Dengue infection protein C

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Modification of the cytoprotective protein C pathway during

Dengue virus infection of human endothelial vascular cellsCarlos Cabello-Gutirrez1; Maria Eugenia Manjarrez-Zavala2; Alejandra Huerta-Zepeda1; Jorge Cime-Castillo1;Vernica Monroy-Martnez1; Benjamn Biruete- Correa3; Blanca H. Ruiz-Ordaz1

1Departamento de Biologa Molecular y Biotecnologa, Instituto de Investigaciones Biomdicas, Universidad Nacional Autnoma de Mxico,

Mxico, DF, Mxico; 2Departamento de Virologa, Instituto Nacional de Enfermedades Respiratorias, Mxico, DF, Mxico; 3Divisin de Obstet-

ricia UMAE, Hospital de Ginecologa y Obstetricia del IMSS Lus Castelazo Ayala, Mxico, DF, Mxico

SummaryDengue fever (DF) is the most prevalent arthropod-borne viraldisease of humans. No safe vaccine is available, there is no ex-perimental animal model and no specific treatment (antiviral)

for Dengue virus (DV) infection exists. The pathogenic mechan-isms of the severe forms of the disease, such as Dengue shocksyndrome (DSS) and Dengue haemorrhagic fever (DHF), inwhich endothelial damage is the pathognomonic sign, are notfully understood. Clinical observations have revealed significantabnormalities in the coagulation and inflammation systems, withincreased levels of soluble thrombomodulin (sTM) in the plasmaof patients with DHF/DSS (grade III or IV). Blood sTM was pro-posed as an early predictor of DSS during the febrile stage. How-ever, the role of the DV in endothelial injury during DSS is un-clear. Here, we present novel insights into the participation ofDV in the downregulation of the thrombomodulin-thrombin-

KeywordsDengue virus, endothelial cells, APC, thrombomodulin, PAR-1,EPCR inflammation, coagulation, cytoprotection

protein C complex formation at the endothelial surface, with areduction in activated protein C (APC). APC is the most impor-tant vasoprotective protein because it downregulates thrombin

generation (by the inactivation of procoagulant factors Va andVIIIa) and has anti-inflammatory, antiapoptotic, and barrier pro-tection properties. These biological functions of APC are associ-ated with the endothelial protein C receptor (EPCR) and pro-tease-activated receptor 1 (PAR-1) signalling pathways, whichlink the coagulation-inflammation responses. We found alter-ations in the antithrombotic and cytoprotective protein C path-ways during DV infection of human endothelial vascular cells,which may explain the vasculopathy observed during DHF/DSS.Clarification of the basic principles that underlie these pro-cesses has important implications for the design of new thera-peutic strategies for DHF/DSS.

Thromb Haemost 2009; 101: 916928

Wound Healing and Inflammation/Infection

Correspondence to:Blanca Hayd Ruiz OrdazDepartamento de Biologa Molecular y BiotecnologaInstituto de Investigaciones BiomdicasUniversidad Nacional Autnoma de MxicoApartado Postal 04510, Mxico, DF, MexicoTel.: +52 55 56 22 89 31, Fax: +52 55 56 22 92 11E-mail: [email protected]

Financial support:This study was supported in part by the PAPIIT and CONACYT programs.

Received: April 30, 2008Accepted after major revision: January 25, 2009

Prepublished online: April 3, 2009doi:10.1160/TH08-04-0271

Introduction

Dengue fever (DF) is the most important mosquito-borne viraldisease in tropical areas. The World Health Organization cur-rently estimates that there may be 50 million Dengue infectionsworldwide every year (1). In 2007 alone, there were more than890,000 reported cases of DF in the Americas, of which 26,000

cases were Dengue haemorrhagic fever (DHF)(1). Dengue virus(DV) infection is caused by any of the four DV serotypes (DV14), which can be transmitted to the human by the bite of the fe-male bloodsucking mosquitoesAedes aegypti andA. albopictus

(2). The clinical spectrum of the disease varies from a mild feb-

rile illness to more severe forms, such as DHF and Dengue shocksyndrome (DSS). DF is characterised by sudden fever, rash,headache, and non-specific signs and symptoms, such as myal-gias, arthralgias, and general weakness (2). DHF/DSS are diffi-cult to differentiate from DF in the acute phase. However, duringdefervescence, symptoms of circulatory failure appear abruptlyand shock or leakage manifestations may occur(2, 3). No effec-

tive strategies exist to prevent the progression of DHF because itspathogenic mechanisms are not fully understood. The haemos-tatic changes involve three main factors: thrombocytopenia,multiple defects in the coagulation-f ibrinolysis system, and vas-cular damage(increased permeability) (24). The vascular en-

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dothelium plays a determining role in the response to injury be-cause it functions as a regulatory interface during haemostasis(coagulation-fibrinolysis) and inflammation, and at the vascularendothelium barrier (5). Under normal conditions, endothelialvascular cells (EVC) promote anticoagulant properties by differ-ent mechanisms (6). The protein C (PC) anticoagulant pathway

provides the main control for the coagulation system (7). Thethrombomodulin-thrombin complex (TM-TB) plays a determin-ing role during the PC activation (APC) process (8, 9). TM is aconstitutively expressed glycoprotein present in high concen-trations in EVC, and is the thrombin-specific receptor (10, 11).APC is a multidomain plasma serine protease, which downregu-lates the generation of thrombin by the inactivation (proteolysis)of procoagulant factors Va and VIIIa (7). APC also has anti-in-flammatory, antiapoptotic, and cytoprotective properties, main-taining the endothelial barrier (12, 13). These biological func-tions of APC are associated with the endothelial protein C recep-tor (EPCR) and the activation of the protease-activated receptor(PAR-1) signalling pathways, which link the blood coagulation

and inflammation responses (5, 14).Under normal conditions or in response to minor injury, thevascular endothelium remains protected because TM sequestersthrombin, which generates adequate local levels of APC, confer-ring a cytoprotective effect (12). However,during profound en-dothelial injury (viral infection or inflammatory stimulus),thrombomodulin is released (soluble TM, sTM) and decreases atthe cell surface level, causing reduced APC generation and tissuedamage (15). Downregulation of the APC anticoagulant pathwaypromotes thrombosis and amplifies the inflammatory and apop-totic processes and EVC dysfunction (15). Recently, Buttep et al.(16) and Sosothikul et al. (4), have reported elevated levels ofsTM in the plasma of DHF patients as result of endothelial in-

jury. However, the role played by DV in the vasculopathy is un-known. In the present study, we evaluated the participation of theTM-TB complex in APC generation in human umbilical vein en-dothelial cells (HUVEC or EVC) infected with DV. We also as-sessed the APC cytoprotective regulatory pathways that involvedPAR-1 and EPCR receptors on endothelial vascular cells in thepresence of DV.

Materials and methods

Viral isolatesTwo different DV isolates of serotype 2 were used, one from a pa-tient with a fatal case of DHF (DHF-DV/D79 isolated in 1979 in

Thailand. Asian genotype) and the other from classical DF (DF-DV/D2M is a Mexican strain. American genotype). Bothsamples were evaluated in terms of their virulence in mice (17),and were designated DHF-DV (elevated virulence) and DF-DV(low to medium virulence), respectively. Both samples werekindly donated by Dr. Duane Gubler from the Center for DiseaseControl in Fort Collins, CO, USA.

Cell lineLLC-MK2 cells of the African green monkey kidney (AmericanType Culture Collection) were used for DV amplification, ti-tration, and purification. For propagation, cells were infectedwith DHF-DV or DF-DV at a multiplicity of infection (MOI) =

0.1. They were incubated at 37C with 5% CO2 until their cyto-pathic effect (CPE) was above 90%, then harvested in cryotubescontaining 0.5% albumin in minimum essential medium, andstored at 70C.

DV titration by lytic plaque assay

LLC-MK2 monolayers were inoculated with 100 l of serial logdilutions of DF-DV or DHF-DV in serum-free medium (100 to109) in duplicate, and incubated for 2 hours (h) at 37C with 5%CO2. The viral inoculum was removed with washing and the cellswere overlayed with 1.5 ml of incomplete medium plus 2.5%methylcellulose. Cultures were incubated until the CPE wasabove 90% (68 days), and then stained with 1% crystal violet.

Establishment of EVC culturesHUVEC were isolated as described previously by Olsen (18).Briefly, the umbilical cord vein was cannulated, washed with sa-line solution plus HEPES and 5 ml of 0.075% trypsin-versenesolution, and incubated for 10 minutes (min) at 37C. The cells

were centrifuged and resuspended in M199 medium containing10% fetal bovine serum, 100 mg/ml porcine heparin, 50 U/ml penicillin/streptomycin, and 25 g/ml vascular endothelialgrowth factor. The cells used in all experiments corresponded tothe third consecutive passage and were verified by their cobble-stone morphology and the presence of the von Willebrandantigen (by immunofluorescence assay). Cell viability was de-termined with trypan blue stain.

Detection of TM in DV-infected EVC by flow cytometricassayTM at the membrane surface was determined by flow cytometricassay (fluorescence-activated cell sorting [FACS]). HUVEC

were infected at different MOI (0.01, 0.1, or 1.0) with DF-DV orDHF-DV for 24 or 48 h or stimulated with 10 ng/ml tumour ne-crosis factor- (TNF-,positive control for reduced TM), whichinduces the downregulation of TM at endothelial cell surfacelevel (19). Cultures were washed in phosphate-buffered saline(PBS), detached with 0.5 M EDTA-PBS, and resuspended inPBS (1 ml) with anti-TM antibody diluted 1:100. After 1 h incu-bation (4C), the samples were centrifuged (3 min, 1,300 rpm)and washed three times with buffer A (150 mM NaCl, 2.5 mMCaCl2, 5 mg/ml bovine serum albumin [BSA] 20 mM Tris-HCl[pH 7.4]). Secondary fluorescein isothiocyanate (FITC)-con-jugated anti-human IgG antibody (Santa Cruz Biotechnology,Santa Cruz, CA, USA) was added, incubated for 1 h at room tem-

perature (RT), and washed with the same buffer. The pellet wasresuspended in cytometry fluid and analysed by FACS.

TM determination by Western blot assayHUVEC membrane proteins were extracted as follows. Briefly,DHF-DV (MOI = 1.0)-infected HUVEC were concentrated bycentrifugation at 3,000 rpm (10 min) and washed with PBS. Thecells were resuspended in RSBNP-40 (1.5 mM MgCl2, 10 mMTris-HCL, 10 mM NaCl, 1% Nonidet P-40) in the presence ofprotease inhibitors. Nuclei and debris were removed by centrifu-gation. The amount of protein was determined with the Bradfordmethod(20). Samples were separated by 10% sodium dodecylsulfate (SDS)-polyacrylamide gel electrophoresis (PAGE),


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transferred to nitrocellulose membranes, and incubated with aprimary antibody (1:1,000) overnight at 4C (anti-TM; SantaCruz Biotechnology). Bound antibody was detected using a per-oxidase-conjugated secondary antibody (1:2,000) in 10 mMTris-HCl (pH 7.6), 150 mM NaCl, and 0.05% Tween 20 in Tris- buffered saline (TBST) for 1 h at RT. The membranes were

washed and the proteins were detected with a western blot lumi-nol reagent.

Identification of sTM in the supernatants ofDV-infected EVCThe presence of free thrombomodulin in the supernatants (SN)of HUVEC cultures infected withDHF-DV (MOI = 1.0) was de-termined after 36 and 48 h by Western blot assay, as previouslydescribed. The samples were centrifuged and concentrated inYM-3Microcon tubes (Millipore, Billerica, MA, USA).

Colocalisation of PAR-1 and EPCR by confocalmicroscopy

We evaluated the colocalisation of EPCR and PAR-1 at cell sur-face level in DV-infected HUVEC. Confluent monolayers werecultured on silane multiwells slides and processed for double im-munofluorescence. Briefly, DV-infected cells with DHF-DV orDF-DV (MOI = 1, 48 h) or negative controls, were fixed (2%paraformaldehyde), washed (3 times PBS-BSA 1%) and incu-bated 1.5 h at room temperature (separately) with specific anti-PAR-1 and EPRC antibodies (1:100; Santa Cruz). Slides werewashed (3 times PBS-BSA1%) and incubated with secondaryantibodies to reveal immunostaining. For EPCR a monoclonalIgG coupled to TxR was used (1:100; Vector, Burlingame, CA,USA) and a fluorescein-conjugated monoclonal anti-IgG(1:100) was employed for PAR-1. Cross-reactivity was excluded

by negative control that showed no immunostaining (non-in-fected cultures in presence of the same solutions). Samples werewashed and covered with fluorescent mounting medium (Vectas-hield, Vector) and examined with an epifluorescence microscope(Nikon Eclipse) equipped with filters for FITC and TxR amongothers. The samples were further analysed with a confocal LSM5 PASCAL Zeiss microscope equipped with argon/krypton laser.Images were collected with 1.30 numerical aperture (NA) oil-immersion (100 X) objective. To avoid bleed-through, FITC la-bels were excited with the 488 line and emitted light was band-passed with 520550-nm filter and TxR labels were excited withthe 543 line and emitted light was long-passed with 650-nmfilter. Confocal images were obtaining using two separate photo-

multiplier channels, either concurrently or in separate runs. Im-ages were separately projected and merged using a pseudocolordisplay showing green for FITC, red for TxR and yellow for co-localisation. Confocal images were analysed with the help of theassistant confocal programs IMARIS 6.0 BITPLANE andIMAGE PRO.

Protein C activation assay in DV-infected endothelialcellsThe amidolytic activity of APC was evaluated using a chromo-genic assay, as described by Cadroy (21). The APC assay wasperformed using confluent monolayers (2 105 cells/well) in24-well culture plates infected at different MOI (0.01, 0.1, or 1.0)

of DF-DV or DHF-DV for 48 h. As the positive control, HUVECwere stimulated with 10 ng/ml TNF-. The cultures were washedwith buffer A, to which was added 100 l of buffer A plus1.5 U/ml human thrombin, 150 nM/l human PC (Sigma, St.Louis, MO, USA), and 5 mM CaCl2 (Sigma). The cells were in-cubated for 2 h at 37C. We collected 50 l of the culture super-

natants and added 450 l of APC chromogenic substrate (S2266,2 mM Chromogenix) in the presence of 2.5 U/ml hirudin(Sigma) to inhibit any free thrombin. After 30 min, the reactionwas stopped with 200 l of 50% (vol/vol) acetic acid. The amid-olytic activity of APC was read at 405 nm.

Phosphorylation of mitogen-activated protein kinases(MAPK1/p38 and MAPK3/ERK1/2)Endothelial cells stimulated with TNF- or infected with DHF-DV (MOI = 1.0) were lysed with cold lysis buffer (50 mM Tris-HCl [pH 7.4], 150 mM NaCl, 1 mM EGTA, 0.25% sodium oxy-cholate, 1 mM Na3VO4, 1 mM NaF, 1 mM of phenylmethylsulfo-nyl fluoride, 1% Triton X-100, 1 g/ml protease inhibitors) for

10 min. The samples were centrifuged (14,000 rpm for 3 min) at4C and resuspended twice in sample buffer, boiled for 10 min,and analysed by SDSPAGE (10%). The proteins were trans-ferred (3.5 h) to 0.45 m nitrocellulose membrane and blockedfor 1 h at RT, using TBST (vol/vol) with 5% fat-free powderedmilk as the blocking solution. The membranes were washed withPBS and 0.05% Tween 20 (PBST) and incubated overnight at4C in the presence of 1:1,000 primary antibody (anti-phospho-ERK, anti-ERK, anti-phospho-p38, or anti-PAR-1 [ATAP];Santa Cruz Biotechnology). The bound antibody was detectedwith a peroxidase-conjugated secondary antibody (Zymed Lab-oratories, South San Francisco, CA, USA) diluted at 1:2,000with BSA (2.5 mg/ml) in PBS, for 1 h at RT. The membranes

were washed and the proteins detected with a Western blot lumi-nol reagent. We performed a parallel assay to evaluate anychanges in the phosphorylation of MAP kinases (MAPK1 or p38and MAPK3 or ERK1/2) in DV-infected EVC pretreated for 2 hwith either a specific inhibitor of ERK phosphorylation (PD98059, 10 nM) at different times (1, 4, or 6 h), a specific p38 ki-nase inhibitor (SB-203580, 1 M), APC (10 ng/ml), a specificanti-PAR-1 antibody, or recombinant human interleukin 8 (IL-8,10 ng/ml). The samples were analysed by Western blotting.

IL-8 determinationIL-8 levels were determined in the supernatants of DV-infectedHUVEC (MOI = 1.0, 0.1, or 0.01) by enzyme-linked immuno-

sorbent assay (ELISA). Conditions were optimised titrating therecombinant human IL-8 (rh IL-8) as described by Kittigul (22).Briefly, a microtiter plate was covered with 100 l (10 g/ml) ofanti-IL-8 monoclonal antibody (R&D Systems Inc., Minneapo-lis, MN, USA) in bicarbonate buffer (pH 9.6) and incubatedovernight at 4C. The plate was washed with 0.05% PBST (pH7.4) and blocked with 200 l of BSA-PBS for 2 h at RT. Thesamples were washed in PBST and 100 l of the following sub-stances was added: recombinant human IL-8 protein or thesupernatant (SN) of DV-infected cells; the SN of uninfectedHUVEC, the SN of EVC stimulated with 10 ng/ml TNF-, and/or the SN of EVC pretreated with the ERK inhibitor PD98059.The cultures were incubated for 1.5 h at 37C, washed, and then


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100 l/well of biotinylated anti-IL-8 antibody (5 g/ml) wasadded and the cells were incubated for 1.5 h at 37C. After thesamples had been washed three times, 100 l/well of streptavi-din-peroxidase (Sigma) diluted 1:5,000 was added and the cellswere incubated for 1 h at 37C, followed by three rounds ofwashing. The color was developed for 15 min using 100 l of tet-

ramethylbenzidine substrate (Sigma). The reaction was stoppedwith 50 l/well of 0.05 M sulfuric acid. The optical densitieswere determined in an ELISA reader at 450 nm, at least fourtimes. A parallel assay was performed on cultures that had beenpretreated with 10 ng/ml APC and/or a specific anti-PAR-1 anti-body for 2 h before they were infected with DHF-DV (MOI =1.0).

Propidium iodide analysisDHF-DV (MOI = 1.0)-infected HUVEC, DHF-DV-infectedHUVEC pretreated with 10 ng/ml APC, and HUVEC stimulatedwith TNF- (10 ng/ml) were fixed with 4% paraformaldehydefor 1 h at RT, washed with PBS (pH 7.45), and incubated with

0.5 mg/ml RNase. The samples were stained with propidium io-dide (50 mg/ml; Sigma) for 15 min at RT, and washed with PBS.The samples were analyzed under a fluorescence microscope.

Vascular permeability assayAn in vitro vascular permeability assay (Chemicon KitECM640) was performed using 24-well culture plates contain-ing culture inserts with symmetrical pores and polyethylenemembranes, which permit the diffusion of molecules. Briefly,HUVEC were seeded at 2 105 cells per collagen-coated insertand grown to confluence, and then infected with DF-DV orDHF-DV (MOI = 1.0) or stimulated with TNF- (positive con-trol) for 48 h. FITC-dextran was then added to the samples for

1 h. Parallel assays were performed using pretreated inserts with10 ng/ml IL-8, 10 ng/ml APC, specific MAPK inhibitors(PD98059 for ERK1/2 and SB203580 for p38), or specific anti-PAR-1 antibody. The extent of cell permeability was determinedby measuring the fluorescence of the solution in each well at 492nm (in triplicate).

Statistical analysisData from at least three independent experiments assayed in trip-licate were expressed as means standard deviation (SD) andevaluated with Students t-test and the Mann-Whitney U test,using the Statistical (version 6) program. The significance levelwas set at p< 0.05.

Results

Detection of TM in DV-infected EVCTM was detected on the surfaces of DV-infected HUVEC (MOI= 0.01, 0.1, or 1.0) by FACS. TM expression was correlated withthe mean fluorescence intensity (MFI). As a positive control,HUVEC were stimulated with 10 ng/ml TNF-. We observedthat DV caused the downregulation of TM in EVC, as shown inFigure 1AD. TM expression decreased to 85% (Fig. 1A and B,pink and purple bars) in the presence of DHF-DV at 48 h post in-fection (p.i.; MOI = 1.0 or 0.1) and this effect was more promi-nent than that observed in the presence of TNF- (35%; green

bar, Fig. 1B and D). HUVEC infected with DF-DV showed a28% reduction in MFI (Fig. 1C and D, pink bar) in the presenceof the highest dose (MOI = 1.0). To confirm these data, we per-formed a parallel immunohistochemical assay of DF-DV orDHF-DV (MOI = 1.0)-infected EVC for 48 h. We observed thepresence of constitutive TM at the surfaces of uninfected cells

(Fig. 1E, panel 1). However, the maximum reduction in TM wasobserved in DHF-DV-infected HUVEC (Fig. 1E, panel 3). Thesesamples presented lower amounts of TM than the positive control(TNF- Fig. 1E, panel 2). Nevertheless, in presence of DF-DV(Fig.1 E, panel 4) the cell surface TM amount was similar to theconstitutive TM (Fig. 1 E, panel 1). These data correlate with ourcytofluorometric results. To evaluate whether the reduction inTM levels was the result of deleterious effects of DV infection oncell viability, we performed a cell viability assay using trypanblue staining of HUVEC cultures infected with DF-DV or DHF-DV isolates (MOI = 0.01, 0.1, or 1.0) for 48 h. EVC viability(Fig. 1F) was around 9096% in all samples (infected andnegative and positive control cells), demonstrating that there was

no cell death attributable to Dengue cytotoxicity.

Evaluation of TM release and PAR-1/EPCRcolocalisationThe possible liberation of TM (sTM) from the cellular mem-brane to the culture supernatants of DHF-DV-infected HUVEC(MOI = 1.0) was evaluated with a Western blot assay. We exam-ined the proteins present in the membrane extracts and those inthe supernatants at different times (6, 12, 24, and 48 h p.i.). Theconstitutive form of TM was detected in the uninfected cells (Fig.1G, upper panel, lane 1), but the amount of this protein graduallydecreased in DV-infected cultures as the period of infection in-creased (Fig. 1G, upper panel, lanes 25). We found no free TM

in the SN of the uninfected cells (Fig. 1G, upper panel, lane 6).The major band detected with the monoclonal antibody cor-responded to a protein of approximately 96 kDa, but in somecases we detected a minor dimeric isoform of TM. Figure 1G(lower panel, lanes 2 and 3) shows that DV infection caused therelease of TM into the culture supernatant at 36 and 48 h p.i. NoTM was present in the SN of the uninfected cells (Fig. 1G, lowerpanel, lane 1). As previously discussed, low levels of TM result indown regulation of APC causing proinflammatory diathesis(12), as in addition to its well-studied anticoagulant effect, APCalso elicits potent cytoprotective and antiinflammatory re-sponses when APC forms complex with endothelial protein C(EPCR) that acquires a different specificity, thus activating

PAR-1. Therefore, we evaluated the presence of PAR-1 andEPCR at endothelial cell surface in presence of DV. We observedby immunolocalisation (using double label and confocal micro-scopy) that EPCR and PAR-1 were colocalised (yellow bandFig.1 I-J, L-M) at cell surface level in DV-infected HUVECs(Fig. 1 H-P). The relative immunofluorescence intensity washigher in presence of DHF-DV than in DF-DV infected cells.

APC activity assay in DV-infected EVCWe also evaluated if downregulation of APC modifies the amid-olytic activity of APC in HUVEC infected with DF-DV or DHF-DV with the help of a chromogenic assay. APC activity was re-duced in the presence of both DV isolates at 48 h p.i. at all doses


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(MOI = 1.0, 0.1, and 0.01) compared with that of the uninfectedcells, which were considered to have 100% amidolytic activity(Fig. 2A). The APC activity in DHF-DV-infected EVC decreased

70% at the highest dose (Fig.2E), which was lower than that inthe TNF--treated cells (Fig. 2B). APC activity in DF-DV-in-fected cells decreased 40% at the highest dose (Fig. 2H). Thesedata suggest that DV infection affects the formation of the TM-TB-PC complex and APC generation, mainly in the presence ofDHF-DV, which is consistent with the cytofluorometry results.

Anti-inflammatory effect of APC in DHF-DV-infectedEVCIn addition to its anticoagulant activity, APC has anti-inflamma-tory properties and downregulates the expression of inflamma-tory interleukins, such as IL-8. We evaluated the consequence ofdecreased APC activity on IL-8 production in endothelial cells in-

fected with different doses of DF-DV or DHF-DV at 48 h p.i,using an ELISA assay. We detected high levels of IL-8 at all dosesof DHF-DV in infected cells (Fig. 3A, bars 35) when compared

with that in uninfected cultures and negative control (Fig. 3A, bar1 and 3) and with the positive control (TNF-, 115.61 pg/ml. Fig.3, bar 2). At MOI = 1.0, the amount of IL-8 was 190 pg/ml; atMOI = 0.1, it was 160 pg/ml; and at MOI = 0.01, it was 150 pg/ml. In cultures infected with high doses of DF-DV (MOI = 1.0),the amount of IL-8 was 28.33 pg/ml (Fig. 3A, bar 9). Cytokine

production in the presence of DHF-DV was higher than that in thepositive control. To evaluate the participation of APC in cytopro-tection, a parallel assay was performed using DHF-DV-infectedHUVEC (MOI = 1.0) that had been pretreated with 10 ng/ml ofrecombinant APC (Fig. 3B). IL-8 production decreased by asmuch as 90% (Fig. 3B, bar 5) with respect to that in DHF-DV-in-fected EVC in the absence of APC (Fig. 3B, bar 4). The same ef-

Figure 1: Detection of constitutive and free TM during DV in-fection of EVC. AD) Determination of constitutive TM in HUVEC byflow cytometry. The histograms (A) and bars (B) correspond to the fol-lowing samples: negative control (red); positive control (green), HUVECcultures stimulated with 10 ng/ml TNF-; uninfected cells (black); DHF-DV-infected EVC at MOI = 0.01 (blue), at MOI = 0.1 (purple), and atMOI = 1.0 (pink). Figure 1C and D correspond to DF-DV-infectedHUVEC. Results are the means SD of at least three independent ex-periments assayed in triplicate. * Indicates p-values when samples in thepresence of DV were compared with uninfected cells (p< 0.05). E) TM

detection by immunohistochemistry assay. Panel 1 shows the constitut-ive form of TM at the cell surfaces of uninfected HUVEC cultures. Panel2 shows the decrease in TM in HUVEC pretreated with 10 ng/ml TNF-.TM was downregulated in DHF-DV-infected EVC (MOI = 1; panel 3) andDF-DV-infected HUVECs (MOI=1; panel 4) show similar TM amounts ofconstitutive TM. The data shown are representative of another three ex-periments that produced similar results. F) HUVEC viability assay. En-dothelial cells were pretreated with 10 ng/ml TNF- (gray bar) or in-fected with DF-DV or DHF-DV at MOI = 0.01, 0.1, or 1.0. Open bar

shows the viability of uninfected HUVEC cultures. SD was calculated foreach sample. G) Upper panel: TM identification in HUVEC cultures byWestern blot assay. Uninfected cells (lane 1). DHF-DV-infected HUVEC(MOI = 1.0) at different time intervals (6, 12, 24, and 48 h p.i., lanes 25,respectively) and the culture supernatants (lane 6 and Fig. 1G lower).Lower panel: sTM in the SN of DHF-DV-infected cells at 36 and 48 h p.i.(lanes 2 and 3, respectively). Uninfected cells (lane 1). The data shownare representative of another three assays with similar results. H-J) Co-localisation of PAR-1 and EPCR receptors by confocal microscopy. Thecolocalisation of PAR-1 and EPCR was represented by the yellow zone.

Panel H shows uninfected cells. Panel I colocalisation in DF-DV infectedHUVEC. Panel J colocalisation in DHF-DV infected HUVEC. The datashown are representative of another three experiments that producedsimilar results. K-P) Localisation of PAR-1 and EPCR by immunostaining.Panel K uninfected cells. Panel L PAR-1/EPCR immunolocalisation in DF-DV infected cells. Panel M PAR-1/ EPCR immunolocalisation in DHF-DVinfected cultures. Panels N-P show independent immunostaining ofPAR-1 and EPCR. The fluorescence values were given in arbitrary units(p


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fect was observed in TNF--stimulated cultures pretreated withAPC (Fig. 3B, bars 2 and 3). However, the poor response of IL-8in presence of the avirulent strain does not permit to evaluate theAPC activity (Fig. 3B, bars 6 and 7).

The mechanisms involved during the induction of IL-8 inDV-infected EVC have remained elusive. In other viral haemor-rhagic fevers, such as Ebola or Bluetongue, and in septic shock,the participation of MAPKs in this process has been established(2325). Therefore, we assessed whether IL-8 expression is

under the control of ERK1/2 kinases in the vascular endothe-lium, using Western blot and ELISA analyses. ERK1/2 acti-vation in DHF-DV-infected EVC (MOI = 1.0) was determined.Figure 4A shows that MAPK ERK1/2 was phosphorylated in thefirst 5 min and the signal gradually increased until 90 min, al-though at 180 min the signal began to decrease. Figure 4B showsthe presence of constitutive ERK in HUVEC cultures. We alsoevaluated whether IL-8 production is under phospho-ERK con-trol, using an inhibitory IL-8 ELISA in the presence of the spe-cific ERK1/2 inhibitor PD98059 at different time intervals (1, 4,and 6 h) before DV infection. Figure 4C shows that IL-8 produc-tion was clearly inhibited in DHF-DV-infected cultures pre-treated for 6 h with PD98059, and a concentration of only 39 pg/

ml IL-8 was detected. To confirm these results, a parallel West-ern blot assay was performed. HUVEC cultures preincubated for6 h with PD98059 showed the lowest ERK phosphorylation (Fig.4C, top square I). We then examined the possible activation ofp38 during DV infection (DHF-DV, MOI = 1.0) of endothelialcells, using a Western blot assay. Figure 5A (lane 3) shows that,during DV infection, p38 was phosphorylated in the first 30 min,as in the positive control (TNF-, lane 2), and as in the presenceof IL-8 (lane 4). To evaluate the specificity of these data, a paral-lel assay was performed in the presence of the p38-specific in-hibitor, SB203580 (10 M). The downregulation of p38 acti-vation was evident in both the DHF-DV-infected cells and theEVC treated with 10 ng/ml IL-8 (Fig. 5A, lanes 5 and 6). A recent

study has indicated that genes that are upregulated by APC arealso induced by agonist peptides of PAR-1 (26). Therefore, wetested whether the APC response is mediated by the PAR-1MAPK pathway, using Western blotting and ELISA. We firstexamined the possible activation of the MAPKs ERK1/2 in thepresence of APC. Figure 5B (lane 2) shows the phosphorylationof ERK1/2 in HUVEC cultures pretreated with APC (10 ng/ml).A parallel assay was performed using specific anti-PAR-1 anti-body. Phospho-ERK1/2 was downregulated in the presence of

anti-PAR-1 antibody, as shown in Figure 5B, lane 6. In a similarassay, phospho-ERK1/2 was determined in DHF-DV-infectedEVC (MOI = 1.0, Fig. 5B, lane 4) in the presence of specific anti-PAR-1 antibody. A significant reduction in phospho-ERK1/2was also observed (Fig. 5B, lane 5). In a parallel IL-8 blockingassay (using specific anti-PAR-1 antibody) in DHF-DV-infectedEVC (MOI = 1.0), IL-8 production was also downregulated inthe presence of anti-PAR-1 antibody (Fig. 5C, lanes 36). Ourdata suggest that the phosphorylation of the MAPKs ERK1/2 in-duced by APC may be PAR-1 dependent, in a similar way to IL-8 production. These novel and important findings indicate themain role of the MAPKs p38 and ERK1/2 in the APCPAR-1signalling pathways regulating IL-8 induction, during DV infec-

tion of endothelial cells.

Role of APC and ERK1/2 or p38 MAPKs in EVCpermeabilityBased on the data presented here, we also explored the involve-ment of APC and ERK1/2 or p38 MAPKs in the endothelial bar-rier function during DV infection of HUVEC cultures (DHF-DV,MOI = 1.0), using a vascular permeability assay (Chemicon), asdescribed in Materials and methods. Endothelial cells treatedwith only culture medium, were used as the positive control formonolayer integrity (Fig. 6B) in which no cell permeability wasobserved. These samples represent the background FITC-dex-tran values. Inserts without cell monolayers were considered the

Figure 2: APC activity assay during DVinfection of EVC. Uninfected HUVEC (con-trol, A). Cultures pretreated with 10 ng/mlTNF- (B). DHF-DV-infected cells at differentMOIs (0.01, 0.1, and 1.0 as bars C, D, and E,respectively). DF-DV-infected EVC at MOI =0.01, 0.1, and 1.0 (bars F, G, and H, respect-ively). Values represent means SD of at leastthree independent experiments assayed in trip-licate. * Indicates p-values (p< 0.05) when

samples in the presence of DV were comparedwith uninfected cultures.


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100% permeability control (Fig. 6A). The DHF-DV-infected cul-tures displayed a notable increase in cell permeability (Fig. 6F),at a level similar to that of the positive controls treated withTNF- or IL-8 (Fig. 6C and D). However, in the DHF-DV-in-fected cells or IL-8-stimulated cultures pretreated with eitherAPC (10 ng/ml, panels E and G) or the specific inhibitors of theMAPKs p38 and ERK1/2 (Fig.6 J and K), a partial protection interms of the endothelial barrier was apparent. The poor response

of DF-DV isolate with respect to the permeability effect dont letus to evaluate the APC outcome (Fig.6 H and I). Our data supporta partial barrier protective role for APC.

Assay of APC cytoprotection during DV infection ofEVCThe protective role of APC during tissue injury has been wellstudied (7, 1213, 2730). We evaluated the possible cytoprotec-

Figure 3: Cytoprotective effect of APCon DV-infected endothelial cells. Wemeasured IL-8 in the presence of DF-DV- orDHF-DV-infected HUVEC (MOI = 0.01, 0.1,and 1.0), as shown in Figure 3A (lanes 49).Figure 3A lane 3 show the IL-8 production inpresence of UV-DV. HUVEC stimulated with 10ng/ml TNF- were used as the positive control(lane 2). In this figure, * indicates the p-value(p< 0.05) when samples in the presence of DVwere compared with uninfected cells. We thenassayed the cytoprotective effects of APC inTNF--treated or DV-infected EVC (MOI = 1)pretreated (2 h) with 10 ng/ml APC (Fig. 3B,lanes 3, 5 and 7). * Indicates p-values (p< 0.05)when samples pretreated with 10 ng/ml APCwere compared with non-pretreated cultures(lanes 2, 4 and 6).


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tive capacity of APC in DHF-DV-infected HUVEC (MOI = 1.0),using propidium iodide staining of proapoptotic nuclei. Culturesinfected with DHF-DV (MOI = 1.0) displayed proapoptotic nu-clei, with high levels of propidium iodide accumulation (Fig. 7C)similar to those found in the presence of 10 ng/ml TNF- (posi-tive control). However, in the DHF-DV-infected cells pretreatedfor 72 h with 10 ng/ml APC before DV infection, a significantprotective effect was observed as a reduction in the numbers ofproapoptotic nuclei (Fig. 7F). A similar effect was observed inHUVEC cultures stimulated with TNF- and pretreated for 72 h

with 10 ng/ml APC (Fig. 7E). We found no apoptotic nuclei in thenegative control (Fig. 7A) or in the presence of UV inactivatedDHF-DV (Fig. 7D).

Discussion

The pathogenesis of DHF/DSS has been explained by two the-ories. Leon Rosens hypothesis (3132) is based on the virulenceof DV isolates and Halsteads theory on immunopathogenesis inthe course of a secondary infection (33). Both hypotheses aresupported by epidemiological and experimental data, both ap-pear to represent different aspects of the same phenomenon, andthey are not mutually exclusive. Both are probably valid but un-

fortunately there are no good animal models of DHF/DSS,which makes studies of their pathogenesis difficult to interpret.DHF/DSS are determined by multiple factors, such as the hostsimmunological status, host age, genetics, intercurrent infections,and DV virulence among others (epidemiological factors).

Current hypotheses are insufficient to explain some clinicalmanifestations, such as the thrombocytopenia and haemocon-centration caused by endothelial activation (damage). Vasculo-pathy is the result of endothelial dysfunction, caused by both, di-rect (viral factors) and indirect (host factors) mechanisms, that

ultimately targets vascular endothelium (making it battlefield)leading to severe disease. Unquestionably, DV can infect en-dothelial cells in vitro (3739), which have been used widely tostudy the pathophysiological changes that occur during DV in-fection, to answer questions raised in patients with DHF/DSS.Most of the DHF/DSS pathogenic mechanistic information hascome from in-vitro experiments and remains to be translated tohuman patients. However, these in-vitro experiments have gener-ated useful knowledge about the mechanisms that may be occur-ring intracellulary.

Because strong and consistent clinical evidence suggests thatvascular activation (or damage) is involved in the pathophysiol-ogy of DHF/DSS, many groups have demonstrated data ex vivo

Figure 4: MAPK ERK1/2 activation in DV-infected EVC. MAPK (ERK 1/2) phosphory-lation (p) was detected by Western blot assayin the presence of DHF-DV. ERK activationwas evident in the first 5 min and gradually in-creased with increasing infection time (Fig. 4A).Maximum p-ERK was observed at 90 min, andit started to decrease at 180 min. Lane 1shows the uninfected HUVEC (Ctrl). Consti-tutive ERK1/2 (Fig. 4B). Figure 4C shows theeffects of the ERK1/2-specific inhibitorPD98059 (preincubated for 1, 4, or 6 h) onIL-8 production during DHF-DV (MOI = 1.0)infection of EVC. * Indicates the p-values whensamples in the presence of the ERK1/2 in-hibitor PD98059 were compared with DHF-DV-infected cells in the absence of inhibitor(p< 0.05).


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(3436), in vitro (32, 3739) andin vivo (4042) for the EVCparticipation. Therefore, Bonner and OSullivan (37) proposedthe use of EVC as an appropriate model for studying DSS. Ac-cordingly, we employed de HUVEC monolayers as our model.Here, we present novel insights into the participation of DV inendothelial cell modifications of the cytoprotective protein Cpathway. Inflammation-coagulation, apoptosis, and barrier pro-tection are essential parts of the hosts defensive response duringtissue damage (5) and have several convergent points of interac-tion (crosstalk) via the protease-activated receptors (PARs).PARs are molecules coupled to G-proteins, which are involved inregulatory signalling pathways (14, 26, 29). The first link be-tween these processes is mediated by the vascular endothelium

(5), which localises and promotes the diverse biochemical trans-formations involved in the cytoprotective APC pathway (PC ac-tivation, APC anticoagulant activity, anti-inflammatory effect,antiapoptotic response, and endothelial barrier stabilisation).Because of its pleiotropic activities, APC has a potential role inthe treatment of complex disorders, including sepsis, thrombo-sis, and ischemic stroke among others (29, 30). Therefore, weevaluated the basic mechanisms that could regulate these pro-cesses during dengue virus infection of EVC. We examined theTM-TB-PC complex at the endothelial surface (which plays akey role in APC generation) in the presence of DV. Recently, But-tep (16) and Sosothikul (4) have reported elevated sTM levels inthe plasma of patients with DF during the toxic phase of the dis-

ease, suggesting damage to (and specific activation of) vascularendothelial cells. Patients with DSS (DHF grade III or IV) showhigher concentrations of sTM than those of patients with DF orDHF grades I and II. In both studies, sTM levels correlated withthe severity of the disease.

Blood sTM was proposed as an early predictor of DSS duringthe febrile stage (16). However, the role of DV in the devel-opment of the vasculopathy that presents during DSS is unclear.We observed that, in the presence of DHF-DV (MOI = 1.0), theamount of TM at the endothelial surface was reduced to 85%relative to that of uninfected cells (Fig. 1A and B). This effectwas more prominent than those in cultures pretreated withTNF- (positive control). The DF-DV-infected HUVEC (MOI =

1.0) showed a 28% reduction in MFI in the cytofluorometricanalysis, and these data are similar to the TNF- response (Fig.1D). We demonstrated that the reduction in TM at the EVC sur-face is associated with the presence of DV, because the surfaceTM gradually decreased as the infection proceeded (Fig. 1G,upper panel). In agreement with this, the level of sTM was elev-ated in the SN of DV-infected cultures, and this was more evidentat 36 and 48 h p.i. (Fig. 1G, lower panel). These data stronglysuggest the participation of DV in EVC alteration and may ex-plain the presence of sTM in the plasma during the febrile phasein patients at risk of DSS, as reported by Buttep (16).

TM is a cell-surface receptor that plays a critical role in en-dothelial anticoagulant activity through its cofactor function

Figure 5: Role of APC, PAR-1, and

MAPKs (p38 and ERK 1/2) in the inflam-matory response of endothelial cells inthe presence of DV. A) Activation of p38during DV infection of EVC (lane 3) or in thepresence of TNF- (lane 2) and IL-8 (lane 4).We evaluated the specificity of these data in aparallel assay in the presence of a p38-specificinhibitor (lanes 5 and 6). B) Activation ofMAPKs ERK1/2 in EVC pretreated with 10 ng/ml APC (lane 2), which is similar to the acti-vation in DHF-DV-infected HUVEC (lane 4) orin cultures stimulated with TNF-.However, adownregulation of p-ERK1/2 was observed inthe presence of DV or in APC-stimulated cul-tures that had been pretreated with anti-PAR-1

specific antibody (lanes 5 and 6). The resultspresented here are representative of anotherthree experiments with similar results. C) Pro-duction of IL-8 in DHF-DV-infected culturespretreated with specific anti-PAR-1 antibody.The results in Figure 5C are the means SD ofat least four independent experiments assayedin triplicate. * Indicates the p-value (p< 0.05)when EVC infected with DV and pretreatedwith anti-PAR-1 antibody were compared withsamples infected with DV but in absence ofanti-PAR-1 antibody.


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Figure 6: Assay of APC participation in endothelial barrier pro-tection. We evaluated the participation of APC and MAPKs (ERK1/2and p38) in the endothelial barrier function during DV infection of EVC,using a vascular permeability assay as described inMaterials and methods.Figure 6 (bar B) shows HUVEC in the presence of culture medium only,which represents the monolayer integrity of the positive control. The in-

serts with no cell monolayer (bar A) represent the 100% permeabilitycontrol. The percentage EVC permeability in the presence of: TNF-(bar C), IL-8 (bar D), APC+IL-8 (bar E), DHF-DV MOI = 1.0 (bar F),

APC+DHF-DV (bar G), DF-DV (barH), APC+DF-DV (bar I),SB203885+DHF-DV MOI = 1.0 (bar J), or PD98059+DHF-DV MOI =1.0 (bar K) are shown. The data are representative of three independentexperiments assayed in triplicate, and are shown as means SD. * Indi-cates the p-value (p< 0.05) when all samples were compared with the in-tegrity control cells, and when pretreated APC samples when compared

with DV-infected HUVECs. indicatesp-value (p> 0.05) when DF-DVwere compared with APC+DF-DV.


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Figure 7: Antiapoptotic APC activity inDV-infected EVC. Propidium iodide stainingof proapoptotic nuclei was determined in thefollowing samples: uninfected HUVEC (negativecontrol, panel A), HUVEC pretreated withTNF- (panel B), DHF-DV-infected EVC MOI= 1.0 (panel C), DHF-DV (UV inactivated) MOI= 1.0 (panel D), TNF--stimulated EVC pre-treated with 10 ng/ml APC (panel E), and DHF-DV (MOI = 1.0)-infected HUVEC pretreatedwith APC (panel F). The assays are represen-tative of at least three experiments with similarresults.


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during the thrombin-catalysed activation of human protein C(the most important vasoprotective molecule). It is well knownthat the downregulation of TM causes low APC levels, promot-ing procoagulant and proinflammatory diathesis (56). Weevaluated the APC activity in HUVEC cultures infected withDF-DV or DHF-DV. We observed the downregulation of the TM-

TB-PC complex (Fig. 2) and found that DV modifies the anti-thrombotic state of the EVC surface. The transformation of theanticoagulant/anti-inflammatory state into a procoagulant/proinflammatory phenotype in vascular cells is of foremost im-

portance. Similarly, high levels of IL-8 production in DHF-DV-infected cultures (Fig. 3A) were detected, which agrees with theprevious clinical observations by Raghupaty (43), who demon-strated a correlation between elevated levels of IL-8 in patientswith grade III or IV DHF/DSS and low levels of IL-8 in DF pa-tients. Our study confirms the participation of APC in the anti-

inflammatory response of EVC during DV infection (Fig. 3B),because DHF-DV-infected cells pretreated with 10 ng/ml APCdisplayed downregulated IL-8 production. We also demonstratedthe contribution of MAPKs (ERK1/2 and p38) to IL-8 ex-pression in DHF-DV-infected cells. The specificity of these datawas confirmed by the assay in which MAPK phosphorylationwas inhibited because the pharmacological inhibition of theseprotein kinases downregulated IL-8 production (Figs. 4C and5B). It has also been reported that APC initiates intracellular sig-nalling via the activation of PAR-1 and EPCR receptors (14, 26).We found that in absence of TM (Fig. 1G), PAR-1 and EPCR arepresent (Fig. 1H-M), which suggests their participation in thisprocess. In a recent study, Bae et al. (14) provided some insight

into the PAR-1-dependent signalling mechanisms, demonstrat-ing that both EPCR and PAR-1 are associated with caveolin-1within lipid rafts of HUVEC in which the occupancy of EPCR byAPC, leads to dissociation of EPCR from caveolin-1 and recruit-ment of PAR-1 to a protective signalling pathway. Figure 5Cshows that in DHF-DV-infected cultures pretreated with specificanti-PAR-1 antibody, IL-8 was downregulated. Similarly, whenDHF-DV-infected cells were pretreated with APC or anti-PAR-1antibody, phospho-ERK1/2 was reduced (Fig. 5B). Meza (44)and Lee (45) reported that the release of IL-8 induces an increasein vascular permeability, during DV infection of endothelialcells. We have presented novel data relating to the endothelialbarrier stabilisation mediated by APC during DV infection of

EVC. Figure 6 (panels E and G) shows that DV-infected cells thathad been pretreated with APC were less permeable than cells nottreated with APC. We also observed that a barrier protective ef-fect can be induced (at least in part) by the MAPKs p38 andERK1/2.

In the present study, we also observed the participation ofAPC in the downregulation of the apoptotic response induced bythe DV infection of EVC. Espina (46) and Matsuda (47) pro-posed that apoptosis contribute to pathogenesis and tissue dam-age during Dengue virus infection. APC reduces organ damagein animal models of sepsis and directly prevents apoptosis in hy-poxic human brain endothelium (48, 49). There is some evidencethat the APC antiapoptotic function is independent of its anti-

coagulant properties, in that two APC mutants with substantiallyreduced anticoagulant activity retain their antiapoptotic effects(50). Both basic and clinical studies have provided an extensivebody of research into the cytoprotective role of the protein Cpathway (1213, 15, 51). Our data support this idea. Figure 8shows a schematic representation of our experimental findings,which may explain different aspects of DHF/DSS pathogenesis.In Figure 8A, we indicate physiological EVC homeostasis viaantithrombotic and cytoprotective effects in uninfected cells.However, during the DV infection of EVC (mainly in the pres-ence of the more aggressive viral isolate), we observed an en-dothelial surface transformation of the anti-inflammatory/anti-coagulant state into a procoagulant/proinflammatory phenotype.

What is known about this topic? There is no safe vaccine, no experimental animal model

and no specific treatment (antiviral) for Dengue virus(DV) infection. The pathogenic mechanisms of the severeforms of the disease, such as Dengue shock syndrome(DSS) and Dengue haemorrhagic fever (DHF) are not

fully understood. The vasculopathy observed during DHF/DSS is the resultof endothelial dysfunction caused by both direct (viralfactors) and indirect (host factors) mechanisms, but todate research on Dengue pathogenesis has almost exclus-ively relied on studies of host factors. Very few reports ofviral factors are available.

Recently, we found that during endothelial vascular cell(EVC) injury caused by an aggressive DV isolate, tissuefactor is up-regulated, which favors thrombin generationand the activation of the protease activated receptortype-1 (PAR-1). We also demonstrated that PAR-1 (in-flammation) and TF (coagulation) receptors can be down-

regulated in the presence of MAPKs (p38 and ERK1/2)specific inhibitors suggesting a link between coagulationand inflammation.

What does this paper add? We reported for the first time, the downregulation of the

cytoprotective protein C pathway (inflammation-coagu-lation, apoptosis and barrier protection) in human en-dothelial vascular cells (EVC) during DV infection.

We present novel data on the specific activation (damage)of EVC by DV, that causes thrombomodulin (TM) re-lease, which affects the formation of the TM-thrombin-protein C complex and APC generation, that modify theantithrombotic/anti-inflammatory state into a procoagu-lant/proinflammatory phenotype of endothelial vascularcells, which could be explain the vascular damage (vascu-lopathy) observed during DHF/DSS.

Furthermore, we observed that APC cytoprotective regu-latory pathways involve the MAPKs, p-38 and ERK 1/2pathways via PAR-1 and EPCR receptors during the DVinfection of EVC.

This study provides new perspectives and has impli-cations in the design of appropriate (e.g. pharmacologicaladministration of APC) therapies to target the thrombo-inflammatory responses that occur during DHF/DSS.


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DV causes TM release (sTM), which affects the formation of theTM-TB complex and alters both the anticoagulant and cytopro-tective protein C pathways. Our data support the involvement ofthe MAPKs p38 and ERK1/2 in this process. This study providesa new perspective and has important implications for the design

of appropriate therapies to target the thrombo-inflammatory re-sponses that occur during DHF/DSS. It is well known that thepharmacological administration of APC reduces the mortalityassociated with severe sepsis in humans (Recombinant HumanProtein C Worldwide Evaluation in Severe Sepsis [PROWESS]clinical trial) (5152) and in murine injury models. The cytopro-

tective effects of APC may also be of fundamental importanceduring the control of DSS.

Acknowledgements

We wish to thank Dr. Daniel Nez, Dr. Escalona and Dr. Ramrez of the

Gynecology and Obstetrics Clinic No. 4 of the Mexican Institute for SocialSecurity (IMSS) for their generous donation of the biological material for es-tablishing primary cultures. We also thank the CONACYT and PAPIIT Pro-gram of DGAPA UNAM, for their financial support. We are grateful to Dr.Fredy Cifuentes, C. Castellanos and Dr. Luis Vaca for their support in theConfocal microscopy image and the Biomedical Science PhD program(PDCB) - UNAM. We thank also to Ms Eva Reyes for writing support.

Figure 8: Role of the TM-TB-PC complex during DV infection ofendothelial cells. In this model, we summarise the possible regulatorysignalling pathways involved in EVC activation, which might explain differ-ent aspects of DHF/DSS pathogenesis. A) Physiological EVC homeostasisin terms of the antithrombotic and cytoprotective effects in uninfectedcells. However, during DHF-DV infection of EVC (panel B), we observedan endothelial surface transformation of the anti-inflammatory/anti-

coagulant state into a procoagulant/proinflammatory phenotype. DV in-duces TM release (sTM), affecting the formation of the TM-TB complex,altering both the anticoagulant and cytoprotective protein C pathwaysand destabilising the endothelial barrier, producing increased vascularpermeability. Our data support the involvement of the MAPKs, p38 andERK1/2, and PAR-1 response in this process.

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Dengue infection protein C