapeutic concentrations may includethe inhibition of NETosis in an EPCR-, PAR3-, and Mac-1-de-pendent manner, providing additional mechanistic insightinto the diverse functions of neutrophils and APC in diseasestates including sepsis.

Activated protein C (APC)3 is a 56-kDa serine protease thatacts as an anticoagulant and anti-inflammatory molecule. Incomplex with protein S, APC inactivates coagulation factors (F)Va and VIIIa, dampening the generation of the thrombin (1– 4).APC is generated from the zymogen protein C via enzymaticcleavage by thrombin. On endothelial cells this reaction isenhanced in a calcium-dependent manner in the presence ofthrombomodulin and endothelial protein C receptor (EPCR)(5– 8). Because of the combined antithrombotic and anti-in-flammatory properties of APC, recombinant APC was usedclinically as a therapeutic for adult severe sepsis before itsremoval in 2011, as no overall benefit to patient outcome wasobserved in a second phase 3 trial performed 10 years after thefirst successful phase 3 trial (9, 10). In severe sepsis therapy,APC increased risk for serious bleeding (9). Thus, a compre-hensive understanding of the structure-activity relationshipsfor the various functional activities of APC may provide therationale for developing different engineered APC mutants thatmaintain the anti-inflammatory or cytoprotective functions ofAPC without adversely affecting thrombin generation or com-promising hemostasis (11, 12).

Neutrophils are the most populous innate immune cellsfound in circulation, playing an indispensable role in surveyingthe body for signs of infection and inflammation (13). Neutro-phils express several of the key receptors responsible for bind-ing and mediating cell signaling of APC, and APC can inhibitneutrophil migration (14 –18).

Classically, the cytoprotective mechanisms of APC havebeen best characterized on the endothelium, where APCcanonically binds EPCR and cleaves protease activated receptor(PAR) 1 at Arg-46, a unique site different from thrombin’scleavage site at Arg-41, activating an intracellular signaling cas-cade to promote barrier function and inhibit apoptosis, a con-cept known as biased agonism (19 –22). Moreover, subsequent

This work was supported by National Institutes of Health Grants R01HL101972(to O. J. T. M) and R01GM116184 (to O. J. T. M. and F. L.); R01HL052246(to J. H. G.); R44HL117589 (to A. G.); T32AI007472 (to L. D. H); andR01GM097746, U19AI062629, and P30GM114731 (to F. L.). The authorsdeclare that they have no conflicts of interest with the contents of thisarticle.The content is solely the responsibility of the authors and does notnecessarily represent the official views of the National Institutes of Health.

This article contains supplemental Figs. S1–S4.1 To whom correspondence should be addressed: 3303 SW Bond Ave., Center

for Health & Healing 13B, Portland, OR 97239. Tel.: 503-418-9350; Fax: 503-418-9311; E-mail: healyl@ohsu.edu.

2 An American Heart Association (AHA) Established Investigator (13EIA12630000).

3 The abbreviations used are: APC, activated protein C; rhAPC, recombinanthuman APC; NET, neutrophil extracellular trap; EPCR, endothelial protein Creceptor; Mac-1, macrophase-1 antigen; MPO, myeloperoxidase; PKC, pro-tein kinase C; PAR3, protease-activated receptor 3; PPACK, D-phenylalanyl-prolyl-arginyl chloromethyl ketone; PAF, platelet activating factor; PLC,phospholipase C; LDH, lactate dehydrogenase; fMLP, formylmethionylleu-cylphenylalanine; PFA, paraformaldehyde; HBSS, Hank’s Balanced SaltSolution; NA, numerical aperture; PMA, phorbol 12-myristate 13-acetate;Gla, �-carboxyglutamic acid-rich.

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studies have made a similar observation that PAR3 on theendothelium is cleaved by APC at Arg-41, a noncanonicalposition, which may in part explain the ability of APC toproduce a PAR3-dependent cytoprotective effect observedin podocytes (23, 24). Our aim was to determine whetherAPC was able to ligate EPCR and employ PAR3 to regulateneutrophil cell survival.

One form of neutrophil cell death that has garnered muchinterest since its initial report in 2004 is the ability of neutro-phils to form extracellular traps (25). Neutrophil extracellulartraps (NETs) are comprised of extruded nuclear DNA and asso-ciated nuclear proteins including histones and granular pro-teins. NETs are hypothesized to act as an additional mechanismto promote pathogen clearance. Recent studies have focused onthe characterization of the mechanisms that result in NETosis,as well as the pathophysiological relevance of this process (26 –33). Since their initial discovery, NETs have been shown to pro-mote activation of select coagulation factors and platelets,thereby exacerbating several disease states including thrombo-sis (34 –36). Based on studies linking the activation of bloodcells and coagulation factors with inflammatory processes lead-ing to thrombosis, the concept of immunothrombosis was pro-posed to encompass the complex reactions and cross-talkoccurring in disease states such as sepsis (37). APC has beenproposed to play a role in inhibiting immunothrombosis, basedin part on the observation that APC was found to cleave extra-cellular histones in animal models of sepsis, suggesting a possi-ble link between NETs and APC function (38 – 40). Although itis known that neutrophils express cognate receptors that bindAPC, it is unknown whether APC can inhibit signaling mecha-nisms that potentiate NETosis. We have recently shown thatAPC binds to leukocytes and NETs in a Gla domain-dependentmanner (41). Herein, we present evidence that APC binds neu-trophils to elicit a receptor-mediated intracellular signalingcascade to inhibit neutrophil activation and NETosis. More-over, infusion of pharmacological levels of APC abrogated neu-trophil activation and myeloperoxidase (MPO) release in anonhuman primate model of bacterial sepsis. Together theseresults suggest that the anti-inflammatory properties of APCmay extend to the inhibition of neutrophil cell death, includingNETosis.

Results

APC binds leukocytes and inhibits NETosis

It was first validated that APC bound neutrophils in a staticadhesion assay. Neutrophils were plated on coverslips coatedwith immobilized APC and assessed for adhesion usingdifferential interference contrast microscopy. Neutrophilsbound to APC to a similar degree as to fibronectin; signifi-cantly fewer neutrophils bound to BSA- or IgG-coated cov-erslips (Fig. 1A). It was next studied whether APC in solutionbound to neutrophils undergoing NETosis. The data showthat soluble APC bound to both the activated neutrophil cellbody and the extracellular DNA during NETosis (Fig. 1B).

Subsequently, whether APC binding to neutrophils had afunctional effect on the process of NETosis was investigated.NETs were formed upon incubation of neutrophils with either

autologous platelet secretome or the protein kinase C (PKC)activator PMA. NET formation was characterized by anincrease in surface area of DNA, detection of citrullinated his-tone 3 (H3) and extracellular appearance of MPO (Fig. 2A).Prior incubation of neutrophils with APC caused a significantreduction in the area of DNA and MPO extruded by neutro-phils during NETosis induced by either platelet secretome orPMA (Fig. 2B and supplemental Fig. S1).

PMA was used to induce NETosis in the subsequent mecha-nistic studies because of the enhanced signal-to-noise ratio forsurface area of DNA observed when NETosis was induced byPMA as compared with platelet secretome. Results show that aminimal concentration of 75 nM APC was sufficient to reducethe extent of DNA surface area, whereas 300 nM APC potentlyreduced PMA-induced NET formation (Fig. 3, A–C). Pretreat-ment of APC with the serine protease inhibitor PPACKreversed the inhibitory effect of APC on NETosis. Equimolarconcentrations of the zymogen protein C or the serine proteasethrombin failed to inhibit the extent of DNA extruded by neu-trophils during NETosis (Fig. 3, A and B).

Figure 1. Neutrophils bind APC. Acid-washed glass coverslips were coatedwith denatured BSA (5 mg/ml), 20 �g/ml IgG, 20 �g/ml fibronectin (FN), or100 �g/ml immobilized APC for 1 h, followed by PBS wash and then blockedwith denatured BSA (5 mg/ml). A, purified human neutrophils (2 � 106/ml)were plated on the coverslips for 1 h at 37 °C, followed by washing and fixa-tion. After imaging, cell adhesion was counted in ImageJ and quantified ascell per field of view (FOV). B, neutrophils (2 � 106/ml) were plated onfibronectin-coated coverslips and were treated with HBSS or PMA (10 nM)for 3 h at 37 °C. Cell samples were washed and vehicle (HBSS buffer) or APC(300 nM) was then incubated for 15 min with the cell samples at 37 °C.Samples were then fixed with 4% PFA and incubated overnight with pri-mary antibodies. The samples were then incubated with Hoechst 33342(1:1000) and secondary antibody Alexa Fluor 488 goat anti-mouse (Invit-rogen) (1:500). Images were normalized to secondary antibody-aloneimages. Shown above (C) are representative images of APC-positive stain-ing in the presence of NETs and neutrophils *, p � 0.05 versus BSA. Data aremean � S.E. n�3.

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To determine whether the effect of APC on NETosis wasbecause of a receptor-ligand interaction or because of the directenzymatic cleavage of extracellular DNA, a wash step was intro-duced after incubation of neutrophils with APC and prior toPMA stimulation. As shown in Fig. 3C, an equivalent level ofinhibition of NETosis was observed for experiments in whichAPC was washed away prior to induction of NETosis, with aminimal concentration of 75 nM APC causing a reduction inPMA-induced NET formation. Moreover, inclusion of PPACKin this experiment eliminated the inhibitory effect of APC with-out having an effect on vehicle-treated, PMA-induced NETosis(supplemental Fig. S2A), whereas pretreatment of neutrophilswith an equimolar concentration of thrombin had no signifi-cant effect on the ability of neutrophils to undergo PMA-in-duced NETosis (supplemental Fig. S2B). Because PPACKirreversibly inhibits APC enzyme activity, this implies APCproteolytic activity is required.

Proteins PAF and A1-AT eliminate APC inhibition of NETosis

Other inhibitors of APC activity were studied. Platelet acti-vating factor (PAF) is a phospholipid that binds EPCR, thereby

inhibiting APC binding to EPCR and signaling (42). NET for-mation was detected by staining for DNA and citrullinated H3(Fig. 4A), and PAF eliminated the inhibitory effect of APC onPMA-induced NETosis, as evidenced by an overall DNA areaincrease from 1380 � 47.8 �m2 in the presence of APC alone to2140 � 40.9 �m2 in the presence of APC with PAF (Fig. 4B).

Under physiological conditions, APC is regulated in part bythe heparin-independent plasma inhibitor, �1-antitrypsin (A1-AT), which blocks the proteolytic activity of APC (43). Experi-ments were designed to test whether A1-AT would block theability of APC to inhibit NETosis. When APC was pretreatedwith A1-AT, the DNA area increased from 1380 � 47.8 �m2 forthe presence of APC alone to 2040.5 � 168.2 �m2 when APCwas pretreated with A1-AT (Fig. 4C). As a control, thrombinwas used in lieu of APC, and in either the presence or theabsence of A1-AT, there was no significant change in the abilityof PMA to induce NETosis, suggesting specificity for APC withregard to inhibiting NETosis.

Neutrophil receptors Mac-1, EPCR, and PAR3 mediate APCinhibition of NETosis

Because our results indicated that both the proteolytic activ-ity and the binding of APC to EPCR are required for APC-mediated inhibition of NETosis, we sought to identify thereceptors that mediate this potential cytoprotective function ofAPC. Purified human neutrophils were treated with function-blocking antibodies prior to incubation with APC and subse-quent induction of NETosis with PMA. APC pretreatmentresulted in a significant decrease in DNA area from 2380 � 45.7�m2 to 1010 � 39.9 �m2. After pretreatment of neutrophilswith blocking antibodies to either CD11b or CD18, it was foundthat only in combination was there blockade of APC-mediatedinhibition of NETosis, as evidenced by the increase to 2230 �93.4 �m2 of DNA area in the presence of the combination ofthese antibodies (Fig. 5A), whereas a blocking antibody toCD11a had no effect (data not shown). The ability of APC toinhibit PMA-induced NETosis was eliminated in the presenceof a blocking antibody to EPCR (RCR-252), as evidenced by acorresponding increase in DNA area to 2260 � 34.4 �m2. Nota-bly, neutrophil pretreatment with a blocking antibody to PAR3blocked the ability of APC to inhibit NETosis, as evidenced bythe increase of DNA area to 2080 � 104 �m2. In contrast, pre-treatment with a specific small molecule PAR1 antagonist,SCH79797, was found to have no significant effect on the abilityof APC to inhibit NETosis (supplemental Fig. S3). As a positivecontrol, platelets pretreated with SCH79797 resulted in theinhibition of TRAP-induced platelet activation (data notshown).

Because the use of an antibody that blocked the cleavage ofPAR3 resulted in the loss of APC-mediated inhibition of NETo-sis, we sought to test whether the PAR3 tethered-ligand pep-tides generated by APC or thrombin cleavages could inhibitNET formation (24). The thrombin-derived P3K peptide,which is generated after the cleavage of PAR3 at Lys-38 bythrombin, had no effect on PMA-induced NET formation. Incontrast to the P3K peptide, the use of the APC-derived P3Rpeptide, which is generated after the cleavage of PAR3 atArg-41 by APC, significantly reduced PMA-induced NETosis

Figure 2. APC treatment inhibits NETosis. Acid-washed glass coverslipswere coated with 20 �g/ml fibronectin and then blocked with denatured BSA(5 mg/ml). Purified human neutrophils (2 � 106/ml) were plated on the cov-erslips for 30 min at 37 °C, then incubated with APC (300 nM) for 30 min at37 °C. Samples were then washed once with PBS and subsequently treatedwith HBSS, platelet secretome, or PMA (10 nM) for 3 h at 37 °C. All sampleswere then fixed with 4% PFA. Samples were incubated overnight with poly-clonal mouse anti-myeloperoxidase antibody (MPO) (1:100) and rabbit anti-cit-rullinated histone 3 antibody (H3Cit) (1:250). Samples were then incubated withHoechst 33342 (1:1000) and secondary antibodies Alexa Fluor 488 goat anti-rab-bit and 546 goat anti-mouse IgG (Invitrogen) (1:500). Images were normalized tosecondary antibody-alone (vehicle control) images. A, representative images ofneutrophils and NETs with DNA, MPO, and H3Cit staining are shown. B, imageswere then analyzed in a custom MATLAB program to quantify each pixel-positivesignal as area DNA per image. *, p � 0.05 versus DMSO � platelet secretome. #,p � 0.001 versus DMSO � PMA. Data are mean � S.E. n � 4.

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in either the presence or absence of APC (Fig. 5B), implying thatPAR3 agonism was sufficient to inhibit NETosis.

APC signals via G�� to inhibit PKC and PI3K signaling-depen-dent NET formation

The signaling pathways that regulate NETosis were studied,first by validation of the pathways by which platelet secretomeand PMA potentiate NETosis. Pretreatment of neutrophilswith the PI3K inhibitor wortmannin eliminated NET formationinduced by either platelet secretome or the PKC activator PMA(Fig. 6A). Moreover, pretreatment of neutrophils with U73122,an inhibitor to phospholipase C (PLC) that acts upstream ofPKC, abrogated NET formation stimulated by platelet secre-tome but not PMA, whereas the inactive analog of the PLCinhibitor U73343 had no effect on NETosis potentiated byeither agonist. Pretreatment of platelet secretome with the ser-ine protease inhibitor PPACK had no effect on the ability ofplatelet secretome to induce NETosis (data not shown).Whether there was a role for cytokine receptor-mediated sig-

naling in platelet secretome-induced NETosis was tested next.Pretreatment of neutrophils with the JAK/STAT inhibitorcucurbitacin I significantly reduced the degree of DNAextruded from neutrophils following exposure to plateletsecretome while having no effect on PMA-induced NETosis(Fig. 6A).

Next, the signaling pathways by which APC inhibits PMA-induced NETosis were characterized. As PMA is a diacylglyc-erol analog, it activates PKC to drive NETosis in vitro in a PI3K-dependent manner (30, 31, 44). First, the role of PKC signalingpathway in PMA-induced NETosis was confirmed. As seen inFig. 6B, pharmacological inhibition of PKC with eitherGF109203X or Ro31– 8220 ablated the ability of PMA to induceNETosis. In contrast, inhibition of PLC or G�� signaling withU73122 or gallein, respectively, had no effect on PMA-inducedNETosis, yet eliminated the ability of APC to inhibit NETosis,consistent with the findings suggesting that APC induces sig-naling downstream of the G protein-coupled receptor PAR3 toinhibit NETosis. Of note, a partial inhibition of NETosis was

Figure 3. APC treatment inhibits PMA-induced NETosis. Acid-washed glass coverslips were coated with 20 �g/ml fibronectin and then blocked withdenatured BSA (5 mg/ml). Purified human neutrophils (2 � 106/ml) were plated on the coverslips for 30 min at 37 °C. A and B, they were then incubated withincreasing concentrations of APC (30, 100, and 300 nM), protein C (300 nM), and thrombin (300 nM) in the presence or absence of PPACK (40 �M) for 30 min at37 °C. Samples were washed and then subsequently treated with HBSS or PMA (10 nM) for 3 h at 37 °C. C, purified human neutrophils (2 � 106/ml) were platedon the coverslips for 30 min at 37 °C, then incubated with increasing concentrations of APC (30, 75, 100, 150, and 300 nM) for 30 min at 37 °C. Samples were thenwashed once with PBS and subsequently treated with HBSS or PMA (10 nM) for 3 h at 37 °C. All samples were then fixed with 4% PFA. Samples were incubatedovernight with polyclonal rabbit anti-neutrophil elastase antibody (1:100). The samples were then incubated with Hoechst 33342 (1:1000) and secondaryantibody Alexa Fluor 546 goat anti-rabbit IgG (Invitrogen) (1:500). Images were normalized to secondary antibody-alone images. Shown in A are representativeimages of PMA-induced NETs in the presence of increasing concentrations of coagulation factors. Images were analyzed in a custom MATLAB program toquantify each pixel-positive signal as area DNA per image, shown in B and C. *, p � 0.001 versus vehicle � PMA. Data are mean � S.E. n � 3.

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observed for gallein in the absence of APC, likely because of thefact that gallein is also known to exhibit a partial off-targetinhibitory effect on the PI3K pathway (Fig. 6B).

The protease and Gla domains of APC mediate the inhibitionof NET formation

Subsequent experiments were designed to determine theregions of APC responsible for inhibiting NETosis. A panel ofrecombinant APC mutants was rationally created with muta-tions within the active site and charged exosite of the proteasedomain, as well as within the Gla domain, as depicted in Fig.7A. Equimolar concentrations of recombinant human APC(rhAPC) inhibited PMA-induced NETosis to the same degreeas plasma-derived APC (Fig. 7B). The proteolytic activity ofAPC was required to inhibit NETosis. Consistent with the find-ings that demonstrated the action of APC was blocked by theserine protease inhibitor PPACK (Fig. 3), the inhibitory func-tion of APC was lost when the active site residue Ser-360 wasmutated to Ala, rendering APC catalytically inactive (S360A-APC) (Fig. 7C).

Recombinant APC mutants with protease domain mutationswere used to determine structure-activity relationships forinhibition of NETosis. APC with the mutation Asn-329 to Gln

(N329Q-APC) exhibited a loss of function and was unable toinhibit NETosis. The N329Q mutation results in the loss of anattached carbohydrate that possibly participates in APC-recep-tor interactions. In the same sequence as Asn-329, substitutionof two glutamic acid residues for alanine (E330A/E333A-APC)resulted in only partial inhibition of PMA-induced NETosis,suggesting that residues 329 –333 are required for normal inhi-bition of NETosis by APC. Mutation of Ser-252 located alongthe opening of the catalytic pocket to the acidic residue Glu(S252E-APC) also eliminated the ability of APC to inhibitNETosis. However, mutation at the same residue to alanine(S252A-APC) did not impair the ability of APC to inhibitNETosis (Fig. 7C).

As mutations in and around the active site seem to play acritical role for APC inhibition of NETosis, we next sought todetermine whether charged exosites on the face opposite fromthe face containing the active site were necessary to inhibitNETosis. A mutation that enhances protein S cofactor activityby mutating the negatively charged Glu-149 to Ala caused apartial reduction in the ability of E149A-APC to inhibit PMA-induced NETosis. Next, it was determined if the positivelycharged exosites in the protease domain played a role in thefunctional inhibition of NET formation by APC. There was a

Figure 4. APC inhibition of NETosis is abrogated by protein platelet activating factor and �1-antitrypsin. Acid-washed glass coverslips were coated with20 �g/ml fibronectin and then blocked with denatured BSA (5 mg/ml). Purified human neutrophils (2 � 106/ml) were plated on the coverslips for 30 min at37 °C and allowed to adhere before APC treatments. A, cells were washed once with PBS and incubated with APC (100 nM) for 30 min at 37 °C. B, to test forinhibition of APC, cells were washed once with PBS and incubated with buffer (vehicle) or platelet activating factor (PAF) (100 nM) for 10 min at 37 °C prior toincubation with APC (100 nM) for 30 min at 37 °C. C, surface-bound cells were incubated with APC (300 nM) and thrombin (300 nM) in the presence or absenceof �1-antitrypsin (�1-AT) (20 �M) for 30 min at 37 °C. Samples were then washed once with PBS and subsequently treated with HBSS or PMA (10 nM) for 3 h at37 °C. They were then fixed with 4% PFA. The samples were incubated overnight with polyclonal rabbit anti-citrullinated histone H3 antibody (1:250) and werethen incubated with Hoechst 33342 (1:1000) and secondary antibody Alexa Fluor 488 goat anti-rabbit IgG (Invitrogen) (1:500). Images were normalized tosecondary antibody-alone images and analyzed in a custom MATLAB program to quantify each pixel-positive signal as the area of DNA per image. *, p � 0.05versus vehicle � PMA. #, p � 0.05 versus APC � PMA. Data are mean � S.E. n � 3.

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partial loss of function for APC when the residues in the calci-um-binding binding loop were mutated to alanine (R229A/R230AAPC). In contrast, mutation of several positivelycharged lysine residues 191–193 to alanine in the 37-loop hadno effect on the inhibitory function of APC, as evidenced by anequivalent degree of inhibition of NETosis for the 3K3A-APCmutant and rhAPC (Fig. 7D).

The protease domain of APC contains an Arg-Gly-Asp(RGD) triad that is canonically recognized by integrins; thus,Asp-180 in the RGD motif was mutated to determine whetherthe RGD motif of APC played a role in the regulation of NETo-sis. D180E-APC had normal ability for inhibition of NETosis(Fig. 7D).

The �-carboxyglutamic acid-rich (Gla) domain of APC playsan essential role mediating binding to both EPCR and phos-phatidylserine exposed on the surface of activated blood andendothelial cells (45). Consistent with data in Fig. 5 implicating

a role for EPCR in mediating the inhibitory function of APC, theGla domain mutation of Leu-8 to Val (L8V-APC) caused a par-tial loss of activity (Fig. 7E).

The anticoagulant function of APC is enhanced when boundto the cofactor protein S, increasing the rate of inactivation ofFVa and FVIIIa. Recently, there was demonstrated a critical rolefor protein S and cleaved FVa acting as cofactors for APC anti-inflammatory function in a mouse model of septic peritonitis(46). Moreover, studies have demonstrated that binding of pro-tein S to APC requires the APC residue Leu-38 in the Gladomain, as mutation to Asp (L38D-APC) impairs APC and pro-tein S interactions (47). The L38D mutation caused a partialloss of the NETosis inhibitory activity of APC. The substitutionof the bulky polar uncharged amino acid, Gln for hydrophobicLeu-38 in APC produced a complete loss of function (Fig. 7E),

Figure 5. EPCR, PAR-3, and Mac-1 are required for APC-mediated inhibi-tion of NETosis. Acid-washed glass coverslips were coated with 20 �g/mlfibronectin and then blocked with denatured BSA (5 mg/ml). Purified humanneutrophils (2 � 106/ml) were plated on the coverslips and allowed to adherefor 30 min at 37 °C. Cells were washed once with PBS. A, cells were incubatedwith CD11b blocking antibody (ab8878) (10 �g/ml), CD18 blocking antibody(CBL158) (1 �g/ml), EPCR blocking antibody (RCR-252) (10 �g/ml), or PAR3blocking antibody (ab-5598) (25 �g/ml) for 10 min at 37 °C prior to incubationwith APC (300 nM) for 30 min at 37 °C. B, cells were incubated with PAR3peptides P3K or P3R (50 �M) representative of cleavage at Lys-38 (P3K) orArg-41 (P3R) in either the presence or absence of APC (300 nM) for 30 min at37 °C. Samples were then washed once with PBS and subsequently treatedwith HBSS or PMA (10 nM) for 3 h at 37 °C. Samples were then fixed with 4%PFA and were incubated overnight with polyclonal rabbit anti-neutrophil ela-stase antibody (1:100) or polyclonal rabbit anti-citrullinated histone H3 anti-body (1:250). The samples were then incubated with Hoechst 33342 (1:1000)and secondary antibody Alexa Fluor 546 goat anti-rabbit IgG (Invitrogen)(1:500). Images were normalized to secondary antibody-alone images andanalyzed in a custom MATLAB program to quantify each pixel-positive signalas the area of DNA per image. *, p � 0.001 versus vehicle � PMA. #, p � 0.05versus APC � PMA. Data are mean � S.E. n � 3.

Figure 6. APC-mediated inhibition of NETosis requires G�� signaling toinhibit PKC and PI3K. Acid-washed glass coverslips were coated with 20�g/ml fibronectin and then blocked with denatured BSA (5 mg/ml). Purifiedhuman neutrophils (2 � 106/ml) were plated on the coverslips, and allowedto adhere for 30 min at 37 °C. Cells were washed once with PBS. A, cells wereincubated with PI3K inhibitor wortmannin (100 nM), PLC inhibitor U73122 (5nM), the inactive analog U73343 (5 nM), or JAK/STAT inhibitor cucurbitacin I(500 nM) for 30 min at 37 °C. B, cells were incubated with PKC inhibitorsGF109203X (5 �M) and Ro31– 8220 (380 �M), PLC inhibitor U73122 (5 nM), orG�� inhibitor gallein (10 �M) for 30 min at 37 °C prior to incubation with APC(300 nM) for 30 min at 37 °C. Samples were then washed once with PBS andsubsequently treated with HBSS, autologous platelet secretome, or PMA (10nM) for 3 h at 37 °C. Samples were then fixed with 4% PFA and incubatedovernight with polyclonal rabbit anti-citrullinated histone H3 antibody(1:250). The samples were then incubated with Hoechst 33342 (1:1000) andsecondary antibody Alexa Fluor 488 goat anti-rabbit IgG (Invitrogen) (1:500).Images were normalized to secondary antibody-alone images and analyzedin a custom MATLAB program to quantify each pixel-positive signal as thearea of DNA per image. *, p � 0.001 versus vehicle � PMA. #, p � 0.001 versusAPC � PMA. Data are mean � S.E. n � 3.

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indicating that residue Leu-38 located on the top of the Gladomain is critical for inhibition of NETosis.

Platelet adhesion to NETs is inhibited by APC

Experiments were designed to determine whether APCcould inhibit platelet-NET binding, which has been implicatedas playing a pathological role in promoting immunothrombosis(34 –36). First, it was validated that platelets bound immobi-lized fibronectin, as measured by staining for the platelet integ-rin CD41 (Fig. 8A). Next, platelet adhesion to fibronectin-bound neutrophils was quantified. The results show thatinduction of NETosis with PMA potentiated platelet adhesion.Pretreatment of neutrophils with APC reduced the degree ofplatelet adhesion by more than 50%, as measured by either thetotal degree of CD41 staining (Fig. 8A) or the overlap of CD41with DNA (Fig. 8B).

LDH release is reduced by APC pretreatment

Experiments were next designed to test if neutrophil celldeath was inhibited by APC. Lactate dehydrogenase (LDH) isan enzyme found in almost all cells including neutrophils, play-ing an integral role in changing lactate to pyruvate, and LDHrelease is a marker of cell death (48). Neutrophil cell death wasinduced with PMA and quantified as percent cytotoxicity,where under vehicle conditions it resulted in nearly 90% cyto-

toxicity (Fig. 8C). When neutrophils were pretreated with APC,the percentage of cytotoxic cells was significantly reduced bynearly 40%.

MPO plasma levels are inhibited by APC infusion in anonhuman primate model of sepsis

As an in vivo proof-of-concept experiment, we studiedwhether the administration of pharmacological doses of APCunder pathophysiological conditions could reduce neutrophilactivation using an established baboon model of bacterial sepsis(40, 49, 50). In this model, either pharmacological doses of APC(3 mg/kg initial bolus followed by infusion of APC at 16 �g/kg/min for 5.4 h) or of vehicle were infused prior to challenge witha lethal dose (LD100) of Escherichia coli (ATCC® 12701) thatresulted in a total concentration of 1–2 � 1010 colony formingunits (cfu)/kg. Using this model, the effect of APC on neutro-phil MPO release was assessed. MPO is a critical antimicrobialenzyme released by activated neutrophils as well as a biomarkerfor NET formation (48, 51). As seen in Fig. 9, E. coli challengeinduced a nearly 7-fold increase in MPO release from a baselineof 31.1 � 4.3 mU/ml to 196.4 � 45.8 mU/ml in nontreatedanimals; in striking contrast, pretreatment of baboons withAPC nearly abrogated MPO release, as evidenced by the level ofMPO release only increasing to 38.6 � 2.7 mU/ml 24 h postchallenge.

Figure 7. Identification of APC residues required for APC-mediated inhibition of NETosis. A, structural scheme for APC with residue mutations numberedbeginning with the active site Ser-360 in the protease domain. B–E, acid-washed glass coverslips were coated with 20 �g/ml fibronectin and then blocked withdenatured BSA (5 mg/ml). Purified human neutrophils (2 � 106/ml) were plated on the coverslips for 30 min at 37 °C, then incubated with 300 nM APC or APCmutants for 30 min at 37 °C. Samples were then washed once with PBS subsequently treated with HBSS or PMA (10 nM) for 3 h at 37 °C. They were then fixed with4% PFA and were incubated overnight with polyclonal rabbit anti-neutrophil elastase antibody (1:100). Samples were then incubated with Hoechst 33342(1:1000) and secondary antibody Alexa Fluor 488 goat anti-rabbit IgG (Invitrogen) (1:500). Images were normalized to secondary antibody-alone images andanalyzed in a custom MATLAB program to quantify each pixel-positive signal as the area of DNA per image. B, compares wtAPC versus rhAPC used to make allmutants, where all mutations within the active site are presented in C, the charged exosite mutations are presented in D, and the Gla domain mutations in APCare presented in E. *, p � 0.001 versus vehicle � PMA. #, p � 0.05 versus rhAPC � PMA. �, p � 0.05 versus L38D � PMA. �, p � 0.05 versus S252A �PMA. Data are mean � S.E. n � 3.

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Discussion

Here, we report that in a purified neutrophil system and in aninitial proof-of-concept experiment using an established in vivononhuman primate model of sepsis, using therapeutic concen-trations of APC was found to inhibit neutrophil activationand induction of cell death. Utilizing a mechanistic in vitroapproach, APC inhibition of PKC- and PI3K-dependent NETo-sis was found to be in part dependent on the neutrophil recep-tors Mac-1, EPCR, and PAR3 (Fig. 10). EPCR and � integrinsexpressed on neutrophils are known to bind APC resulting in areduction in migration (14, 15, 18), and our results now suggesta role for the receptors EPCR and Mac-1 in APC-mediatedinhibition of NETosis. Based on molecular studies with endo-thelial cells and podocytes, APC ligates EPCR and cleaves PAR1and PAR3 at noncanonical residues differing from the throm-bin cleavage sites (19, 24). Together with our results, these stud-ies substantiate the biased agonism hypothesis in which throm-bin and APC differentially cleave the same receptors, thus

Figure 8. APC inhibits platelet adhesion to NETs and LDH release. A and B, acid-washed glass coverslips were coated with 20 �g/ml fibronectin and thenblocked with denatured BSA (5 mg/ml). Purified human neutrophils (2 � 106/ml) were plated on the coverslips and allowed to adhere for 30 min at 37 °C. Cellswere washed once with PBS and incubated with HBSS or 300 nM APC 30 min at 37 °C. Samples were then washed once with PBS and subsequently treated withHBSS or PMA (10 nM) for 3 h at 37 °C. The samples again were washed once with PBS and incubated with autologous platelets (2 � 107/ml) and allowed toadhere for 45 min at 37 °C. Samples were then washed once with PBS and fixed with 4% PFA and were incubated overnight with polyclonal rabbit anti-CD41antibody (1:100). They were then incubated with Hoechst 33342 (1:1000) and secondary antibody Alexa Fluor 488 goat anti-rabbit (Invitrogen) (1:500). Imageswere normalized to secondary antibody-alone images and analyzed in a custom MATLAB program to quantify each pixel-positive signal as (A) the area of FITC(CD41) per image. In (B) each pixel positive for both CD41 and DNA signals per image was quantified. *, p � 0.001 versus vehicle � PMA. C, purified humanneutrophils (4 � 105/ml) were incubated in solution with 300 nM APC for 30 min at 37 °C. Samples were then incubated with HBSS or PMA (20 nM, final) andincubated for 3 h at 37 °C. 10� lysis buffer was added to duplicate columns and incubated for 45 min at 37 °C. Supernatants from samples and the LDH-positivecontrols were transferred to new 96-well plates prior to incubation with reaction mixture for 30 min at 37 °C. The reaction was stopped and absorbance wasmeasured at 490 nm and 680 nm. Percent cytotoxicity was quantified per the manufacturer’s instructions (Thermo Fisher). *, p � 0.05 versus vehicle � PMA.Data are mean � S.E. n � 3.

Figure 9. Infusion of APC reduces plasma levels of MPO in an LD100E. coli baboon model of sepsis. In this established baboon model ofsepsis, Papio c. anubis-treated animals were initially dosed with 3 mg/kgbolus of APC at time (T) � 10 min. From T � 0 min to T � 320 min, APC wasinfused intravenously at 16 �g/kg/min. A lethal dose (1–2 � 1010 cfu/kg)of E. coli, ATCC 12701 serotype O86:K61(B7) was infused intravenouslyover 2 h (from T � 0 to T � 120 min). Blood samples were obtained at 0, 2,4, 8, and 24 h and plasma samples were isolated. MPO in plasma sampleswas determined using Fluoro MPO (fluorescent myeloperoxidase detec-tion kit) (Cell Technology) per the manufacturer’s instructions. Data aremean � S.E. LD100 animals n � 8; LD100 � APC animals n � 4.

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initiating different signaling pathways to culminate in disparateeffects on cell survival (11, 19, 23, 52). PAR3 was identified in1997 as the second thrombin receptor in humans, followingPAR1 (53). Initially, PAR3 was thought to be a nonsignalingreceptor. PAR3 was shown to replace PAR1 and facilitate PAR4activation by thrombin (54). PAR1 and PAR3 heterodimerizeon endothelial cells to help explain thrombin-mediated signal-ing (55). Thus far, it has been shown that thrombin, APC, andFXa can cleave PAR3 (24, 53, 56, 57). APC-mediated cytopro-tective signaling on neuronal cells, podocytes, and endothelialcells occurs in part via the activation of PAR3 and requiresEPCR (24, 58 – 60). It remains to be determined whether APCgenerated in plasma from endogenous protein C inhibits neu-trophil activation or extends cell survival.

In neutrophils, basal mRNA and cell surface expression ofPAR3 remains relatively constant in response to various inflam-matory agonists (16, 61). APC-mediated PAR3 signaling onneutrophils inhibited NETosis, whereas equimolar concentra-tions of thrombin-mediated PAR3 signaling had no effect onNETosis. The striking finding that the PAR3 P3R peptide inhib-its NETosis implies that neutrophil-expressed PAR3 biasedagonism is sufficient to regulate neutrophil survival.

With the novel observation that PAR3 but not PAR1 had anessential role in APC-mediated inhibition of NETosis, it is anopen question whether there are two signaling pathways initi-ated that feed into one common path under pathophysiologicalconditions. It is possible that Mac-1 and a PAR3-EPCR com-plex each initiate distinct pathways to activate inhibitory signal-ing; alternatively, Mac-1/PAR3/EPCR may form a larger com-plex to initiate signaling, similar to the clustering of EPCR/PAR1 in caveolae found on endothelial cells (62). Our NETosisinhibition data add to the repertoire of mechanisms by whichadministration of APC exerts anti-inflammatory effects.

Prior to its removal from clinical practice, recombinant APCwas used in adult patients with severe sepsis based on therationale that the anticoagulant and anti-inflammatory effectsof APC would provide a clinical benefit in this patient popula-tion (10). Additional benefits may derive from the ability ofAPC to cleave cytotoxic extracellular histones in animal modelsof sepsis (40), supporting the concept that APC exerts its anti-inflammatoryeffectsthroughbothcell-dependentandcell-inde-pendent pathways. To date, it has not been clear whether APChas a direct cytoprotective action on neutrophil cell survival. Asour results demonstrated, APC inhibited neutrophil cell deathbut did not affect elastase release (supplemental Fig. S4). Thisraises the possibility that APC may exert anti-inflammatoryactivity by inhibiting cellular functions that potentially drive oramplify pathophysiological disease, whereas APC may preservefunctions required for homeostatic surveillance. Under selectinflammatory conditions, the interactions between plateletsand NETs are hypothesized to potentiate immunothrombosis;therefore, we tested if APC reduced platelet-NET binding (34,35, 63). Our results suggest that through inhibition of neutro-phil cell death, APC reduces platelet-NET binding and thusmay reduce the prothrombotic phenotype of NET-promotedimmunothrombosis.

Because APC has well characterized anticoagulant and mul-tiple cell signaling actions, it should become possible throughrational design to select for specific functions of APC (11).Thus, one of our goals is to develop APC mutants that canprovide improved treatments for a variety of disease states, as insepsis where these proteins may allow for the better control ofinflammation and immune system homeostasis without com-promising hemostatic mechanisms (11). Introduction of singleor double amino acid substitutions within APC can signifi-cantly modulate its anticoagulant or anti-inflammatory func-tions. Proof-of-concept studies using APC mutants have dem-onstrated beneficial selection for specific functions of APC on avariety of cell types and through use of in vivo mouse models,where select mutants have now advanced to clinical trial (47, 58,63– 69). Using the protein C database ProCMD (70), five of thesites where mutations were introduced into APC for our studywere found to occur in humans; however, the amino acidsintroduced in our study did not reflect the mutated amino acidreplacements seen in affected humans. Multiple APC mutantsinhibited PMA-induced NETosis. Select mutations in the pro-tease domain, as in 3K3A-APC, yielded a normal degree of inhi-bition of NETosis, similar to rhAPC. This APC signaling selec-tive mutant is in clinical trials for ischemic stroke (71). Incontrast, multiple mutations in the protease domain have a par-tial or total loss of function, demonstrating that certain residuesin the protease domain are essential for the ability of APC sig-naling to inhibit NETosis. Moreover, mutations within the Gladomain of APC result in the partial or entire loss of function ofAPC, highlighting the additional importance of the Gla domainto mediate both the binding and orientation of APC on neutro-phils to inhibit NET formation. These findings support therationale for development of APC mutants that retain cytopro-tective and anti-inflammatory functions but shed anticoagulantactivity for potential clinical use in selected inflammatory dis-ease states.

Figure 10. Schematic overview for APC-mediated inhibition of NETosis.In vitro, select stimuli including cytokines, bacteria, the PKC-agonist PMA, andthe platelet secretome induce PKC-dependent NETosis. Our observationsshow a mechanism whereby proteolytically active APC requires Mac-1, EPCR,and PAR3 to induce intracellular, cytoprotective signaling that results in thedownstream inhibition of NET formation.

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Because the in vitro studies show that APC modulates neu-trophil cell death, we determined whether infusion of APC at apharmacologically relevant dose could reduce neutrophil acti-vation using an established nonhuman primate model of bac-terial sepsis as an initial proof-of-concept experiment. 3K3A-APC is currently being studied at high doses, up to 0.5 mg/kg, inhealthy adults and is well tolerated (71). Here we showed thatbaboons infused with lethal doses of E. coli, when using recom-binant wild-type APC at 3 mg/kg followed by infusion for �5 hat 0.96 mg/kg/h, inhibited MPO release. This suggests a link invivo between APC and neutrophil function and points towardfuture in vivo studies. Of potential relevance, a previous studyobserved that in heart surgery patients, a negative correlationbetween neutrophil MPO activity in the myocardium and gen-eration of APC was observed post surgery (72).

In conclusion, our discoveries uncover a novel mechanism bywhich pharmacological concentrations of APC utilize neutro-phil-expressed Mac-1, EPCR, and PAR3 to activate intracellularsignaling that culminates in the inhibition of PMA-inducedNETosis. Furthermore, a pilot proof-of-concept experimentusing a nonhuman primate model of bacterial sepsis demon-strates that infusion of APC reduced neutrophil activation invivo. Fundamental questions remain about how these threereceptors ligate APC and initiate the specific intracellular sig-naling pathways that are activated to result in inhibition, andthe role of endogenous APC in reducing NETosis in vivo.Future work will aim to identify the precise signaling networksthat result in this inhibition of NETosis by APC.

Experimental procedures

Reagents

Human plasma-derived APC was a gift from the AmericanRed Cross (73). Recombinant human APC and mutant APCproteins were cloned, expressed, and purified as described pre-viously (64 – 66).

The PKC inhibitors GF109203X and Ro31– 8220; as well asthe G�� inhibitor gallein, were purchased from Tocris Bio-Techne (Minneapolis, MN). Polymorphprep was from Axis-Shield Density Gradient Media (Oslo, Norway). The cell-per-meable DNA dye Hoechst 33342 was from Invitrogen. AlexaFluor conjugated anti-mouse antibodies, rabbit polyclonal anti-CD41 (ab63983), rabbit polyclonal anti-histone H3 (ab5103),polyclonal rabbit anti-neutrophil elastase antibody (ab21595),rabbit monoclonal anti-CD11a (ab52895), rat monoclonal anti-C11b (ab8878), and rat monoclonal anti-EPCR/CD201 (RCR-252) (ab81712) were from Abcam (Cambridge, MA). Mousemonoclonal antibody to CD18 (CBL 158) was from EMD Mil-lipore (Darmstadt, Germany). Blocking antibody to proteaseactivated receptor, PAR3 (H-103, sc-5598) was purchased fromSanta Cruz Biotechnology (Dallas, TX). Anti-human mouseantibody to protein C/APC (AHPC-5071) was purchased fromHaematologic Technologies Inc. (Essex Junction, VT). PAR3-derived peptides GAPPNSFEEFPFS (P3R) and TFRGAPPNS-FEEF (P3K) were synthesized and purified to �95% purity fromSynthetic Biomolecules (San Diego, CA). Pierce™ LDH Cyto-toxicity Assay Kit (88953) was purchased from Thermo FisherScientific. Neutrophil Elastase Activity Assay Kit (600610) was

purchase from Cayman Chemical (Ann Arbor, MI). Escherichiacoli serotype O86:K61(B7), ATCC® 12701, was purchased fromATCC (Manassas, VA) All other reagents were purchased fromSigma-Aldrich or previously described sources (74).

Preparation of leukocytes

Human leukocytes were purified as described previously(75). Briefly, human blood was drawn in accordance with anOregon Health & Science University Institutional ReviewBoard-approved protocol from healthy donors by venipunctureinto citrate-phosphate-dextrose (1:7 v/v). Blood was layeredover an equal volume of PolymorphprepTM and centrifuged at500 � g for 45 min at 18 °C. The lower layer containing neutro-phils was subsequently collected and washed with Hank’s Bal-anced Salt Solution (HBSS) by centrifugation at 400 � g for 10min. To remove red blood cells from the sample, the pellet wasresuspended in sterile H2O for 30 s, followed by immediateaddition of 10� PIPES buffer (250 mM PIPES, 1.1 mM CaCl2, 50mM KCl, pH 7.4), to dilute the leukocyte suspension. After cen-trifugation at 400 � g for 10 min, the pellet was resuspended inHBSS containing 2 mM CaCl2, 2 mM MgCl2, and 1% w/v bovineserum albumin (BSA).

Platelet isolation and secretome preparation

Washed platelet isolation was carried out as described previ-ously (76). Thrombin-induced (1 unit/ml) platelet aggregationwas allowed to proceed for 15 min at 37 °C after its onset andthe platelet secretome, containing platelet-derived soluble mol-ecules, was isolated as described previously (77). Briefly, acti-vated platelets were pelleted by sequentially centrifuging twiceat 1000 � g for 10 min in the presence of a protease inhibitormixture to prevent protein degradation. Platelet pellet was dis-carded and the autologous supernatant (secretome) was col-lected and recentrifuged at 13,000 � g for 10 min to remove anymicroparticles. Hirudin (40 �g/ml) was added to the superna-tant to neutralize thrombin activity. The platelet supernatantwas used to stimulate NETosis as described below.

Immunofluorescence microscopy

Acid-washed glass coverslips were coated with 20 �g/mlfibronectin and then blocked with denatured BSA (5 mg/ml).Initially, purified human neutrophils (2 � 106/ml) were platedon the coverslips for 30 min at 37 °C. In select experiments,APC or thrombin was pretreated with PPACK dihydrochloride(40 �M) for 15 min prior to incubation with neutrophils. Sam-ples were subsequently treated with HBSS or phorbol 12-my-ristate 13-acetate (10 nM) for 3 h at 37 °C prior to fixationwith 4% paraformaldehyde (PFA) followed by incubation withblocking buffer (PBS containing 10% FBS and 5 mg/ml FractionV BSA). Cells were stained with primary antibody (1:100 or1:250) in blocking buffer at 4 °C overnight. Secondary goat anti-mouse IgG antibodies conjugated with Alexa Fluor 546 (1:500)and Hoechst 33342 (10 �g/ml) in blocking buffer were addedand incubated for 2 h in the dark. Coverslips were mountedonto glass slides using Fluoromount G and visualized with aZeiss Axiovert fluorescence microscope (Axio Imager; CarlZeiss, Göttingen, Germany) equipped with a 40�/1.3 numeri-

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cal aperture (NA) oil immersion objective and an air-coupledlens providing Köhler illumination at an NA of 0.17.

To examine platelet binding, following fixation with 4% para-formaldehyde, platelets were permeabilized with blocking solu-tion (1% BSA, 0.1% SDS in PBS). Platelets were then stainedwith indicated antibodies (1:100) overnight at 4 °C in blockingbuffer. Secondary Alexa Fluor antibodies (1:500) with Hoechst(1:1000) were added in blocking buffer for 2 h in the dark. Aftermounting with Fluoromount G, platelets and neutrophils wereimaged on a Zeiss Axiovert fluorescence microscope (AxioImager; Carl Zeiss) equipped with a 63� NA oil immersionobjective and an air-coupled lens providing Köhler illuminationat an NA of 1.40. All images were recorded with a charge-cou-pled device camera (AxioCam MRc5 12-bit camera; Carl Zeiss)under software control by Slidebook 5.5 (Intelligent ImagingInnovations, Denver, CO) as described previously (76).

Neutrophil static binding assay

Acid-washed glass coverslips were coated with BSA (dena-tured and filtered 5 mg/ml), IgG (20 �g/ml), fibronectin (20�g/ml), or APC (100 �g/ml) for 1 h, followed by washing andthen blocked with denatured BSA (5 mg/ml) for 1 h. Initially,purified human leukocytes (2 � 106/ml) were plated on thecoverslips for 1 h at 37 °C. Cells were then washed again withPBS and fixed with 4% PFA followed by mounting and imaging.

APC binding assay

Purified human leukocytes (2 � 106/ml) were stimulatedwith HBSS or PMA (10 nM) for 3 h at 37 °C on fibronectin-coated glass coverslips. For colocalization experiments, adher-ent cells were washed and incubated with vehicle (HBSS buffer)or APC (300 nM) for 15 min at 37 °C. Subsequently, sampleswere washed with PBS and fixed with 4% PFA followed by incu-bation with blocking buffer (PBS containing 10% FBS and 5mg/ml Fraction V BSA). Cells and coagulation factors werestained anti-PC/APC Abs (100 �g/ml) in blocking buffer at 4 °Covernight. Secondary goat anti-mouse IgG antibodies conju-gated with Alexa Fluor 488 (1:500) and Hoechst 33342 (10�g/ml) in blocking buffer were added and incubated for 2 hin the dark. Coverslips were mounted onto glass slides andvisualized.

NET formation assays

In select experiments, purified human leukocytes (2 � 106/ml) were plated on fibronectin-coated coverslips for 30 min at37 °C. APC (300 nM) and thrombin (300 nM) were incubated for15 min in the presence or absence of PPACK (40 �M) or �1-an-titrypsin (20 �M) prior to incubation with leukocytes for 30 minat 37 °C. For select experiments, neutrophils were pretreatedwith platelet activating factor (PAF, 100 nM) for 30 min at 37 °Cprior to incubation with 100 nM APC for an additional 30 min at37 °C.

In separate experiments, cells were incubated with PKCinhibitors GF109203X (5 �M) or Ro31– 8220 (380 �M); thePI3K inhibitor wortmannin (100 nM); the PLC inhibitorU73122 (5 nM) or the inactive analog U73343 (5 nM); the JAK/STAT inhibitor curcurbitacin I (500 nM); the blocking CD11bantibody (ab8878) (10 �g/ml), the blocking CD18 antibody

(CBL158) (1 �g/ml); the blocking EPCR antibody (RCR-252)(10 �g/ml); the blocking PAR3 antibody (H-103) (25 �g/ml); orthe PAR3 peptides derived from cleavage at Arg-41 (P3R) (50�M) or Lys-38 (P3K) (50 �M) for 10 –30 min at 37 °C before use.In select experiments, leukocytes were pretreated with 30 –300nM recombinant wild-type or mutant APC for 30 min at 37 °C.Samples were then washed once with PBS and subsequentlytreated with HBSS, PMA (10 nM), platelet secretome treatedwith PPACK, or platelet secretome for 3 h at 37 °C. Cell sampleswere then fixed and imaged.

Platelet adhesion assay to leukocytes and NETs

In select experiments, purified human leukocytes (2 � 106/ml) were plated on fibronectin-coated coverslips for 30 min at37 °C. Leukocytes were pretreated with APC (300 nM) for 30min at 37 °C prior to washing and stimulation with HBSS orPMA (10 nM) for 3 h at 37 °C. Samples were then washed oncewith PBS and incubated with autologous purified platelets (2 �107/ml) and allowed to adhere for 45 min at 37 °C. In selectwells, platelets were allowed to adhere to a fibronectin-coatedcoverslip in the absence of neutrophils. Samples were thenwashed once with PBS and fixed with 4% PFA followed by stain-ing, mounting, and imaging.

Image analysis

The fluorescent intensities of each image were adjustedbased on signals detected in leukocyte or platelet samples in theabsence of primary antibodies. Quantification of fluorescentimages was performed using a custom algorithm in MATLAB(MathWorks, Natick, MA). This algorithm first establishesglobal thresholding parameters based on a training set of fluo-rescence image data to then systematically quantify intensityprofiles across treatment conditions. The area of the total fieldof view is calculated to be 12510 �m2. Quantitative comparisonof treatment conditions was achieved by the normalization offluorescence intensities per image to determine the area ofDNA or respective fluorescent channels.

LDH activity assay

Primary human leukocytes (4 � 105/ml) were purified andincubated in solution with 300 nM APC for 30 min at 37 °C in a96-well plate format. In brief, samples were then incubated withHBSS or PMA (20 nM, final) and after mixing were incubatedfor 3 h at 37 °C. 10� lysis buffer was added to duplicate columnsand incubated for 45 min at 37 °C. Supernatant from all tripli-cate samples, including lysed samples and an LDH-positivecontrol were transferred to new 96-well plates prior to incuba-tion with reaction mixture for 30 min at 37 °C. The reaction wasstopped and absorbance was measured at 490 nm and 680 nm.Percent cytotoxicity was quantified as per the manufacturer’sinstructions (Thermo Fisher).

Neutrophil elastase activity assay

Primary human leukocytes (4 � 105/ml) were purified andincubated in solution with 300 nM APC or neutrophil elastaseinhibitors; CMK (475 �M) or Sivelestat (46 nM) for 30 min at37 °C in a 96-well plate format. Samples were then treated withvehicle, fMLP (10 �M, final) � cytochalasin B (5 �g/ml, final), or

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PMA (50 nM, final) for 2 h at 37 °C. Supernatants were collectedand incubated with substrate solution, protected from light for1.5 h at 37 °C. Fluorescence was measured at excitation 485 nm,emission 525 nm. Elastase was quantified as mU/ml using thestandard curve generated and in comparison to the neutrophilelastase-positive control as per manufacturer’s instructions(Cayman Chemical).

Nonhuman primate model of bacterial sepsis

Studies were reviewed and approved by the Institutional Ani-mal Care and Use Committee at Oklahoma Medical ResearchFoundation. In this established baboon (Papio anubis) model ofsepsis, animals (n � 4) were initially dosed with 3 mg/kg bolusof APC T � 10 min. At T � 0 min to T � 320 min, APC wasinfused at 16 �g/kg/min i.v. A lethal dose (1–2 � 1010 cfu/kg) ofE. coli, ATCC 12701 serotype O86:K61(B7) was infused intra-venously over 2 h (from T 0 to T � 120 min). The control group(n � 8) consisted in historical experiments where animals werechallenged with the same dose of bacteria under similar exper-imental conditions but not treated with APC. Blood sampleswere obtained at 0, 2, 4, 8, and 24 h and plasma samples wereisolated. MPO in plasma samples was determined using FluoroMPO (fluorescent myeloperoxidase detection kit) (Cell Tech-nology, Fremont, CA) as per the manufacturer’s instructions.

Statistical analysis

Two-way analysis of variance with Tukey’s post hoc correc-tion or one-way analysis of variance with Welch’s post hoc cor-rection was used to assess statistical significance among param-eters across multiple normally distributed cell parameters. pvalues of 0.05 or less were considered statistically significant.At least four images per treatment condition were obtainedand experiments were performed in triplicate. All values arereported at means � S.E. of the mean unless otherwise stated.

Author contributions—L. D. H. participated in the conception of thestudy, designed and performed experiments, analyzed data, andwrote the manuscript. C. P. contributed to experimental design andediting the manuscript. J. A. F. and X. X. performed experimentsand expressed and purified crucial APC mutants. A. M. contributedby performing experiments and editing the manuscript. N. A. T. andT. T. C. performed experiments and analyzed data. R. S. K. and F. L.performed and analyzed the animal study and participated in editingthe manuscript. A. G., F. L., J. H. G., and O. J. T. M. participated inthe conception of the study, designed experiments, and edited themanuscript.

Acknowledgments—We thank Jevgenia Zilberman-Rudenko for herstimulating comments and all the volunteers who kindly donatedtheir blood for this study.

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John H. Griffin and Owen J. T. McCartyKeshari, Nyiawung A. Taku, Tiffany T. Chu, Xiao Xu, András Gruber, Florea Lupu,

Laura D. Healy, Cristina Puy, José A. Fernández, Annachiara Mitrugno, Ravi S.in vivoactivation

andin vitroActivated protein C inhibits neutrophil extracellular trap formation

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