Microparticles Provide a Novel Biomarker To Predict Severe Clinical Outcomes of Dengue Virus Infection

Nuntaya Punyadee; Dumrong Mairiang; Somchai Thiemmeca; Chulaluk Komoltri; Wirichada Pan-ngum; Nusara Chomanee; Komgrid Charngkaew; Nattaya Tangthawornchaikul; Wannee Limpitikul; Sirijitt Vasanawathana; Prida Malasit; Panisadee Avirutnan

doi:10.1128/JVI.02207-14

. 2014 Nov 19;89(3):1587–1607. doi: 10.1128/JVI.02207-14

Microparticles Provide a Novel Biomarker To Predict Severe Clinical Outcomes of Dengue Virus Infection

Nuntaya Punyadee ^a,^b, Dumrong Mairiang ^a,^c, Somchai Thiemmeca ^a,^b, Chulaluk Komoltri ^d, Wirichada Pan-ngum ^e, Nusara Chomanee ^f, Komgrid Charngkaew ^f, Nattaya Tangthawornchaikul ^a,^c, Wannee Limpitikul ^g, Sirijitt Vasanawathana ^h, Prida Malasit ^a,^c, Panisadee Avirutnan ^a,^c,^✉

Editor: A García-Sastre

PMCID: PMC4300736 PMID: 25410854

ABSTRACT

Shedding of microparticles (MPs) is a consequence of apoptotic cell death and cellular activation. Low levels of circulating MPs in blood help maintain homeostasis, whereas increased MP generation is linked to many pathological conditions. Herein, we investigated the role of MPs in dengue virus (DENV) infection. Infection of various susceptible cells by DENV led to apoptotic death and MP release. These MPs harbored a viral envelope protein and a nonstructural protein 1 (NS1) on their surfaces. Ex vivo analysis of clinical specimens from patients with infections of different degrees of severity at multiple time points revealed that MPs generated from erythrocytes and platelets are two major MP populations in the circulation of DENV-infected patients. Elevated levels of red blood cell-derived MPs (RMPs) directly correlated with DENV disease severity, whereas a significant decrease in platelet-derived MPs was associated with a bleeding tendency. Removal by mononuclear cells of complement-opsonized NS1–anti-NS1 immune complexes bound to erythrocytes via complement receptor type 1 triggered MP shedding in vitro, a process that could explain the increased levels of RMPs in severe dengue. These findings point to the multiple roles of MPs in dengue pathogenesis. They offer a potential novel biomarker candidate capable of differentiating dengue fever from the more serious dengue hemorrhagic fever.

IMPORTANCE Dengue is the most important mosquito-transmitted viral disease in the world. No vaccines or specific treatments are available. Rapid diagnosis and immediate treatment are the keys to achieve a positive outcome. Dengue virus (DENV) infection, like some other medical conditions, changes the level and composition of microparticles (MPs), tiny bag-like structures which are normally present at low levels in the blood of healthy individuals. This study investigated how MPs in culture and patients' blood are changed in response to DENV infection. Infection of cells led to programmed cell death and MP release. In patients' blood, the majority of MPs originated from red blood cells and platelets. Decreased platelet-derived MPs were associated with a bleeding tendency, while increased levels of red blood cell-derived MPs (RMPs) correlated with more severe disease. Importantly, the level of RMPs during the early acute phase could serve as a biomarker to identify patients with potentially severe disease who require immediate care.

INTRODUCTION

Each year, up to 390 million people are infected by dengue virus (DENV), the most significant mosquito-borne virus of the Flaviviridae family, posing serious socioeconomic and health burdens globally (1). Four serotypes of viruses (DENV serotype 1 [DENV-1], DENV-2, DENV-3, and DENV-4) are transmitted to humans by Aedes mosquitoes to establish both endemic and epidemic transmission cycles primarily in tropical and subtropical countries (1). Although the majority of DENV infections in humans are subclinical, a small fraction of infected individuals develops clinical symptoms ranging from a self-limiting mild flu-like illness, namely, dengue fever (DF), which usually resolves without any complications, to a life-threatening capillary leakage syndrome called dengue hemorrhagic fever (DHF) or dengue shock syndrome (DSS) (2). DHF/DSS is marked by vascular leakage, resulting in hemoconcentration accompanied by thrombocytopenia and abnormalities in liver function and coagulation, a constellation that may result in hemorrhage, shock, organ failure, and, ultimately, death (3). Currently, there are no vaccines or specific therapeutics for severe DENV infection. Rapid and reliable diagnosis along with immediate and appropriate fluid replacement is the key for successful clinical management to achieve a positive outcome in patients with DHF/DSS.

Microparticles (MPs) are a heterogeneous population of small cell-derived vesicles generated by an active energy-dependent cellular process called “vesiculation” or “ectocytosis” which can occur either spontaneously or in response to various stimuli, such as cell activation, apoptosis, or stress (4). MPs are characterized by their size (diameter, 0.1 to 1 μm), the presence of negatively charged phospholipids (phosphatidylserine [PS]) on their surfaces, and an antigenic profile pointing to their cellular origin (4). Under normal physiological conditions, constitutive vesiculation is an ongoing process for the majority of cells, and thus, significant levels of MPs originating from different cells can always be detected in the blood (5). Changes in the level, the cellular origin, and the population composition of MPs in the circulation may indicate distinct pathological conditions, such as cancer, tissue injury, autoimmunity, and inflammation, as well as cardiovascular, hematological, and infectious diseases (4). An elevation in the level of circulating MPs was found to be associated with systemic infection, such as sepsis (6), human immunodeficiency virus infection (7), malaria (8, 9), and active chronic hepatitis C (10). Growing evidence suggests the potential usage of MPs as diagnostic biomarkers (11).

It is unclear whether MPs play a role in dengue pathogenesis. During the acute phase of dengue disease, cellular activation and apoptosis directly induced by DENV as well as inflammatory mediators and reactive oxygen species generated from the interaction between viruses (or infected cells) and the host immune system may trigger many types of cells to shed MPs. We therefore performed experiments to assess this possibility. The methodology used to detect and characterize MPs produced by DENV-infected cells in vitro was set up and extended to the ex vivo analysis of MPs in blood specimens taken from patients at multiple time points and with different levels of clinical severity, leading to the novel findings reported here. Our results point to multiple roles of MPs in dengue pathogenesis and provide a potential novel biomarker that can distinguish DHF patients from DF patients.

MATERIALS AND METHODS

Cells and viruses.

All transformed cell lines used in this study were obtained from ATCC. HepG2 human hepatocellular carcinoma cells were grown in Dulbecco's modified Eagle medium (DMEM; Gibco) supplemented with 10% fetal bovine serum (FBS; Gibco) with 1% nonessential amino acids (Gibco) and 1 mM sodium pyruvate (Biochrome). MEG-01 human megakaryoblast cells and U937 human myelomonocyte cells were cultured in RPMI 1640 medium (Gibco) supplemented with 10% FBS. EAhy926 human umbilical cord vein endothelial cells were cultured in Dulbecco's modified Eagle medium–nutrient mixture F-12 (DMEM/F-12; Gibco) supplemented with 10% FBS. African green monkey Vero cells were grown in minimum essential medium (MEM; Gibco) with 10% FBS. C6/36 Aedes albopictus mosquito cells were grown in Leibovitz-15 medium (L-15; Gibco) supplemented with 10% FBS and 10% tryptose phosphate broth (Sigma). Primary human umbilical cord vein endothelial cells (HUVECs) were isolated as previously described (12) and grown in medium 199 (M-199; Gibco) containing 10% FBS on 1% gelatin-coated surfaces. Primary human peripheral blood mononuclear cells (PBMCs) were isolated from buffy coats obtained from healthy volunteers by Ficoll-Hypaque (Pharmacia) density gradient centrifugation. All cell culture media were supplemented with 2 mM l-glutamine (Sigma), 100 U/ml penicillin G, and 100 μg/ml streptomycin sulfate (Sigma).

To generate virus stocks, the Hawaii strain of DENV-1, the 16681 strain of DENV-2, the H87 strain of DENV-3, and the H241 strain of DENV-4 were propagated in C6/36 cells. The viral titer was determined by a focus-forming assay on Vero cells. Briefly, 10-fold serially diluted (range, 10⁻¹ to 10⁻⁷) virus stock was added in duplicate to individual wells of tissue culture 96-well microtiter plates containing Vero cells at ∼90% confluence. Virus was incubated with Vero cells for 2 h at 37°C, after which the wells were overlaid with 1.5% gum tragacanth (Sigma) in MEM with 4% FBS. After culture at 37°C for 72 h, the overlay medium was removed and the wells were washed with phosphate-buffered saline (PBS). Cells were fixed with 3.7% formaldehyde in PBS for 10 min at room temperature and permeabilized with 2% Triton X-100 (Fluka) in PBS (Triton X-100 buffer) for 10 min at room temperature. Cells were stained with the culture supernatant of mouse monoclonal antibody (MAb) specific to flavivirus envelope protein (4G2) at 37°C for 1 h. Following several washes, the wells were incubated with horseradish peroxidase (HRP)-conjugated anti-mouse IgG (1:1,000 dilution in Triton X-100 buffer; Dako) for 30 min at 37°C. The wells were washed, and infectious foci were visualized with 3,3′diaminobenzidine tetrahydrochloride (DAB) substrate (Sigma) after a 5- to 10-min incubation at room temperature. The wells were rinsed with water and dried prior to analysis.

DENV infection and MP isolation.

Cells (1 × 10⁶) were infected with DENV-2 at multiplicities of infection (MOIs) of 1 and 5. Supernatants and cells were harvested at 24, 48, and 72 h after infection. Harvested cells were washed twice, and the percentage of apoptotic cells was measured by use of a fluorescein isothiocyanate (FITC)-conjugated annexin V (AnV) detection kit I (BD Biosciences) according to the manufacturer's instruction and analyzed by use of a FACSCalibur flow cytometer (Becton, Dickinson). Data were processed with CellQuest software (Becton, Dickinson). To quantitate the percentage of infected cells, harvested cells were fixed with 2% formaldehyde (BDH, England), permeabilized with 0.1% Triton X-100 (Sigma), stained with 100 μl undiluted hybridoma culture supernatants of anti-DENV-2 NS1 MAb clone 2G6 (13) followed by 6 μg/ml of FITC-labeled rabbit anti-mouse IgG (Dako), and analyzed by flow cytometry. For MP isolation, culture supernatants from mock- and DENV-infected cells were collected at the times indicated below and serially centrifuged at 500 × g and at 13,000 × g (10 min each) to remove cell debris and small aggregates, respectively. A final centrifugation step at 20,000 × g for 60 min was performed to pellet the MPs. The MP pellets were washed thoroughly with PBS and kept at −70°C until MP analysis. In some experiments, wild-type 293T cells and 293T cells stably expressing T cell immunoglobulin domain and mucin domain 1 (TIM-1; kindly provided by Ali Amara) were infected with DENV-2 at an MOI of 0.3 for 2 days until both cells and supernatants were harvested and processed as described above.

To determine MP generation in cells infected with the four serotypes of DENV, HepG2 cells were infected with each serotype of DENV at an MOI of 5 in duplicate. One set of infected cells was UV irradiated at 3,000 J/s for 10 min (Hoefer UVC500 UV cross-linker) to inactivate DENV. Supernatants and cells were harvested at 48 h after infection. Analyses of infection efficiency and apoptosis level and MP isolation were performed as described above.

Analysis of MPs generated from DENV-infected cells.

MPs from culture supernatants (1 ml) of mock- and DENV-infected HepG2 cells were isolated as described above. Isolated MPs were stained with 3 μl of FITC-conjugated AnV (BD Biosciences) plus 5 μl of 10-fold-concentrated AnV buffer for 30 min on ice. AnV buffer (300 μl; 150 mM NaCl, 2 mM CaCl₂, 10 mM HEPES, pH 7.4) was added prior to flow cytometric analysis. MPs were gated by the use of standard beads of defined size (BD Biosciences). Only events with a size of less than 1 μm were further evaluated for the percentage of AnV-positive MPs. Events within this size limit were stopped at 10,000 counts.

For analysis of DENV antigens expressed on the surfaces of MPs, 40 μl of 10-fold-concentrated mock- and DENV-infected MPs was incubated with 5 μl of Fc receptor blocking solution (BioLegend) for 15 min on ice, followed by a 1-h incubation on ice with 1 μg/ml of FITC-conjugated anti-NS1 2G6 MAb or 1 μg/ml of allophycocyanin (APC)-conjugated anti-envelope protein (anti-E) 4G2 MAb. FITC- and APC-conjugated mouse MAbs against unrelated antigens were used as negative controls for NS1 and E staining, respectively. After a 30-min incubation with phycoerythrin (PE)-conjugated AnV (3 μl; BD Biosciences) on ice, AnV buffer (300 μl) was added and the solution was transferred into a tube with a known density of fluorescent TruCount beads (BD Biosciences) and analyzed using a BD FACSCalibur flow cytometer. PE-conjugated mouse IgG specific to unrelated antigens was used as a negative control for AnV staining. MP acquisition gating was performed as described above. Absolute numbers of MPs were calculated according to the following formula: (number of events of MPs)/(number of events of TruCount beads collected) × (TruCount Bead count per test/sample volume) × dilution factor.

Transmission electron microscopy.

HepG2 cells infected with DENV at an MOI of 5 were harvested at 48 h postinfection and fixed with 1% glutaraldehyde in 0.1 M sodium cacodylate buffer at 4°C overnight. Fixed cells were then treated with 1% osmium tetroxide in Millonig buffer (120 mM Na₂HPO₄, 100 mM NaOH, 30 mM d-glucose [pH 7.4]) for 1 h at 4°C and later dehydrated with graded ethanol solutions: 70%, 80%, 90%, and 100%. Dehydrated samples were transferred from 100% ethanol to a 50:50 mixture of ethanol-propylene oxide and finally to 100% propylene oxide before being infiltrated with epoxy resin (Ted Pella Inc., Redding, CA) and polymerized at 60°C for 48 h. Ultrathin sections were cut on an ultramicrotome using a Diatome diamond knife (Leica EM UC7) and then mounted on grids and stained with 2% uranyl acetate and lead citrate solution. Images were collected by a transmission electron microscope (JEM 1230; JEOL, Peabody, MA).

Negative staining.

MPs from culture supernatants of mock- and DENV-infected HepG2 cells were isolated as described above. A 10-fold-concentrated MP preparation (3 μl) was adsorbed on a glow-discharged Formvar-carbon-coated nickel grid (Electron Microscopy Sciences, Hatfield, PA) for 5 min. Excess sample was removed by a filter paper, followed by a wash with distilled water. The grids were then stained with 1.5% phosphotungstic acid (PTA) (pH 6.6) for 2 min, and air-dried grids were visualized using an FEI Tecnai T20 transmission electron microscope operated at 200 kV.

Immunogold labeling.

Expression of PS on the surface of DENV MPs was evaluated by immunogold labeling with AnV as previously described, with some modifications (14). Briefly, a 10-fold-concentrated MP preparation was adsorbed on a glow-discharged Formvar-carbon-coated nickel grid for 5 min. Afterward, the grids were stained with 3 μl FITC-conjugated AnV in 10-fold-concentrated AnV buffer for 1 h at room temperature. Excess staining was removed by a filter paper, and the grids were then incubated with blocking buffer (PBS containing 1% bovine serum albumin [BSA]; Aurion, Wageningen, the Netherlands). After a 10-min incubation at room temperature, the grids were washed with PBS containing 0.1% BSA and then incubated with 10-nm gold nanoparticle-conjugated mouse anti-FITC MAb (Aurion) for 1 h at room temperature. Subsequently, the grids were washed with 0.1% BSA–PBS, stained with 1.5% PTA for 2 min, and then washed with distilled water and air dried. The specificity of AnV immunogold labeling on MPs was verified by staining the grid that was adsorbed with PBS instead of MPs, followed by the immunolabeling procedure described above. The samples were observed using an FEI Tecnai T20 transmission electron microscopy operated at 200 kV.

In vitro experiments with immune complexes (ICs).

Healthy adult volunteers provided blood for this study following the provision of written informed consent approved by the Institutional Review Board of the Faculty of Medicine Siriraj Hospital (protocol number 159/2557). Columns, buffers, and the method for construction of a high-gradient magnetic separator (HGMS) apparatus for isolating cells from whole blood were described previously (15). Briefly, the isolation of human erythrocytes by using HGMS (X-Zell Biotech) was as follows: 2 ml of EDTA-blood from healthy volunteers was centrifuged at 800 × g for 10 min, washed three times with 5 ml of PBS containing 0.1% BSA (Sigma), and centrifuged at 500 × g for 5 min after each wash. Subsequently, 250 μl of packed red blood cells (RBCs) was resuspended in 750 μl of PBS containing 3% BSA, the suspension was mixed with 20 μl CD45 HMxBeads (X-Zell Biotech), and the mixture was incubated for 20 min on a constantly rotating shaker at room temperature. After incubation, the suspension was centrifuged at 500 × g for 5 min to remove unbound beads, followed by the addition of 1 ml PBS containing 3% BSA and 15 μl of CD45 HMxBeads for a second round of incubation on a constantly rotating shaker at room temperature for 20 min. During the second round of incubation, an HMX column was prepared for purification of RBCs. Briefly, the column was filled with PBS containing 1% BSA, and after removing the remaining air bubbles by gentle finger tapping, a 26-gauge 1-in. needle was connected to the stopcock. The column was placed between the poles of the dipole magnet and equilibrated for 5 to 10 min with PBS containing 1% BSA. After finishing the second round of incubation, the suspension was applied to the top of the column and the flowthrough was collected in a 15-ml tube. The flowthrough was centrifuged at 500 × g for 5 min, washed once with DGVB⁺⁺ (2.5 mM barbital sodium, 139 mM dextrose, 71 mM NaCl, 0.1% [wt/vol] gelatin, 0.15 mM CaCl₂, 1 mM MgCl₂), and finally, checked for purity (>99.9%) by flow cytometry (FACSCalibur; BD Biosciences).

In vitro analysis of IC opsonization and vesiculation.

Purified DENV NS1 (100 ng/ml) (16) was incubated with 5-μg/ml IgG fractions purified from pooled dengue patient convalescent-phase serum (PCS) in the presence of 5 × 10⁶ purified RBCs diluted in DGVB⁺⁺ and 10% autologous serum for 1 h at 37°C. After centrifugation, the supernatants were collected for MP analysis and the cells were washed once with 1 ml of DGVB⁺⁺. Subsequently, 7.5 × 10⁶ autologous PBMCs diluted in 100 μl of DGVB⁺⁺ were added and the mixture was incubated for 30 min at 37°C. Afterwards, the supernatants were collected for MP analysis and the cells were washed once with 0.1% BSA–PBS, followed by flow cytometric analysis. Negative controls, which were run in parallel, consisted of purified NS1 diluted with 5 μg/ml of PCS in the presence of 20 mM EDTA (to inhibit complement activation) or with 5 μg/ml of pooled non-dengue patient serum (PND).

To determine the binding of ICs to human erythrocytes before and after addition of PBMCs, cells were washed with 0.1% BSA–PBS and stained with 20 μg/ml of anti-C3d clone YB2-39HL (17) diluted in 0.1% BSA–PBS for 1 h on ice, followed by 4 μg/ml Alexa Fluor 488 goat anti-rat IgG antibody (Invitrogen) for 30 min on ice. After washing, complement fragment deposition on erythrocytes was analyzed by flow cytometry and the data were processed with FlowJo software (Tree Star).

MPs were quantified after in vitro IC experiments. Briefly, 34 μl of supernatant was incubated with 5 μl of 10-fold-concentrated AnV buffer and 3 μl of FITC-conjugated AnV either with 3 μl of PE-conjugated anti-CD45 or with 5 μl of PE-Cy5-conjugated anti-CD235a at a dilution of 1:100. After 30 min, 300 μl of AnV buffer was added and the mixture was transferred into a tube with a known number of fluorescent TruCount lyophilized pellet beads and analyzed by flow cytometry as described above.

Patients.

This study was performed at the Khon Kaen Provincial Hospital and Songkhla Hospital between the epidemic seasons of 2010 and 2013. Written informed consent was sought from the parents or guardians of all participants in accordance with the guidelines of the ethical committees of the Ministry of Public Health (protocol number 92/2550), Faculty of Medicine Siriraj Hospital (protocol number 349/2550), the Khon Kaen Provincial Hospital, and the Songkhla Hospital. Acute dengue virus infection was diagnosed by using reverse transcription-PCR (RT-PCR)-based DENV gene identification or by using DENV-specific IgG and IgM capture enzyme-linked immunosorbent assay (ELISA) (18, 19). The inclusion criteria for the patients enrolled in the study were described elsewhere (16). Patients who tested positive for DENV were classified into the DF or DHF group according to the World Health Organization (WHO) 1997 guideline (2), whereas those with negative RT-PCR and DENV antibody results were assigned to the control group, labeled the group with other febrile illness (OFI). Samples used for this study were derived from patients that ranged in age from 5 to 15 years (mean age, 10.3 years); 36% of samples were obtained from females, and 64% were obtained from males. Of a total of 53 samples included in this study, 24 samples were from DHF cases, 19 were from DF cases, and 10 were from OFI cases. Of the 19 DF patients, there were 3 cases with DENV-1 infection, 6 cases with DENV-2 infection, 7 cases with DENV-3 infection, and 1 case with DENV-4 infection. Of the 24 DHF patients, there was 1 case with DENV-1 infection, 16 cases with DENV-2 infection, and 3 cases with DENV-3 infection. All dengue cases were secondary infections. Data were derived from plasma samples collected daily during the febrile phase and paired convalescent-phase samples obtained at 2 weeks and 2 months after defervescence. Study day 0 (defervescence) was defined as the calendar day on which the patient's temperature fell and stayed below 37.8°C. A summary of the subjects' characteristics is shown in Table 1.

TABLE 1.

Demographic, clinical, and key laboratory characteristics of patients enrolled in the study^a

Characteristic	Result for the following patient groups:			P value
Characteristic	OFI patients (n = 10)	DF patients (n = 19)	DHF patients (n = 24)	P value
No. (%) male patients	5 (50)	10 (53)	19 (79)	0.1012
Age (yr)	9.5 (6, 10.8)	10 (8, 11)	11 (8.3, 13.8)	0.1824
Body wt (kg) at enrollment	22.5 (18.5, 35.7)	33.0 (23.2, 49.0)	35.5 (25.5, 46.0)	0.0683
Platelet nadir (10⁹/liter)	184.5 (161.0, 198.0)	96.0 (52.5, 140.0)	27.5 (21.2, 48.8)	<0.0001
Day of acute illness of platelet nadir	−0.5 (−1.8, 0.0)	0 (0, 1)	0 (0, 1)	0.0454
Maximum RBC count (10¹²/liter)	4.7 (4.5, 4.9)	5.4 (4.9, 5.8)	5.5 (5.2, 6.0)	0.0040
Day of acute illness of maximum RBC count	0 (−2, 1)	0.0 (−0.5, 0.5)	0 (−1, 1)	0.5826
No. (%) of patients with mucosal bleeding	0 (0)	5 (29)	12 (71)	0.0078
Day of acute illness of first episode of mucosal bleeding	—	−1 (−1.5, −0.5)	0 (−1, 0)	0.2007

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Data are presented as the number (percentage) for categorical variables and median (25th, 75th percentiles) for continuous variables. OFI, other febrile illness; DF, dengue fever; DHF, dengue hemorrhagic fever; RBC, red blood cell; —, OFI patients did not have mucosal bleeding.

Blood sample collections.

Human peripheral blood was collected in Vacutainer K3-EDTA tubes (BD Biosciences) and processed within 1 h after blood collection. Blood samples were centrifuged at 750 × g for 20 min at 4°C to obtain platelet-rich plasma (PRP), followed by a second centrifugation step at 1,250 × g for 15 min at 4°C to obtain platelet-poor plasma (PPP). PPP was carefully removed, aliquoted, snap-frozen in liquid nitrogen, and stored at −80°C until MP analysis.

Labeling of plasma MPs.

The cellular origins of plasma MPs were determined using fluorescence-conjugated MAbs specific to cellular markers. DENV antigens associated with plasma MPs were identified using fluorescence-conjugated MAbs specific to E and NS1. Briefly, PPP (15 μl) was incubated with 3 μl of Fc blocker for 30 min on ice prior to the addition of 3 μl FITC-conjugated AnV to identify MPs, 1 μl APC-conjugated anti-DENV E or NS1 MAbs, and 5 μl PE- or peridinin chlorophyll protein (PerCP) complex-labeled MAbs specific for CD235a (glycophorin A; for red blood cell-derived MPs), CD41a (glycoprotein IIb/IIIa; for platelet-derived MPs), CD14 (for monocyte-derived MPs), CD66 (for granulocyte-derived MPs), CD3 (for T cell-derived MPs), CD19 (for B cell-derived MPs), and CD56 (for NK cell-derived MPs). In some experiments, fluorochrome-conjugated isotype-matched control IgG1 (MOPC-21) and IgG2a (UPC-10) antibodies were used instead of MAbs specific for cellular markers. All antibodies except APC-conjugated anti-DENV MAbs were purchased from BD Biosciences; APC-conjugated anti-DENV MAbs were generated in-house using an APC conjugation kit (Biolegend) according to the manufacturer's instructions. After a 30-min incubation in the dark, 300 μl of AnV buffer was added and the mixture was transferred into a tube with a known density of fluorescent TruCount beads and analyzed by flow cytometry. To avoid any sedimentation of beads, the samples were thoroughly mixed prior to flow cytometric analysis. All of the staining buffers and fluorochrome-labeled antibodies were clarified by centrifugation at 13,000 × g for 10 min to remove small aggregates before staining.

MP procoagulant activity.

MP procoagulant activity was assayed with a functional chromogenic method on micro-ELISA plates (Zymuphen MP-activity) according to the manufacturer's instructions.

Statistical analysis.

For the in vitro studies, data were collected from at least three independent experiments. Data sets were compared using a two-tailed, unpaired t test. Multiple comparisons were performed using an analysis of variance (ANOVA) test. Correlation coefficients were estimated on the basis of simple linear regressions and Pearson product-moment correlation using GraphPad Prism (version 5) software. Statistical significance was achieved at a P value of <0.05.

For the ex vivo studies, the demographics of three patient groups (the DF, DHF, and OFI groups) were compared using the Fisher exact test and the Kruskal-Wallis test for categorical and numeric data, respectively. The levels and dynamics of MPs, RBCs, and platelets among the patient groups during the acute phase of infection were analyzed by use of a mixed model using a quadratic form. There were some missing data, since the disease day at admission of each patient was different. Only patients with data on at least three consecutive days were included in the analysis. Since the levels of AnV-positive (AnV⁺) MPs, CD41⁺ MPs, CD235⁺ MPs, RBCs, and platelets were positively skewed, a logarithmic transformation was applied. In addition, the ratios in percentage terms between CD41a⁺ MPs and AnV⁺ MPs and between CD235⁺ MPs and AnV⁺ MPs were used as another means to normalize CD41a⁺ MPs and CD235⁺ MPs, respectively.

To determine whether MP, RBC, and platelet levels and dynamics were significantly different among the patient groups, t tests were performed on mixed models and P values were calculated on the basis of the Satterthwaite approximation for denominator degrees of freedom. To determine whether MP, RBC, and platelet levels on the same date between two patient groups or between two dates for a single patient group were significantly different, general linear hypotheses and multiple comparisons for parametric models were used on mixed models.

To determine whether MP, RBC, and platelet levels among patient groups were associated with factors other than DENV infection, such as genetic background, the levels of total AnV⁺ MPs, CD41a⁺ MPs, CD235⁺ MPs, RBCs, and platelets at 14 days and 60 days after defervescence were tested with Welch's t test for any significant differences among patient groups. Finally, receiver operating characteristic (ROC) curves of absolute CD235⁺ MPs and percentage of CD235⁺ MPs were constructed to assess the performances of the earliest levels of CD235⁺ MPs before defervescence as a biomarker for predicting the severity of dengue virus infection.

All statistical analyses of the ex vivo data were performed with the R programming language (version 3.0.2), with the lme4 package used for mixed model construction, the lmerTest package used for P value calculation from the Satterthwaite approximation, the multicomp package used for general linear hypotheses and multiple comparisons, and the ROCR package used for ROC curve construction. Graphs and charts were plotted with GraphPad Prism 5 software.

RESULTS

DENV infection leads to apoptotic death and MP shedding.

Analogous to the results of previous studies (20, 21), infections of a human hepatocyte cell line (HepG2) and primary human umbilical cord vein endothelial cells (HUVECs) by DENV-2 led to apoptotic death in an MOI-dependent manner (Fig. 1). To assess MP generation from different cell types induced by DENV-2, culture supernatants were harvested at various time points after infection. MPs were isolated by sequential centrifugation and labeled with the PS-binding protein annexin V (AnV) prior to flow cytometric analysis. Standard microsphere beads of 1 and 3 μm in diameter (Fig. 2A) were used as size markers to identify the MP population consisting of particles smaller than 1 μm, as assessed by the logarithmic amplification of forward scatter (FSC) and side scatter (SSC) signals (Fig. 2B, region R1) along with AnV binding (Fig. 2C and D, left upper quadrants). Consistent with increased DENV-induced apoptotic death over time after infection (Fig. 1), the percentages of AnV⁺ MPs produced by DENV-2-infected HepG2 cells and HUVECs rose and were significantly higher than those spontaneously produced by mock-infected cells (Fig. 2E and F). Other potential target cell types for DENV in humans, including an endothelial cell line (EAhy926), primary monocytes, a monocytic cell line (U937), and a megakaryocyte cell line (MEG-01), also shed MPs in response to DENV infection (Fig. 2G to J). Infection with all four serotypes of DENV of HepG2 cells also led to apoptotic death (Fig. 1E and F) and MP shedding (Fig. 2K). The amount of MP generation likely depends on the degree of apoptotic cell death, the process of which requires the active replication of DENV, as incubation of inactivated UV-irradiated DENV with cells did not induce apoptotic cell death (Fig. 1E and F) or MP shedding (Fig. 2K). In support of this, the cells overexpressing T cell immunoglobulin domain and mucin domain 1 (TIM-1) protein, which was recently identified to be an entry receptor for DENV (22), enhanced DENV susceptibility, leading to more apoptotic death and, consequently, levels of MP generation higher than those in the wild-type control cells (data not shown).

FIG 1 — DENV infection causes apoptotic cell death. (A to D) HepG2 cells (A and B) and HUVECs (C and D) were infected with DENV-2 at MOIs of 1 and 5 and harvested at 1, 2, or 3 days postinfection for intracellular NS1 staining to determine the percentage of infected cells (A and C) and AnV labeling to measure the percentage of cells that had undergone apoptosis (B and D). (E and F) HepG2 cells were infected with DENV-1 (strain Hawaii), DENV-2 (strain 16681), DENV-3 (strain H87), and DENV-4 (strain H241) at an MOI of 5 and harvested at 2 days postinfection to determine the percentage of infected cells (E) and the percentage of cells that had undergone apoptosis (F). In parallel, cells were treated with equal volumes of UV-irradiated DENV (UV-DENV), harvested at 2 days posttreatment, and processed in the same way as were cells incubated with live DENV. Data are the mean ± SD from three to four independent experiments. Asterisks denote statistically significantly differences between infected or apoptotic cells and mock-infected cells (*, P < 0.05; **, P < 0.001; ***, P < 0.0001).

Fig 2 — DENV infection induces MP production from various cell types. (A to D) Representative histogram and density plots showing the gating protocol for MPs. (A) Histogram profiles of FSC of the 1- and 3-μm-diameter calibrator beads which were used as size markers for MP identification. (B) MPs were defined as events with a size of less than 1 μm, which are gated in the R1 window. (C and D) Size-selected events (gate R1) are plotted as a function of their fluorescence for specific AnV-FITC binding against FSC. Positively labeled events in the left upper quadrants are considered AnV⁺ MPs. Examples of density plots showing the percentage of AnV⁺ MPs in culture supernatants of mock-infected (C) and DENV-infected (D) HepG2 cells harvested at 48 h postinfection are depicted. (E to J) The percentages of AnV⁺ MPs generated by mock- and DENV-infected HepG2 cells (E) and HUVECs (F) at 1, 2, or 3 days, EAhy926 (G) and U937 (I) cells at 2 days, primary monocytes (H) at 1 day, and MEG-01 cells (J) at 7 days postinfection were measured. (K) The percentage of AnV⁺ MPs generated from HepG2 cells treated for 2 days with UV-irradiated or live viruses of all four DENV serotypes was analyzed. All cell types were infected by DENV at an MOI of 5. Data are the mean ± SD from three to four independent experiments. Asterisks denote statistically significantly differences between the percentage of AnV⁺ MPs produced by DENV-infected cells and the percentage produced by mock-infected cells (*, P < 0.05; **, P < 0.001). (L) Transmission electron micrograph of a DENV-infected HepG2 cell displaying small vesicles of 80 to 200 nm in size (arrowheads) near the cell periphery. (M and N) Budding MPs at higher magnification. (O to Q) Negative staining on the grids absorbed by buffer (O), isolated MPs (P), and sucrose density-purified virus particles (Q) released from DENV-infected HepG2 cells are depicted. (R to T) Immunogold labeling of isolated MPs. (S and T) The clusters of 10-nm gold particles (black dots) at the periphery of bilamellar vesicular structures of MPs indicate the externalization of AnV-bound PS at the outer leaflet of the MP membrane. (R) Grids adsorbed with buffer instead of MPs that then underwent the same immunogold labeling procedure used for the isolated MPs in the images shown in panels S and T.

Transmission electron microscopy of DENV-infected HepG2 cells at 48 h postinfection demonstrated a heterogeneous population of MPs of various sizes and shapes around the cell periphery (Fig. 2L). Some MPs appeared to be budding from the cell surface (Fig. 2L, arrowheads). Most MPs had an electron density similar to that of the cytoplasm and clearly defined margins (Fig. 2M and N). To investigate the MPs in greater detail, naturally liberated MPs were isolated from the supernatants of DENV-infected HepG2 cells at 48 h postinfection, processed by negative staining, and observed under a transmission electron microscope. Electron micrographs of isolated MPs showed intact double-layered membranous structures (Fig. 2P) with heterogeneous sizes between 0.1 and 1 μm (data not shown). Background negative staining adsorbed by buffer on the grids was used as a negative control (Fig. 2O). For comparison, virions were purified from the same batch of culture supernatants from which MPs were isolated and were visualized by negative-staining electron microscopy. Similar to what had been described by a previous study for the structure of human cell-derived DENV virions generated at 37°C using cryo-electron microscopy (23), negative-staining electron micrographs of sucrose density gradient-purified DENV produced by infected HepG2 cells revealed bumpy or spiky virions (diameter, ∼50 nm), which are characteristics of immature or partially mature viruses, as well as some deformed particles (Fig. 2Q). Binding of AnV to PS on the surface of MPs was also confirmed by immunogold labeling of negatively stained MPs (Fig. 2S and T). The specificity of AnV immunogold labeling on MPs was verified by staining a grid adsorbed with buffer instead of MPs, followed by the same immunogold labeling procedure described above (Fig. 2R).

MPs released from DENV-infected cells express viral antigens on their surfaces.

Because DENV-infected cells express viral proteins, i.e., envelope protein (E) (24) and nonstructural protein NS1 (25), on their surfaces, infected cell-derived MPs were examined for the presence of these viral antigens. To obtain an accurate number of MPs, the known density of TruCount beads, which were identified by their size on the basis of the logarithmic amplification of the FSC and SSC signals (Fig. 3A to C, region R2), was used as an internal reference. MPs generated from mock-infected (Fig. 3B) and DENV-infected (Fig. 3C) HepG2 cells were localized in the R1 region (as described for Fig. 2) and specifically labeled with phycoerythrin (PE)-conjugated AnV (Fig. 3F and G) but not with the negative control, an unrelated PE-conjugated IgG (Fig. 3D and E). Of note, the buffer used in the experiments showed very few and negligible background noise signals within the same MP gate (Fig. 3A). AnV⁺ MPs in DENV-infected culture supernatants were further analyzed for DENV E and NS1 antigen expression by double immunofluorescence staining with an allophycocyanin (APC)-conjugated anti-E MAb and a fluorescein isothiocyanate (FITC)-conjugated anti-NS1 MAb, respectively (Fig. 3J). The number of DENV-infected cell-derived AnV⁺ MPs labeled with IgG isotype controls (Fig. 3H) and the number of AnV⁺ MPs acquired from mock-infected supernatants labeled with anti-E or anti-NS1 MAbs were not significant (Fig. 3I). The absolute numbers of MPs were also calculated by using the known density of TruCount beads (Fig. 3K). The supernatants from DENV-infected cells contained a significantly higher total number of AnV⁺ MPs than the supernatants from mock-infected cells (P = 0.032). Notably, the majority of AnV⁺ MPs derived from DENV-infected cells were negative for viral E and NS1 (E⁻ and NS1⁻, respectively) antigens on their surfaces (Fig. 3K). About one-third of the total AnV⁺ MPs produced by DENV-infected cells were positive for at least one of the viral antigens, E and NS1 (Fig. 3K).

FIG 3 — MPs generated from DENV-infected cells express E and NS1 antigens on their surfaces. MPs isolated from 10-fold-concentrated culture supernatants of mock- and DENV-infected cells were labeled with PE-conjugated AnV (AnV-PE), APC-conjugated anti-E (E-APC), and FITC-conjugated anti-NS1 (NS1-FITC) MAbs and analyzed by flow cytometry. The absolute numbers of MPs generated in culture supernatants were measured on the basis of a known number of fluorescent TruCount beads. (A to C) Representative flow cytometric dot plots of MPs in buffer (A) and culture supernatants of mock-infected (B) and DENV-infected (C) HepG2 cells are depicted. Region R1 represents the FSC/SSC light scatter gate of MPs (size, <1 μm). Region R2 represents the known density of TruCount beads. Data acquisition was stopped when the number of TruCount beads reached 2,000 events. (D to G) Examples of density plots of MPs (events in the R1 gate) in mock-infected (D and F) and DENV-infected (E and G) cell culture supernatants stained with an unrelated PE-conjugated IgG (IgG-PE) (D and E) as negative controls or AnV-PE (F and G) are displayed. The percentages of total AnV⁺ MPs in the R1 gate (left upper quadrants) detected in the culture supernatants of mock- and DENV-infected cells are depicted. AnV⁺ MPs were further analyzed for E and NS1 expression. (H to J) Representative density plots of AnV⁺ MPs from DENV-infected cells stained with FITC- and APC-conjugated isotype control Abs (IgG-FITC and IgG-APC, respectively) (H) and AnV⁺ MPs from mock-infected (I) and DENV-infected (J) cells double stained with FITC-conjugated anti-NS1 MAb clone 2G6 (anti-NS1-FITC) and APC-conjugated anti-E MAb clone 4G2 (anti-E-APC). The percentages of AnV⁺ MPs positive for NS1 alone (left upper quadrants), E alone (right lower quadrants), and both E and NS1 (right upper quadrants) generated by mock-infected (I) and DENV-infected cells (J) are depicted. (K) The absolute numbers of total AnV⁺ MPs and AnV⁺ MPs negative for both E and NS1 (E⁻ NS1⁻), positive for E alone (E⁺ NS1⁻), positive for NS1 alone (E⁻ NS1⁺), and positive for both E and NS1 (E⁺ NS1⁺) were determined by using TruCount beads of known density (region R2 in panel A). Data are the mean ± SD from four independent experiments. Asterisks note statistically significantly differences between the percentage of AnV⁺ MPs produced by DENV-infected cells and the percentage produced by mock-infected cells (*, P < 0.05; **, P < 0.001).

The relatively low detectable levels of DENV antigen-containing MPs might reflect the fact that low levels of antigens are expressed on the surface of infected cells. During outward budding of the plasma membrane to release MPs, the DENV antigen intensity on the surface of very small particles like MPs might be too low to be positively identified by flow cytometry. Alternatively, a fraction of DENV antigen-negative MPs might be released from infected cell-derived cytokine-induced activation of uninfected cells, as not all cells (∼50 to 60%) were infected by DENV at the time of the analysis (2 days postinfection at an MOI of 5; Fig. 1A). It is worth noting that the DENV E antigen detectable on the surface of some MPs might be due to the binding of specific antibodies to virions that attach to the MP surface. Our preliminary data showed that infected cell-derived MPs were positive for DENV RNA, as detected by nested RT-PCR, and contained infectious activity which was reduced by half after washing the isolated MPs with acidic (pH 3.0) glycine buffer (data not shown). It is also possible that non-virion-associated E protein might be expressed on the surface of infected cells and thus may be incorporated into MPs generated from these cells. This explanation is supported by the study of Ng and Corner (24), as they found by immunoelectron microscopy that DENV E was located in clumps on the plasma membrane of infected cells without an association with DENV virions. Likewise, NS1 has been shown to incorporate into the plasma membrane of infected cells via a glycosylphosphatidylinositol linkage or by lipid raft association (26, 27). Additionally, secreted soluble NS1 can bind back to the plasma membrane of uninfected cells through the interaction with specific sulfated glycosaminoglycans (28). These phenomena could contribute to the positivity of NS1 detection on the surface of MPs. Taken together, DENV infection resulted in the shedding of MPs of various sizes that may or may not harbor viral proteins on their surfaces.

Circulating MPs in dengue patients.

We next measured and analyzed the cellular origins of circulating MPs in the blood of DENV-infected patients. Flow cytometric analysis of circulating AnV⁺ MPs was performed in a blinded manner. MPs were identified according to their standard size and AnV labeling as described above. Demographic information on all subjects enrolled in this study is shown in Table 1, and a summary of the infecting DENV serotype in patients is provided in Materials and Methods. The demographics were not significantly different among the patient groups. To our surprise, the total counts of AnV⁺ MPs were significantly reduced in DENV-infected patients compared with those in the other febrile illness (OFI) group during the febrile illness, except on day −2 (Fig. 4A; Table 2). No significant differences in the results for paired convalescent-phase samples collected after defervescence (day 14 and day 60) were observed among the groups (Fig. 4A; Table 2).

Fig 4 — Circulating MP levels in DENV-infected patients. Platelet-poor plasma was obtained from patients with DF, DHF, and OFIs (acute febrile diseases other than dengue) on different disease days during the acute phase (days −2, −1, 0, and 1) and the convalescent phase (days 14 and 60). Day 0 (defervescence) was defined as the calendar day on which the patient's temperature fell and stayed below 37.8°C. (A to G) The absolute numbers of AnV⁺ MPs (A) and AnV⁺ MPs expressing specific platelet marker CD41a (CD41a⁺ MPs) (B) and erythrocyte marker CD235 (CD235⁺ MPs) (C) and the percentages of CD41a⁺ MPs (D) and CD235⁺ MPs (E) from total AnV⁺ MPs in the plasma of DF, DHF, and OFI patients were quantified by immunofluorescent labeling and flow cytometric analysis. (F) Numbers of platelets measured in individual patients from each group on each day over the course of the acute phase and 14 days after defervescence. Each dot represents the absolute number (A to C) or the percentage (D and E) of total AnV⁺ MPs (A) or AnV⁺ MPs derived from platelets (B and D) or erythrocytes (C and E) and the numbers of platelets (F) and RBCs (G) for an individual patient (n = 3 to 15 patients on each disease day). Solid bars indicate the mean. Asterisks denote the means that are statistically significantly different between the indicated groups of patients (*, P < 0.05; **, P < 0.001; ***, P < 0.0001). (H and I) The correlation coefficients between the absolute counts of CD41a⁺ MPs and platelet numbers (H) and CD235⁺ MPs and RBC counts (I) were calculated. The linear regression, correlation coefficient, and P value are presented in the graphs. (J) The percentages of AnV⁺ MPs expressing CD41a⁺, CD235⁺, and other CD markers (CD41a⁻ CD235⁻) in the plasma of OFI, DF, and DHF patients are displayed by disease day.

TABLE 2.

Absolute number of AnV⁺ MPs in three groups of patients^a

Day	Total AnV⁺ MPs (mean ± SD no. of events/μl)			P value
Day	OFI patients	DF patients	DHF patients	OFI vs DF patients	OFI vs DHF patients	DF vs DHF patients
−2	7,027 ± 1,973	5,202 ± 1,068	5,668 ± 774	0.393	0.458	0.744
−1	11,060 ± 2,863	4,800 ± 761	4,606 ± 947	0.007	0.011	0.879
0	7,429 ± 1,385	4,434 ± 546	4,324 ± 648	0.021	0.031	0.901
1	16,370 ± 7,583	5,110 ± 560	3,468 ± 413	0.031	0.004	0.023
14	20,410 ± 6,650	14,380 ± 2,629	13,940 ± 2,701	0.317	0.285	0.906
60	ND	7,713 ± 1,326	9,604 ± 1,580	ND	ND	0.300

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MPs, microparticles; OFI, other febrile illness; DF, dengue fever; DHF, dengue hemorrhagic fever; ND, not done.

We further went on to identify the cellular phenotypes of those circulating MPs using a panel of specific MAbs to CD markers belonging to platelets (CD41a), erythrocytes (CD235), monocytes (CD14), granulocytes (CD66), T cells (CD3), B cells (CD19), and NK cells (CD56). Unexpectedly, only CD41a⁺ MPs and CD235⁺ MPs made up the two major populations of AnV⁺ MPs in the blood of acutely ill patients (50 to 75% of total AnV⁺ MPs), while the levels of MPs generated from other cell types, especially those that have been shown to be targets for DENV replication in vivo, including monocytes/macrophages (29) and lymphocytes (30), were remarkably low (data not shown).

Similar to the total AnV⁺ MP levels, the absolute numbers of CD41a⁺ (platelet-derived) MPs in the DHF group were significantly lower than those in the DF and OFI groups on almost every disease day during the acute phase (Fig. 4B; Table 3). Interestingly, we observed a rise in the number of CD235⁺ (red blood cell-derived) MPs starting as early as day −2 in the DHF group, and the increased numbers reached statistical significance at day −1; the numbers were still significantly higher than those in the other groups through day 14 (Fig. 4C; Table 4). At day 60 after defervescence, the CD235⁺ MP levels were comparable among all groups (Fig. 4C; Table 4). Importantly, during febrile illness the percentage of CD235⁺ MPs in the DHF group was significantly higher than those of the DF and OFI groups, whereas the percentage of CD41⁺ MPs in the DHF was significantly reduced compared with those of other groups. However, the percentages of both CD235⁺ and CD41a⁺ MPs were undistinguishable among all groups during the convalescent phase (Fig. 4D and E).

TABLE 3.

Absolute number of AnV⁺ MPs expressing the specific platelet marker CD41a in three groups of patients^a

Day	CD41a⁺ MPs (mean ± SD no. of events/μl)			P value
Day	OFI patients	DF patients	DHF patients	OFI vs DF patients	OFI vs DHF patients	DF vs DHF patients
−2	3,503 ± 926	2,911 ± 611	1,485 ± 337	0.589	0.021	0.042
−1	7,291 ± 2,414	2,637 ± 695	1,299 ± 329	0.018	0.0002	0.072
0	4,784 ± 1,320	2,041 ± 429	1,117 ± 221	0.049	0.0001	0.049
1	11,920 ± 6,238	4,182 ± 1,067	811.2 ± 126	0.061	0.002	0.002
14	15,060 ± 5,178	9,595 ± 2,476	10,280 ± 2,155	0.288	0.317	0.834
60	ND	6,399 ± 1,318	7,696 ± 1,465	ND	ND	0.531

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MPs, microparticles; OFI, other febrile illness; DF, dengue fever; DHF, dengue hemorrhagic fever; ND, not done.

TABLE 4.

Absolute number of AnV⁺ MPs expressing the specific erythrocyte marker CD235 in three groups of patients^a

Day	CD235⁺ MPs (mean ± SD no. of events/μl)			P value
Day	OFI patients	DF patients	DHF patients	OFI vs DF patients	OFI vs DHF patients	DF vs DHF patients
−2	876.3 ± 217	1,138 ± 216	2,852 ± 928	0.449	0.199	0.134
−1	1,084 ± 218	1,946 ± 208	1,946 ± 208	0.372	0.056	0.0002
0	534.8 ± 123	1,065 ± 159	2,167 ± 260	0.053	0.001	0.002
1	857.3 ± 288	934.7 ± 147	2,092 ± 343	0.795	0.076	0.012
14	529.9 ± 104	752.9 ± 104	1,221 ± 185	0.029	0.029	0.042
60	ND	731.5 ± 92	926.4 ± 181	ND	ND	0.355

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MPs, microparticles; OFI, other febrile illness; DF, dengue fever; DHF, dengue hemorrhagic fever; ND, not done.

The number of platelets was also monitored during the acute phase of infection until recovery at day 14. As expected, the platelet count in the DHF group was distinctly reduced during the acute phase of illness (Fig. 4F; Table 5), consistent with the lower absolute number (Fig. 4B; Table 3) and percentage (Fig. 4D) of CD41a⁺ MPs compared with those in the DF and OFI groups. Platelet numbers returned to within the normal range at 2 weeks after defervescence (Fig. 4F; Table 5). To test whether the increase of CD235⁺ MPs exclusively observed in the DHF group was caused by an increase in the concentration of RBCs, RBC levels in all groups were analyzed over time. Unlike platelets, no statistically significant difference in RBC levels was observed between the DHF and DF groups during the acute phase of illness (day −2 to day 1) or during the convalescent phase (day 14) (Fig. 4G; Table 6). The elevated RBC levels in DHF patients, especially within the period from 24 h before to 24 h after defervescence (days −1, 0, and 1), were likely due to plasma leakage, thereby resulting in a relative rise in the concentration of RBCs due to hemoconcentration. In this study, all DHF patients had clinical evidence of plasma leakage, including a ≥20% rise of hematocrit and/or the accumulation of fluid in pleural or abdominal cavities, the two major WHO criteria for DHF (2, 3).

TABLE 5.

Number of circulating platelets in three groups of patients^a

Day	Mean ± SD platelet count (10³/μl)			P value
Day	OFI patients	DF patients	DHF patients	OFI vs DF patients	OFI vs DHF patients	DF vs DHF patients
−2	169 ± 14	172 ± 20	124 ± 15	0.917	0.146	0.072
−1	202 ± 14	133 ± 18	66 ± 9	0.053	<0.0001	0.001
0	199 ± 10	113 ± 16	44 ± 7	0.003	<0.0001	0.0002
1	233 ± 16	94 ± 13	43 ± 8	<0.0001	<0.0001	0.002
14	406 ± 29	463 ± 30	401 ± 26	0.248	0.912	0.138
60	ND	ND	ND	ND	ND	ND

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OFI, other febrile illness; DF, dengue fever; DHF, dengue hemorrhagic fever; ND, not done.

TABLE 6.

Number of circulating RBCs in three groups of patients^a

Day	Mean ± SD RBC counts (10⁶/μl)			P value
Day	OFI patients	DF patients	DHF patients	OFI vs DF patients	OFI vs DHF patients	DF vs DHF patients
−2	4.8 ± 0.2	5.3 ± 0.3	5.2 ± 0.1	0.185	0.103	0.817
−1	4.6 ± 0.1	5.2 ± 0.2	5.3 ± 0.1	0.158	0.004	0.524
0	4.6 ± 0.1	5.2 ± 0.2	5.5 ± 0.1	0.059	0.0005	0.221
1	4.5 ± 0.1	5.3 ± 0.2	5.5 ± 0.1	0.017	0.002	0.326
14	4.7 ± 0.1	5.1 ± 0.2	4.9 ± 0.1	0.152	0.105	0.492
60	ND	ND	ND	ND	ND	ND

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RBC, red blood cell; OFI, other febrile illness; DF, dengue fever; DHF, dengue hemorrhagic fever; ND, not done.

A simple linear regression showed that the log₁₀-transformed plot of platelet counts had a strong positive correlation with the log₁₀-transformed level of CD41a⁺ MPs (R = 0.702, 95% confidence interval [CI] = 0.627 to 0.764, n = 212, P < 0.0001; Fig. 4H) These data suggest that the lower platelet numbers (thrombocytopenia) induced by DENV may partly contribute to the reduced CD41a⁺ MP plasma levels observed in DF cases and especially in DHF cases (Fig. 4B and D). Conversely, there was a weak positive correlation between the log₁₀-transformed level of CD235⁺ MPs and the log₁₀-transformed level of RBC counts (R = 0.323, 95% CI = 0.197 to 0.438, n = 212, P < 0.0001; Fig. 4I), indicating that the increased concentration of RBCs (or hematocrit levels) due to plasma leakage did not primarily account for the highly elevated CD235⁺ MP levels detectable in DHF patients (Fig. 4C and E). Strikingly, the phenotypic characterization of circulating AnV⁺ MPs demonstrated that CD235⁺ MPs constituted the major population in the blood of the DHF group (≥50% of total AnV⁺ MPs) during the acute phase of illness, whereas during the convalescent phase, CD41a⁺ MPs were the most abundant AnV⁺ MPs (>65%) in the blood (Fig. 4J), and this result was similar for all groups. Indeed, CD41a⁺ MPs are the major MP population in the bloodstream of healthy individuals (5).

Elevation of CD235⁺ MPs in DENV infection is associated with DHF: a potential novel biomarker for plasma leakage.

The absolute counts and percentages of total AnV⁺ MPs, CD41a⁺ MPs, and CD235⁺ MPs were assessed to determine whether they were affected by disease severity in patients using a mixed model of multivariate statistical analysis because there were some missing data at some time points. Only data for patients with at least 3 consecutive measurements were included in the analysis. This resulted in data for 31 patients (17 DHF, 10 DF, and 4 OFI patients) and 107 observations for all parameters. The predicted total levels of AnV⁺ MPs, CD41a⁺ MPs, and CD235⁺ MPs were obtained from the quadratic mixed model against time (Fig. 5B, D, and F). For comparison, the mean and standard deviation (SD) of each variable over time, based on all available data, are also displayed (Fig. 5A, C, and E). DENV infection, but not disease severity, resulted in the reduction of the total levels of AnV⁺ MPs (DF versus OFI patients, P = 0.022; DHF versus OFI patients, P = 0.002; DF versus DHF patients, P = 0.237; Fig. 5B). However, in contrast to the total levels of AnV⁺ MPs, which were affected only by DENV infection, the level of CD41a⁺ MPs was significantly affected by both DENV infection and disease severity (DF versus OFI patients, P = 0.003; DHF versus OFI patients, P < 0.0001; DF versus DHF patients, P = 0.031; Fig. 5D). Furthermore, the level of CD41a⁺ MPs was significantly lower in DHF patients than in DF patients on every disease day except day −1 (day −2, P = 0.009; day −1, P = 0.062; day 0, P = 0.023; day +1, P < 0.001; Fig. 5D).

Interestingly, the level of CD235⁺ MPs was elevated only in DHF patients, while there was no difference in the level of CD235⁺ MPs between DF and OFI patients (DF versus DHF patients, P < 0.001; DF versus OFI patients, P = 0.307; DHF versus OFI patients, P < 0.001; Fig. 5F). The elevation of CD235⁺ MPs in DHF patients was found on every disease day during the acute phase of illness (for DF versus DHF patients, day −2, P = 0.005; day −1, P < 0.0001; day 0, P < 0.001; day +1, P = 0.032; for DHF versus OFI patients, day −2, P = 0.019; day −1, P = 0.010; day 0, P < 0.001; day +1, P < 0.0001; Fig. 5F). Notably, the levels of CD235⁺ MPs in DHF patients remained relatively constant during the course of the acute phase of illness (for DHF patients, day −2 versus day −1, P = 0.118; day −1 versus day 0, P = 0.951; day 0 versus day +1, P = 0.146; Fig. 5F).

The percentages of CD41a⁺ and CD235⁺ MPs among all AnV⁺ MPs was also evaluated by mixed model analysis (Fig. 5H and J). The mean and standard deviation of both parameters over time are also displayed (Fig. 5G and I). Similar to the absolute counts, the percentage of CD41a⁺ MPs was affected by both disease severity and infection with DENV (DF versus DHF patients, P = 0.039; DF versus OFI patients, P < 0.0001; DHF versus OFI patients, P < 0.0001; Fig. 5H), and differences between the DF and DHF groups were found on every disease day (day −2, P = 0.002; day −1, P = 0.020; day 0, P = 0.032; day +1, P = 0.009; Fig. 5H). While DHF patients had remarkably low platelet-derived MP levels, the percentage of CD235⁺ MPs, on the other hand, was exclusively elevated in the DHF group (DF versus DHF patients, P < 0.0001; DF versus OFI patients, P = 0.168; DHF versus OFI patients, P < 0.0001; Fig. 5J) and significantly different from that in the other groups on every disease day during the acute phase of illness (for DF versus DHF patients, day −2, P < 0.0001; day −1, P < 0.0001; day 0, P < 0.0001; day +1, P < 0.0001; for DHF versus OFI patients, day −2, P < 0.001; day −1, P < 0.0001; day 0, P < 0.0001; day +1, P < 0.0001; Fig. 5J). The percentage of CD235⁺ MPs in DHF patients was relatively constant during the course of illness (for DHF patients, day −2 versus day −1, P = 0.238; day −1 versus day 0, P = 0.131; day 0 versus day +1, P = 0.875; Fig. 5J).

The fact that the increase in CD235⁺ MPs was observed in DHF patients as early as day −2 and CD235⁺ MP levels were constantly elevated during the febrile phase led us to reason that the earliest single measurement of CD235⁺ MPs might be used as a biomarker to predict the severity of illness in dengue patients or the development of DHF. To test this, the earliest measurements (median disease day of sample collection = day −2; range = day −6 to day −1; Table 7) of the absolute count and percentage of CD235⁺ MPs prior to defervescence were collected from all patient groups and assessed through receiver operating characteristic (ROC) analysis. Interestingly, the areas under the ROC curves (AUCs) indicated a good prediction performance, with AUCs being equal to 0.824 (95% CI = 0.693 to 0.954) and 0.903 (95% CI = 0.808 to 0.997) for the absolute count (Fig. 5O) and percentage (Fig. 5P) of CD235⁺ MPs, respectively. If a cutoff absolute count of CD235⁺ MPs was set at 2.982 on a log₁₀ scale, it would predict the development of DHF with 94.7% sensitivity, 60.0% specificity, a 69.2% positive predictive value, and a 92.3% negative predictive value. If a cutoff percentage of CD235⁺ MPs was set at 29.67%, it would predict severity with 84.2% sensitivity, 80.0% specificity, an 84.2% positive predictive value, and an 87.5% negative predictive value. Because of the high negative predictive values of the absolute count and the percentage of CD235⁺ MPs at the cutoffs described above, the measurement of CD235⁺ MPs levels at the early febrile phase of dengue virus infection may potentially be used to screen out a patient with mild disease (DF) who may not require hospitalization.

TABLE 7.

Characteristics of the earliest samples from each patient included in the ROC analysis of CD235⁺ MPs as a biomarker^a

Characteristic	Result for the following patients:
Characteristic	OFI patients (n = 6)	DF patients (n = 14)	DHF patients (n = 19)
Absolute CD235⁺ MP count	743.7 (691.6, 1,099.5)	892.3 (533.0, 1,324.8)	1,708.7 (1,147.2, 2,340.8)
Log₁₀ (absolute CD235⁺ MP count)	2.87 (2.84, 3.03)	2.95 (2.73, 3.12)	3.23 (3.06, 3.37)
% CD235⁺ MPs	7.98 (7.18, 10.05)	19.97 (11.70, 28.97)	45.45 (35.14, 65.10)
Day of disease that samples were collected	−2 (−2.75, −1.25)	2 (−2, −1)	−2 (−2, −1)

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Data are presented as the median (25th, 75th percentile). OFI, other febrile illness; DF, dengue fever; DHF, dengue hemorrhagic fever; MPs, microparticles.

The changes of the other parameters (RBC and platelet counts) over time were also analyzed with mixed-effects models (Fig. 5L and N). The mean and standard deviation of each parameter over time were also displayed (Fig. 5K and M). Platelet counts were affected by disease severity and DENV infection (DF versus DHF patients, P < 0.001; DF versus OFI patients, P = 0.009; DHF versus OFI patients, P < 0.0001; Fig. 5L). However, on day −2 the platelet count was not significantly different among the three patient groups (DF versus DHF patients, P = 0.102; DF versus OFI patients, P = 0.508; DHF versus OFI patients, P = 0.065; Fig. 5L). The platelet counts of DHF patients started to be significantly lower than those of the other groups on day −1 (DF versus DHF patients, P = 0.002; DHF versus OFI patients, P < 0.0001), while the platelet counts of DF patients started to differentiate from those of OFI patients on day 0 (P = 0.006). There was no difference in the dynamics of RBC counts throughout the acute phase, except between DHF and OFI patients (DF versus DHF patients, P = 0.349; DF versus OFI patients, P = 0.131; DHF versus OFI patients, P = 0.026; Fig. 5N).

Circulating platelet-derived MPs and red blood cell-derived MPs carry DENV antigens on their surfaces.

In vitro, DENV-infected cells generated MPs that contained E and NS1 antigens on their surfaces (Fig. 3). To evaluate the relevance of these findings in dengue patients, circulating CD41a⁺ MPs and CD235⁺ MPs were analyzed for DENV E and NS1 antigen content by immunofluorescence staining (Fig. 6). Interestingly, the absolute numbers of NS1-positive (NS1⁺) CD235⁺ MPs but not the absolute numbers of E-positive (E⁺) CD235⁺ MPs (Fig. 6C) were significantly higher in the DHF group during the acute phase (for day −1, DHF versus DF patients, P = 0.012; DHF versus OFI patients, P = 0.048; for day 0, DHF versus DF patients, P = 0.052; DHF versus OFI patients, P = 0.031; for day 1, DHF versus DF patients, P = 0.008; DHF versus OFI patients, P = 0.037; Fig. 6D). However, there was no significant difference in the absolute counts of CD41a⁺ MPs carrying DENV E (Fig. 6A) or the absolute counts of CD41a⁺ MPs carrying NS1 (Fig. 6B) in the DHF group from those in DF and OFI patients.

FIG 6 — Circulating MPs derived from platelets and red blood cells harbor DENV antigens on their surfaces. Platelet-derived MPs (AnV⁺ CD41a⁺) and erythrocyte-derived MPs (AnV⁺ CD235⁺) were further analyzed for DENV E (A and C) and NS1 (B and D) expression on their surfaces. The absolute numbers of E⁺ CD41a⁺ MPs (A), NS1⁺ CD41a⁺ MPs (B), E⁺ CD235⁺ MPs (C), and NS1⁺ CD235⁺ MPs (D) were quantified by immunofluorescent labeling and flow cytometric analysis. Each dot represents the absolute number of each population of MPs for an individual patient on different disease days. Solid bars indicate the means. Asterisks denote the means that are statistically significantly different between the indicated groups of patients (*, P < 0.05; **, P < 0.001).

Complement activation of NS1–anti-NS1 ICs triggers MP shedding from red blood cells.

Next, we explored why and how RBCs of DHF patients shed more MPs, as shown by the elevation of AnV⁺ CD235⁺ MPs (Fig. 4 and 5). To maintain homeostasis, one important function of RBCs is to trap potentially harmful immune complexes (ICs) in the circulation. Complement receptor type 1 (CR1) on RBCs binds to complement fragment-opsonized ICs and transfers them to mononuclear phagocytes in the liver and spleen (31, 32). Circulating ICs have been detected in DHF patient serum during the acute phase (33, 34). Additionally, high levels of NS1 have been found in dengue patients, and they were positively correlated with disease severity (16, 35). We thus reasoned that, in patients with DHF/DSS, which is mostly associated with a secondary infection (3), circulating ICs formed between the virion or NS1 and their specific antibodies should increase dramatically due to the presence of an extensive amount of viral antigens in the bloodstream (viremia and NS1 antigenemia) together with a rapid rise in the levels of antivirion (36) and anti-NS1 (37) antibodies, followed by an anamnestic secondary immune response. High levels of circulating ICs in DENV infection may overburden the RBCs' function of IC clearance and, consequently, trigger those RBCs to shed MPs. We hypothesized that RBC vesiculation may occur either during the processes of immune adherence (binding of complement-opsonized ICs to CR1 on RBCs) or during the transfer of ICs from RBCs to phagocytic cells in the liver and spleen, thereby resulting in increased RBC-derived MPs in DHF patients (Fig. 4 and 5).

To test this hypothesis, we developed an in vitro model of IC transfer and investigated whether this process triggers RBCs to shed MPs (Fig. 7). Interestingly, incubation of RBCs with ICs (consisting of purified NS1 plus IgG from pooled dengue patient immune serum [PCS]) and complement led to not only the binding of C3d-opsonized ICs to CR1 (Fig. 7B) but also a slight increase of CD235⁺ MP generation (Fig. 7E). Inhibition of complement activation by EDTA or pretreatment of RBCs with function-blocking anti-CR1 MAb clone 3D9 (38), but not with an isotype control, prior to the addition of a mixture of NS1 and PCS abrogated the binding of ICs to CR1, as determined by negative C3d staining (Fig. 7A and B) and MP shedding from RBCs (P = 0.013; Fig. 7E). Low levels of CD235⁺ MP generation were also observed when RBCs were incubated with serum alone or with a mixture of NS1 and purified IgG from pooled DENV-nonimmune donors (PND) (Fig. 7E).

To mimic the phenomenon of IC transfer, autologous peripheral blood mononuclear cells (PBMCs) were added to RBCs carrying complement-opsonized ICs. As expected, C3d-opsonized NS1–anti-NS1 ICs deposited on CR1 of RBCs were completely removed after PBMC exposure (Fig. 7D). More importantly, this process dramatically augmented MP shedding from RBCs (Fig. 7F). However, the MP shedding from PBMCs was not changed, as determined by the detection of antibodies against common leukocyte surface marker CD45 (Fig. 7G). Taken together, these results suggest that removal of complement-opsonized ICs bound to RBCs via CR1 by mononuclear cells can trigger RBCs to shed MPs and might explain the elevated CD235⁺ MP levels in DHF patients.

A decrease in platelet-derived MPs in dengue patients is associated with a bleeding tendency.

Hemorrhagic manifestations are commonly associated with DENV infection (2). They can potentially complicate clinical management and cause significant morbidity and mortality, especially when occurring massively in the gastrointestinal tract (3). Despite being more frequently associated with DHF, a bleeding tendency can independently occur in the absence of plasma leakage, i.e., in DF patients (3). Many factors have been shown to contribute to bleeding in DENV infection, including thrombocytopenia, reduced thrombin formation, and increased fibrinolysis activity (39, 40), yet the pathogenesis of bleeding complications is not fully understood. To further investigate whether circulating MPs influence the outcome of bleeding, all dengue virus-infected patients were separated into two groups according to their history of bleeding, i.e., a bleeding group (total number of patients = 18; number of DHF patients = 13, number of DF patients = 5) and a no-bleeding group (total number of patients = 25; number of DHF patients = 11, number of DF patients = 14), regardless of plasma leakage. The dengue patients who had evidence of bleeding on any day during febrile illness were categorized into the bleeding group. Bleeding episodes included nose or gum bleeding, ecchymosis, gastrointestinal hemorrhage (blood in vomitus or in stool), and hypermenorrhea. Platelet counts (Fig. 8A and B), total AnV⁺ MP levels (Fig. 8D and E), CD41a⁺ MP levels (Fig. 8G and H), and CD235⁺ MP levels (Fig. 8J and K) on each disease day were plotted according to bleeding episodes. Declining levels of platelets, total AnV⁺ MPs, and CD41a⁺ MPs were displayed more prominently in the bleeding group over the course of the acute phase of illness, reaching the lowest values on day +1, and were statistically significantly different between the two groups (platelets, P = 0.021; total AnV⁺ MPs, P = 0.013; CD41a⁺ MPs, P = 0.008). On the contrary, in the no-bleeding group, the levels of those parameters started rising on day +1 and returned to normal on day 14, when both groups exhibited similar levels (P > 0.05). Interestingly, while the levels of CD235⁺ MPs were clearly different between DF and DHF patients (Fig. 4C and 5E and F), it was not affected by hemorrhagic manifestations (Fig. 8J and K).

Fig 8 — Blood levels of platelet-derived MPs but not red blood cell-derived MPs correspond to procoagulant MP activity and bleeding predisposition. DENV-infected patients with at least one episode of bleeding and patients with no mucosal bleeding during the admission were classified as the bleed and no-bleed groups, respectively. Bleeding episodes included epistasis, nose bleeding, gastrointestinal hemorrhage, and hypermenorrhea. (A to Q) The numbers of platelets (A), total AnV⁺ MPs (D), CD41a⁺ MPs (G), and CD235⁺ MPs (J) of DENV-infected patients with and without bleeding episodes during the acute phase and at day 14 after defervescence (convalescent phase) are depicted. The means and standard errors of log₁₀-transformed values were analyzed for platelets (B), total AnV⁺ MPs (E), CD41a⁺ MPs (H), and CD235⁺ MPs (K). Predicted platelets (C), total AnV⁺ MPs (F), CD41a⁺ MPs (I), and CD235⁺ MPs (L) in each group were also analyzed by the use of quadratic mixed-effects models. The levels of platelets (M), total AnV⁺ MPs (N), CD41a⁺ MPs (O), CD235⁺ MPs (P), and procoagulant MPs (Q) in DENV-infected patients with and without bleeding episodes on the day of the platelet count nadir were plotted. Asterisks denote the means that are statistically significantly different between the indicated groups of patients (*, P < 0.05; **, P < 0.001; N.S., not significant). (R to U) The correlation coefficients between the absolute counts of procoagulant MPs and the numbers of CD41⁺ MPs (R and S) and CD235⁺ MPs (T and U) in patients with (R and T) or without (S and U) bleeding episodes were calculated. The linear regression lines, correlation coefficients (R), and P values are presented in the graphs.

To obtain more accurate results for the analysis of these continuous variables over time, mixed-effects models were examined to assess whether the overall changes in the levels of these factors were related to the outcome of bleeding (Fig. 8C, F, I, and L). Using the quadratic models, 11 bleeding and 16 no-bleeding patients whose data were available on at least three consecutive days were included in the analysis. In the models, the outcome of bleeding and the disease day were treated as fixed effects, while each case was treated as a random effect. Interestingly, the pattern of the changes in platelet counts over time was not significantly different between dengue patients with bleeding and those without bleeding (P = 0.132; Fig. 8C). Likewise, bleeding status did not affect the levels of total AnV⁺ MPs during the course of the acute phase of illness (P = 0.272; Fig. 8F). However, similar to the analysis of the mean (Fig. 8A and D), the multivariate analysis showed that platelet counts (P = 0.032; Fig. 8C) and total AnV⁺ MP levels (P = 0.003; Fig. 8F) were significantly different between the two groups only on day +1. Interestingly, the reduction of CD41a⁺ MP levels over time in the bleeding group was significantly different from that in the patients without bleeding (P = 0.043; Fig. 8I). Moreover, the levels of CD41a⁺ MPs could be used to differentiate between the two groups on almost every disease day except day −1 (day −2, P = 0.047; day −1, P = 0.176; day 0, P = 0.035; day +1, P < 0.0001; Fig. 8I). Nonetheless, no differences in the levels of CD235⁺ MPs were observed among the different groups of patients (P = 0.271; Fig. 8L). Overall, these results imply that the levels of CD41a⁺ MPs but not those of CD235⁺ MPs might be associated with the outcome of bleeding.

The surface of CD41a⁺ MPs, the most abundant MPs in healthy human blood (5), has approximately 50- to 100-fold more procoagulant activity than the surface of activated platelets (41). They provide a low-grade procoagulant state to maintain homeostasis in the blood of healthy individuals (5, 42). We reasoned that under circumstances in which the number of platelets significantly decreases, like in dengue, a small change in the level of circulating CD41a⁺ MPs, which could also be caused by factors other than thrombocytopenia, could largely influence the tendency to bleed. To address this, blood samples from each patient taken on the day of the platelet level nadir were examined to determine whether decreased levels of CD41a⁺ MPs directly affected procoagulant MP activity in the blood of those patients who had one or more bleeding episodes during the acute phase of illness. As expected, procoagulant MP activity (P = 0.006; Fig. 8Q) along with total AnV⁺ MP levels (P = 0.027; Fig. 8N) and CD41a⁺ MP levels (P = 0.005; Fig. 8O), but not CD235⁺ MP levels (P = 0.807; Fig. 8P), were significantly lower in the bleeding group than in patients without any bleeding episodes during the acute phase of febrile illness. Of note, the platelet level nadir was also lower in the bleeding group, but the difference did not reach statistical significance (P = 0.054; Fig. 8M). All parameters measured during the convalescent phase, including procoagulant MP activity, were indistinguishable between the two groups (Fig. 8A, D, G, J, and Q). The levels of CD41a⁺ MPs might reflect the procoagulant MP activity in the blood of dengue patients, especially in the bleeding group, as a weak positive correlation was observed (for the bleeding group, R = 0.436 and P < 0.001 [Fig. 8R]; for the no-bleeding group, R = 0.252 and P = 0.025 [Fig. 8S]), but this association was not observed with CD235⁺ MPs (for the bleeding group, R = −0.087 and P = 0.492 [Fig. 8T]; for the no-bleeding group, R = −0.067 and P = 0.559 [Fig. 8U]). These data suggest that platelet-derived MPs but not red blood cell-derived MPs might contribute to the bleeding tendency that occurs in dengue patients.

DISCUSSION

MPs have been recognized to play important roles in many biological processes. In response to various stimuli, including virus infection, cells shed parts of their plasma membrane and cytoplasmic content into small vesicular structures with different sizes, characteristics, and functions. Herein we show that DENV infected various kinds of susceptible cells in vitro, resulting in apoptotic death and MP release. These MPs harbored viral E and NS1 on their surfaces. Electron microscopy techniques, including negative staining and immunogold labeling, revealed greater detail about the MPs generated from hepatocytes (one of the major target cells for DENV replication in vivo [43]), which were morphologically distinct from the virions produced in the same culture system. Nevertheless, the molecular mechanism underlying MP generation after DENV infection is still unknown, that is, whether it is directly linked to the process of cells undergoing apoptosis or it is a consequence of cell activation in response to viral infection leading to increased intracellular calcium, which in turn causes membrane vesiculation.

It is still questionable as to what could be the significance of this phenomenon in DENV infection. A previous study has also described membrane vesicular structures within the cytoplasm and in the culture supernatants of DENV-infected cells and has proposed their function as being sites of virus replication (44). Alternatively, DENV may use these secreted MPs as a tool to escape from host immune surveillance and disseminate infection. In fact, this immune evasion strategy via secreted MPs has been previously described for other viruses. Microvesicles released from infected cells facilitate the spread of human immunodeficiency virus and herpes simplex virus to neighboring uninfected cells by transferring either viral constituents or the host's membrane proteins required for viral entry to other null cells, thereby increasing the number of susceptible cells (45 –47).

Changes in the levels and origin profiles of circulating MPs have been described in many pathological conditions. Our results from the ex vivo analysis of MPs in the circulation of DENV-infected patients were initially surprising. The total numbers of MPs identified by AnV labeling of surface-exposed PS and platelet-derived (AnV⁺ CD41a⁺) MPs were decreased during the acute phase. Thrombocytopenia (in which the platelet number in blood is decreased), a key clinical manifestation of DENV infection (2), may contribute to these surprising results. In support of this, DHF patients whose platelet counts were significantly lower than those of DF patients, especially at the critical period (days 0 and +1; Fig. 4F), also had significantly lower platelet-derived MP levels (Fig. 4B). In fact, platelets are the major source of MPs in healthy individuals (5). In our study, a strong positive correlation between the numbers of platelets and CD41a⁺ MPs was observed (Fig. 4H). Notably, various factors could enhance MP shedding from platelets during the acute phase of DENV infection, such as platelet activation (48), platelet apoptosis (49), and direct infection of platelets by DENV (50). Indeed, a small number of circulating AnV⁺ CD41a⁺ MPs carried DENV E and NS1 antigens, similar to the findings for infected cell-derived MPs, as demonstrated in our in vitro model of DENV infection. However, the existence of DENV antigen-carrying CD41a⁺ MPs in the blood during the acute phase of illness (Fig. 6A and B) was not direct evidence for DENV infection of platelets or megakaryocytes (which also express CD41a). Alternatively, FcγII receptors on the surface of platelets might bind to DENV-antibody complexes, as previously shown by Wang et al. (51), or NS1-antibody complexes, resulting in shedding of E⁺ or NS1⁺ CD41a⁺ MPs, respectively, without actual DENV infection.

In contrast to our findings, a previous study by Hottz et al. reported a higher number of platelet-derived MPs in dengue patients than in healthy controls (52). This discrepancy may be due to distinct characteristics of dengue patients between the two studies, including age (children versus adults), the degree of thrombocytopenia, and disease severity, as well as the method for MP enumeration. The protocol used by Hottz et al. (52) to obtain the number of circulating CD41a⁺ MPs per 100 platelets did not represent the absolute number of MPs in the patients' circulation. Using this method of flow cytometric analysis, the plasma platelet concentration could influence the number of MP counts; i.e., for samples with higher platelet concentrations, like those from healthy controls, the amount of time required to acquire platelet counts of 100 is less, and thus, MP counts are lower than those in dengue patients, whose circulating platelet numbers are usually low during the acute phase of illness even when they have mild diseases (2). The present study provided more accurate information on the total circulating MP concentration by using TruCount beads to obtain the absolute number of MPs per μl plasma (53). Moreover, comparative analysis among groups of patients (DF patients versus DHF patients versus nondengue OFI patients) at multiple time points has revealed similar dynamic changes between platelet-derived MP levels (Fig. 4B) and platelet counts (Fig. 4F), which are decreased over the course of the acute phase of illness (for day −2 to day +1, DHF patients < DF patients < OFI patients) and rise back to within normal limits during the recovery period and are equal in DHF, DF, and OFI patients on day 14 and day 60.

The functional significance of decreased AnV⁺ CD41⁺ MP levels in DENV infection may relate to the procoagulant property of the negatively charged PS exposed on MPs, which provides the binding site for coagulation factors and serves as a platform for assembly of the prothombinase complex, resulting in the amplification of prothrombin cleavage and thrombin production (54). Circulating CD41a⁺ MPs possess up to 100-fold more procoagulant activity than activated platelets (41) and help to maintain homeostasis in healthy individuals (5, 42). Increased amounts of circulating platelet-derived MPs are linked to hypercoagulable states in several pathological conditions (4, 55 –58). Conversely, patients with an inherited bleeding disorder, Scott syndrome, develop severe hemorrhagic manifestations due to a defect in scramblase activity, leading to the lack of surface exposure of PS (thus resulting in an inadequate ability to promote a procoagulable state) and to the inability of platelets and other blood cells to generate MPs (59). Another bleeding disorder that results in a decreased capacity of platelets to shed MPs has also been described (60). In the same line of thought, our results showed that the levels of CD41a⁺ MPs reflected the procoagulant activity of circulating MPs, especially in the bleeding group. Therefore, together with thrombocytopenia, diminished total circulating AnV⁺ MP levels and especially AnV⁺ CD41a⁺ MP levels might also contribute to abnormal hemostasis in some cases of DENV infection.

Surprisingly, our ex vivo analysis of circulating MPs showed a significant increase in the levels of AnV⁺ CD235⁺ MPs in DHF patients (but not DF and OFI patients) during the acute phase (Fig. 4 and 5). However, the role of RBCs in DENV infection has not been previously described, and importantly, RBCs could not support DENV replication. Of note, the increase in AnV⁺ CD235⁺ MP levels was not due to a difference in RBC levels, which indicates the degree of hemoconcentration. One of the key functions of RBCs is to maintain homeostasis by trapping potentially harmful ICs which could deposit in tissues and induce injury. This is accomplished by complement opsonization, followed by binding to complement receptor type 1 (CR1), expressed on the surface of RBCs, the so-called immune adherence process (31, 61, 62). ICs bound to RBCs are then transferred to phagocytic cells in the liver and spleen (31, 62). The finding that the level of NS1 containing AnV⁺ CD235⁺ MPs was significantly increased in DHF patients (Fig. 6D) prompted us to hypothesize that ICs that form between NS1 and its specific antibody are involved in the MP shedding process of RBCs.

We have shown that removal of complement-opsonized NS1–anti-NS1 ICs bound to erythrocytes by mononuclear cells triggered in vitro RBC-derived MP shedding. These results suggest that the IC transfer mechanism might cause RBC vesiculation in dengue patients. However, the precise cellular mechanisms involved in the transfer of RBC-bound ICs from RBCs to macrophages or phagocytic cells and the generation of CD235⁺ MPs during this process remain to be elucidated. The transfer mechanism could be driven by a higher number of CR1 copies on the monocyte than on the RBCs (63) or by the close juxtaposition of the macrophage to the RBC-bound ICs that leads to macrophage-associated protease cleavage of CR1 and a release of the IC from the RBC (64). It is possible that instability of RBC membranes may occur during transfer of CR1-bound ICs from erythrocytes to acceptor phagocytic cells via Fc receptors (64, 65) and thus trigger RBCs to shed MPs.

Our data on MP shedding from RBCs in response to the removal of surface-bound ICs by mononuclear cells point to the possibility of increased circulating red blood cell-derived MPs (RMPs) in other IC diseases, such as systemic lupus erythematosus (SLE). Increased levels of MPs from various cellular origins, including platelets, leukocytes, and endothelial cells, but not from RBCs have previously been reported (66 –69). The reason why DHF patients but not SLE patients showed a significant increase in AnV⁺ CD235⁺ MP levels remains to be answered. The level of circulating MPs likely depends on the balance between their rates of generation and clearance. It is possible that the transport of ICs for clearance in the liver and spleen in dengue patients might be defective due to the pathology of those organs directly caused by DENV infection. Indeed, a recent pathology study at autopsy of tissues obtained from patients who have died from DHF/DSS, led by our group, has demonstrated that the liver and spleen are the two major organs found to be heavily infected (43). Alternatively, the amount and nature of ICs, including their size and whether they are soluble or cell associated, determine the magnitude and efficiency of complement fixation (70) and may also influence the degree of MP shedding from RBCs. The marked inefficiency of complement C5 cleavage relative to the efficiency of C3 cleavage and the inefficiency of C5b-9 generation during complement activation by soluble ICs compared with the efficiency of C5b-9 generation by ICs on particles or cells likely result in the detection of lower levels of C5a and soluble C5b-9 (SC5b-9) complexes in the plasma of patients with ongoing, complement-consuming IC diseases like SLE (33). In support of this idea, in contrast to SLE patients, high levels of circulating SC5b-9 were observed in DENV-infected patients, especially in those with DHF/DSS (16), suggesting more efficient complement activation and possibly also differences in the nature/characteristics of complement activators generated during the acute phase of DENV infection. Consistent with these findings, it has been demonstrated that the large antigen-antibody complexes formed on the surface of infected cells efficiently fix complement reaching the terminal pathway, as determined by the generation of fluid-phase SC5b-9 as well as the deposition of C5b-9 membrane attack complexes on cell surfaces (16, 20). Taken together, these factors may also contribute to the discrepancy between the RMP levels observed in dengue patients and those observed in SLE patients.

The physiological roles of RMPs have not been well elucidated. Many physiological and pathological conditions have been shown to induce RMP generation. During their life span, RBCs lose approximately 20% of their hemoglobin content through vesicle formation (71). Stored RBCs also lose their membrane integrity, thereby resulting in hemolysis and RMP generation that may contribute to complications associated with transfusion (66, 72, 73). Additionally, RMP formation could be triggered with different types of stimuli, such as shear stress, complement attack via the insertion of the terminal complement C5b-9 in the membrane of RBCs, oxidative stress, and proapoptotic stimulations (74 –76). Increased RMP levels have been documented in different pathological conditions, including sickle cell disease (77), hereditary hemolytic anemia (78), paroxysmal nocturnal hemoglobinuria (73), atherosclerosis (79), and malaria (9, 80). RMPs might also be involved in immunomodulation by inhibiting macrophage release of proinflammatory cytokines (81) or, conversely, acting as potent stimulators of cells of the innate immune system (82). Nevertheless, the functional roles of RMPs in dengue require further investigation.

Although the biological process behind RMP elevation in DHF patients is not yet fully understood, RMP elevation itself has a strong potential for use as a biomarker for differentiating potentially severe dengue disease (DHF) from milder dengue fever. During the early febrile phase, clinical signs and symptoms as well as currently available laboratory tests cannot differentiate the two clinical syndromes. We have shown that the elevation of RMP levels occurred at a very early time point of the disease and the elevated RMP level remained constant throughout the acute phase. Consequently, an early measurement of RMP levels is potentially useful to identify patients who will subsequently develop plasma leakage (DHF) as well as prevent patients with mild disease from unnecessary hospitalization. This is important, as during a dengue epidemic where limited resources are available, overhospitalization has been shown to contribute to a significant increase in morbidity and mortality, especially in countries where dengue is endemic (83, 84).

In summary, the present study has provided some evidence of the roles of MPs in dengue pathogenesis. MPs generated from erythrocytes and platelets comprised two major populations in the circulation of dengue patients. Elevated levels of RMPs directly correlated with DENV disease severity, whereas a significant decrease in platelet-derived MP levels was associated with a bleeding tendency. Removal of complement-opsonized NS1–anti-NS1 ICs bound to erythrocytes by mononuclear cells triggered erythrocytes to shed MPs in vitro, a process that might explain the increased RMP levels in severe dengue. RMPs are a potential biomarker to predict the development of DHF.

ACKNOWLEDGMENTS

This work was supported by a Mahidol University research grant (to P.A.) The clinical cohorts were supported by the Office of the Higher Education Commission and Mahidol University under the National Research Universities Initiative (to P.M.). P.A. has been supported by a Siriraj Chalermprakiat grant and a research lecturer grant, Faculty of Medicine Siriraj Hospital, Mahidol University. N.P. and S.T. are Ph.D. scholars in the Royal Golden Jubilee Ph.D. Program.

We thank Ali Amara for kindly supplying wild-type 293T cells and 293T cells stably expressing TIM-1, Chunya Puttikhunt and Watchara Kasinrerk for providing anti-DENV NS1 MAb, Bunpote Siridechadilok for advise and help in electron microscopy experiments, Prapat Suriyaphol for constructive comments on data analysis, and Richard Hauhart and John Atkinson for helpful discussion and critical reading of the manuscript.

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Microparticles Provide a Novel Biomarker To Predict Severe Clinical Outcomes of Dengue Virus Infection

Nuntaya Punyadee

Dumrong Mairiang

Somchai Thiemmeca

Chulaluk Komoltri

Wirichada Pan-ngum

Nusara Chomanee

Komgrid Charngkaew

Nattaya Tangthawornchaikul

Wannee Limpitikul

Sirijitt Vasanawathana

Prida Malasit

Panisadee Avirutnan

Roles

ABSTRACT

INTRODUCTION

MATERIALS AND METHODS

Cells and viruses.

DENV infection and MP isolation.

Analysis of MPs generated from DENV-infected cells.

Transmission electron microscopy.

Negative staining.

Immunogold labeling.

In vitro experiments with immune complexes (ICs).

In vitro analysis of IC opsonization and vesiculation.

Patients.

TABLE 1.

Blood sample collections.

Labeling of plasma MPs.

MP procoagulant activity.

Statistical analysis.

RESULTS

DENV infection leads to apoptotic death and MP shedding.

FIG 1.

Fig 2.

MPs released from DENV-infected cells express viral antigens on their surfaces.

FIG 3.

Circulating MPs in dengue patients.

Fig 4.

TABLE 2.

TABLE 3.

TABLE 4.

TABLE 5.

TABLE 6.

Elevation of CD235+ MPs in DENV infection is associated with DHF: a potential novel biomarker for plasma leakage.

FIG 5.

TABLE 7.

Circulating platelet-derived MPs and red blood cell-derived MPs carry DENV antigens on their surfaces.

FIG 6.

Complement activation of NS1–anti-NS1 ICs triggers MP shedding from red blood cells.

FIG 7.

A decrease in platelet-derived MPs in dengue patients is associated with a bleeding tendency.

Fig 8.

DISCUSSION

ACKNOWLEDGMENTS

REFERENCES

ACTIONS

PERMALINK

RESOURCES

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Elevation of CD235⁺ MPs in DENV infection is associated with DHF: a potential novel biomarker for plasma leakage.