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The Journal of Immunology, 2008, 181, 5865 -5874
Copyright © 2008 by The American Association of Immunologists, Inc.
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A Complex Interplay among Virus, Dendritic Cells, T Cells, and Cytokines in Dengue Virus Infections1

Wanwisa Dejnirattisai*, Thaneeya Duangchinda{dagger}, Chen-Lung Steve Lin{ddagger}, Sirijitt Vasanawathana§, Meleri Jones, Michael Jacobs, Prida Malasit{dagger},||, Xiao-ning Xu#, Gavin Screaton2,* and Juthathip Mongkolsapaya2,*,||

* Department of Immunology, Division of Medicine, Hammersmith Hospital, Imperial College, London, United Kingdom; {dagger} Medical Biotechnology Unit, National Center for Genetic Engineering and Biotechnology, National Science and Technology Development Agency, Pathumthani, Thailand; {ddagger} Division of Surgery, Oncology, Reproductive Biology and Anesthetics, Faculty of Medicine, Hammersmith Hospital, Imperial College, London, United Kingdom; § Pediatric Department, Khon Khan Hospital, Ministry of Public Health, Khon Kaen, Thailand; Department of Infection, University College London, Hamstead Campus, Rowland Hill Street, London, United Kingdom; || Medical Molecular Biology Unit, Faculty of Medicine, Siriraj Hospital, Mahidol University, Bangkok, Thailand; and # Medical Research Council Human Immunology Unit, Weatherall Institute of Molecular Medicine, John Radcliffe Hospital, University of Oxford, Oxford, United Kingdom


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Severe dengue virus (DV) infections can cause the life-threatening condition dengue hemorrhagic fever, which is characterized by a severe plasma leak, thrombocytopenia, hemorrhage, and, in severe cases, circulatory collapse and death. There is now much evidence that pre-existing immunity to DV can enhance disease when an individual becomes infected on a second or sequential occasion. It has been shown that in contrast to infected dendritic cells (DC), noninfected bystander DC underwent maturation in dengue infection. In this study, we show that TNF-{alpha} and type I IFN contribute to the maturation of bystander DC, whereas the inhibition of DV-infected DC maturation can be overcome by activated T cells. Furthermore, IFN-{gamma}-inducible chemokines, CXCL9, 10, and 11 produced by infected DC are greatly amplified in the presence of DV-specific T cells. The chemokine secretion is also enhanced in coculture of HUVEC with either DV-infected DC or activated T cells. Finally, we found a close correlation between the serum level of these three chemokines and disease severity.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Dengue virus (DV)3 infection is a major emerging disease of tropical and subtropical countries, and rates of infection and hemorrhagic fever are increasing at an alarming rate. It is estimated that nearly half of the world’s population is at risk of infection and up to 50 million people are infected each year with frequent epidemic activity in Southeast Asia, South America, and the Western Pacific regions (1). DV are positive-strand RNA viruses belonging to the Flaviviridae family. Infection occurs following the bite of an infected mosquito, usually Aedes aegypti. Many infections are asymptomatic, whereas in the clinically overt group symptoms begin after an incubation period of 5–7 days. These include high fever, headache, rash, and bone and muscle pains. In the majority of cases, symptoms will lessen as the fever remits and patients will recover; this relatively uncomplicated illness is known as dengue fever (DF). In a proportion of cases (5–30%), the disease is more severe, with evidence of plasma leak, thrombocytopenia, bleeding, and shock. These cases are known as dengue hemorrhagic fever (DHF), which carries a high mortality without expert clinical intervention.

Four distinct serotypes of DV cocirculate in endemic areas, and these four viruses show substantial sequence divergence of ~30%. Immunity to one serotype does not provide protection against infection by one of the others. There is now very good epidemiological evidence that the majority of DHF cases occur in individuals who suffer secondary or sequential DV infections. This implies that immunity to the previously encountered virus is not only ineffective at preventing a secondary infection, but may also enhance disease (2, 3). Furthermore, serious complications occur in DHF at the time of defervescence as virus is cleared from the blood, and vascular leak is transient, implying that DHF is primarily an immunopathology rather than direct virus-induced cytopathology (4).

Much work on DV immunopathogenesis has been performed, and a number of hypotheses have been put forward to explain both the enhancement of disease in secondary infection and the pathophysiology of the vascular leak, which is the hallmark of DHF. There is no animal model of DHF, and almost all in vitro work has been limited to studying the effect of DV infection on a single cell type. In this study, we examine the interaction between the virus, dendritic cells (DC), T cells, and endothelial cells, and find a complex interplay leading to amplification of chemokine release. We further studied chemokine production in patients with DV and found a close correlation between disease severity and the serum level of the IFN-{gamma}-inducible chemokines CXCL9, 10, and 11.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Samples

Blood samples were taken from 50 patients after patient consent and approval from the ethical committee of Khonkaen Hospital, Thailand. Acute DV infection was identified by RT-PCR-based DV gene identification or DV-specific IgM capture ELISA. Disease severity was classified according to World Health Organization criteria. The day of defervescence was defined as day 0, the day before defervescence as day –1, the day after defervescence as day +1, and so forth.

Of the patients enrolled in the study, 14 patients were classified as DF, 13 as DHF I, 13 as DHF II, and 10 as DHF III. Blood samples were collected in 5 mM EDTA, and plasma was separated by centrifugation, aliquoted, and stored at –70°C. PBMCs were isolated from whole blood by Ficoll-Hypaque density-gradient centrifugation and cryopreserved until tested.

Abs and cells

Abs against CD83, CD80, CD86, HLA-DR, CD107a, CD8, CD38, IFN-{gamma}, anti-TNF-{alpha}, and CXCL10 (BD Pharmingen); CXCR3 (R&D Systems); and human IgG (DakoCytomation) were used. Anti-TNF-{alpha} blocking Ab were provided by A. Cope (Kennedy Institute of Rheumatology, Imperial College, London, U.K.) and anti-IFN-{alpha}/βR2 neutralizing Ab were purchased from PBL IFN Source. Dengue hyperimmune human serum (hemagglutination titer ≥1/25,600) was provided by Armed Forces Research Institute of Medical Science (Bangkok, Thailand), and anti-DV nonstructural protein 1 (NS1), 2G6 generated in our laboratory was conjugated with allophycocyanin using Phycolink allophycocyanin conjugation kit (Prozyme) (5). Primary HUVEC were purchased from Cambrex and maintained in EGM-2 bulletkit medium, as recommended by the company. HLA-A*11 DV-restricted (NS3 aa 134–144) and HLA-A*2-restricted (E aa 211–219) CTL clones were generated by dengue serotype 2 peptides, as previously described (6, 7). CD14– PBMC, CD4+ T cells, and CD8+ T cells taken from healthy donors were isolated by negative or positive selection using MACS MicroBead (Miltenyi Biotec). Activated T cells were generated by treating cells with 100 ng/ml PMA and 500 ng/ml ionomycin for 18 h. Cells were then washed with medium before further use.

Virus stock

DV serotype 2, strain 16681, was propagated in C6/36 cells, and virus supernatant was collected and stored at –80°C. Virus stocks were free from endotoxin and mycoplasma as measured by Limulus amebocyte lysate assay (BioWhittaker) and PCR mycoplasma detection set (Takara Bio), respectively.

The titers of virus were determined by a focus-forming assay on PS clone D cells and expressed as focus-forming U/ml.

DC preparation and infection with DV

CD14+ cells were isolated from PBMC by positive selection using CD14 MicroBead mAbs (Miltenyi Biotec). To promote differentiation into immature DC, purified CD14+ cells were cultured for 5–6 days in RPMI 1640 supplemented with 10% FCS (R10), 20 ng/ml human rGM-CSF (First Link), and 25 ng/ml human rIL-4 (eBioscience) at 37°C in a 5% CO2 atmosphere. DC were infected with mock or DV at multiplicities of infection of 0.5 for 2 h. Cells were washed twice with RPMI 1640 before performing an experiment.

DC coculture

T cells or HUVEC were cocultured with mock-infected or DV-infected DC, and cell culture supernatants were collected for further analysis of cytokine and chemokine productions. To detect CD107a or cytokine/chemokine-producing cells, monensin (Golgistop; BD Pharmingen) and brefeldin A (BFA; Sigma-Aldrich) were added to the coculture, as previously described (8).

Cytotoxic activity of CTL to DV-infected DC was measured by standard 4-h chromium-released assay. Specific 51Cr release was calculated as follows: ((experimental release – spontaneous release)/(maximum release – spontaneous release)) x 100.

FACS analysis

Cell surface molecules, CD80, CD83, CD86, HLA-DR, CD8, CD38, and CXCR3, were detected with Abs at 4°C for 30 min. Thereafter, cells were fixed and permeabilized in FACS permeabilization buffer II (BD Pharmingen). Intracellular staining was achieved by staining with Abs to DV NS1 (2G6), IFN-{gamma}, TNF-{alpha}, and CXCL10. Cells were analyzed by a FACSCalibur.

Measurement of cytokine and chemokine concentrations

Cytokines were measured using a Bio-Plex human cytokine assay kit (Bio-Rad). In addition, levels of CXCL9, CXCL10, and CXCL11 were evaluated using Quantikine ELISA kits (R&D Systems) or a Beadlyte human cytokine kit (Upstate Biotechnology), according to the manufacturer’s instructions. Bio-Plex and Beadlyte analysis were performed using the Bio-Plex array reader (Bio-Rad), according to the manufacturer’s instruction.

Statistical analysis

The statistical significance of difference in the levels of cytokine between study groups was analyzed using Mann-Whitney U test. Value of p < 0.05 was considered to be statistically significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
DV-infected DC induce maturation of bystander DC and activate DV-specific T cells

DC are believed to be an important site of DV replication in vivo (9). One of the major receptors on DC for the DV envelope protein is DC-specific ICAM-grabbing nonintegrin (DC-SIGN) (10, 11). Immature DC can be readily infected, whereas mature DC are relatively refractory to infection (9). In contrast to bystander, noninfected DC, which up-regulate MHC and costimulatory molecules, DV-infected DC become stunned and fail to mature (12, 13). We were keen to investigate which factors contributed to bystander DC maturation and whether infected DC were capable of activating DV-specific T cells.

Monocyte-derived DC were infected with DV serotype 2, and the infection efficiency was determined 24 h later by intracellular staining for DV NS1. Infection efficiencies of between 30 and 60% were routinely obtained (Fig. 1a), and, as expected, the infected DC showed lower levels of HLA-DR, CD80, CD83, and CD86 expression than their noninfected counterparts (Fig. 1b). We wondered whether TNF-{alpha} and type I IFN produced by DV-infected DC were important in the maturation of bystander, noninfected DC (14). To test this, we cultured infected DC in the presence of neutralizing Ab to TNF-{alpha} and/or anti-IFN-{alpha}/βR2 blocking Ab (Fig. 1c). Up-regulation of CD80 expression on bystander cells was not significantly inhibited by the presence of these Abs. In contrast, CD86 and HLA-DR up-regulation were inhibited by the presence of either Ab, and the combination of TNF-{alpha} and anti-IFN-{alpha}/βR2 blocking Ab also almost completely blocked CD83 up-regulation on bystander cells. Taken together, these results imply that both TNF-{alpha} and type I IFN are key cytokines involved in the maturation of bystander-noninfected DC during DV infection.


Figure 1
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  		FIGURE 1. DV-infected DC induce T cell responses. A, DC were treated with mock or infected with DV for 24 h, and the infection was detected intracellularly by mAb specific to DV NS1 protein. B, Expression of cell surface maturation molecules on mock-treated DC and DV-infected DC, which were divided into two groups: DV-negative and DV-positive cells. Matched isotype controls are represented by the dashed line. These plots are representative of at least three independent experiments. C, DV-infected or mock-treated DC were cultured for 24 h in the absence or presence of anti-TNF-{alpha}, anti-IFN-{alpha}/βR2, or anti-TNF-{alpha} plus anti-IFN-{alpha}/βR2, and changes in surface maturation molecules were assessed. The results were expressed as mean fluorescence intensity (MFI) ± SE from three independent experiments (MFI was calculated by subtracting the MFI obtained with the corresponding isotype control). D, DV-specific T cells (CTL) were cocultured with HLA-matched DV-infected or mock-infected DC for 6 h in the presence of monensin and BFA. CTL responses were detected by IFN-{gamma} and TNF-{alpha} production and CD107a, a surrogate marker of degranulation/cytotoxicity (CTL:DC = 1:1). CTL were identified based on a lymphocyte gate and CD8+ population by flow cytometry. E and F, Results of a standard 4-h 51Cr release assay in coculturing CTL with 51Cr-labeled DV-infected or mock-treated HLA-matched DC (CTL:DC = 10:1).

 
Viral inhibition of up-regulation of costimulatory molecules on DC may be a mechanism of immune escape by impairing T cell activation, which may render Ag-specific T cells anergic. In primary DV infection, DV subversion of DC Ag presentation may serve to blunt the immune response, but one of the hallmarks of DV infection is the increase in disease severity in secondary infection. We were therefore keen to discover whether infected DC that failed to up-regulate costimulatory molecules could be recognized by memory T cells and whether such DC could be modulated by T cells, a situation pertinent to a secondary DV infection.

We added DV-specific CD8+ T cells to DV-infected DC. This was achieved by mixing infected, HLA-matched DC and a DV-restricted T cell clone. The CD8+ T cells in this assay were able to recognize the DV-infected DC, as evidenced by CD107a staining (a surrogate marker of degranulation), together with IFN-{gamma} and TNF-{alpha} production in the CD8-positive population (Fig. 1d). Furthermore, DV-infected cells were readily killed by specific T cells in a 51Cr release assay (Fig. 1e). The CTL clone used in these assays was generated using a DV serotype 2 peptide, but was able to recognize DC infected with the three other serotypes of DV to varying degrees (Fig. 1f). Together, these results strongly suggested that DV-specific memory T cells are capable of recognizing DV-infected DC, and in response produce proinflammatory cytokines and develop cytotoxic activity.

DV inhibition of DC maturation can be overcome by activated T cells

We next set out to discover how widespread the block to DC maturation was in infected DC. DV-infected DC were incubated with a variety of activating ligands for TLR, including LPS, poly(I:C), R848, and CL097, as well as L18 MDP (a NOD2 ligand), IFN-{gamma}, and TNF-{alpha}. In all cases, the responses of the DV-infected DC were blunted when compared with either the mock-infected or uninfected bystander cells (Fig. 2a). We next studied whether the presence of activated T cells, as in secondary DV infection, influences DC maturation. We cocultured DV-infected DC with resting or activated autologous CD4+ T cells, CD8+ T cells, or CD14– PBMC (Fig. 2b). Resting T cells had no effect on DC maturation, but activated T cells were able to restore DC maturation to levels comparable to those seen in mock-infected cells and almost to the same levels seen in the bystander DC. As expected, activated T cells caused maturation of mock-infected DC, as evidenced by up-regulation of CD86 expression. To investigate further whether the effect of activated T cells on DC maturation was mediated by soluble factors or direct cell contact, T cells were activated with PMA and ionomycin and then cocultured in a Transwell system in which the DV-infected DC were separated from activated T cells. We found activated CD4+ T cells, CD8+ T cells, and CD14– PBMC composed of both CD4+ and CD8+ T cells were able to restore maturation of DV-infected DC, suggesting that activated T cell-derived soluble factors were required for DC maturation (Fig. 2c). Taken together, these data show that maturation of DV-infected DC via both innate pattern recognition receptors and in response to individual proinflammatory cytokines is inhibited, but the block to maturation is incomplete and can be overcome by the presence of activated T cells.


Figure 2
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  		FIGURE 2. TLR ligands and activated T cells induce maturation of DV-infected DC. DV-infected or mock-treated DC were cultured for 24 h in the presence of TLR ligands or cytokines (A), or cocultured directly with resting or PMA/ionomycin-activated autologous CD4+ T cells, CD8+ T cells, or CD14– PBMC (B). Mock- or DV-infected DC were also cultured with T cells in a Transwell assay (C). Cells were stained with mAb against DC maturation markers and anti-DV NS1, followed by FACS analysis. The DC population was identified based on size and granularity. Expression of cell surface maturation molecules on mock-treated DC and DV-infected DC was defined into two groups: DV-negative and DV-positive cells. The results were expressed as MFI ± SE from three independent experiments.

 
DV-infected DC produce high levels of IFN-{gamma}-inducible chemokines

The IFN-{gamma}-inducible chemokines CXCL9 (monokine induced by IFN-{gamma}), CXCL10 (IFN-{gamma}-inducible protein-10), and CXCL11 (IFN-inducible T cell {alpha} chemoattractant) all interact with the same receptor (CXCR3) expressed on activated and memory T cells, and function as T cell chemoattractants (15, 16, 17, 18, 19, 20, 21). Because activated T cells appeared to have an important influence on maturation of DV-infected DC, we investigated whether infected DC produced T cell chemoattractants. CXCL9, 10, 11, and the proinflammatory cytokine TNF-{alpha} were quantified in supernatants from immature DC that were mock infected, infected with DV, or incubated with UV-inactivated DV (Fig. 3a). Live DV infection induced production of all four chemokines/cytokines, in particular CXCL10 and TNF-{alpha}.


Figure 3
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  		FIGURE 3. DV induce cytokine/chemokine production by DC. A, DC were treated with mock, or infected with UV-inactivated DV, or DV. B, DC were treated with mock or infected with DV in the presence or absence of CTL. Culture supernatants were collected at 48 h and measured for cytokine/chemokine levels. The results are shown as mean ± SE from three independent experiments. C, CTL were cocultured with either mock-treated or DV-infected DC for 6 h in the presence of monensin and BFA. IFN-{gamma}, TNF-{alpha}, and CXCL10 production were then detected by flow cytometry. CTL and DC populations were identified based on size, granularity, and CD8 expression. These plots are representative of three independent experiments.

 
To determine the effect of memory T cells in this scenario, we again mixed infected, HLA-matched DC and a DV-restricted T cell clone. The presence of T cells greatly increased production of CXCL9 and CXCL10, with less effect on CXCL11 and TNF-{alpha} (Fig. 3b). To determine whether DC or T cells were the source of chemokine/cytokine production, we performed intracellular staining for CXCL10 and TNF-{alpha}. As expected, CXCL10 was produced exclusively by DC, whereas TNF-{alpha} was produced by both DC and T cells (Fig. 3c).

DV infection resets the sensitivity of DC to IFN-{gamma} stimulation

CXCL9, 10, and 11 production are induced by IFN-{gamma} (15, 22, 23). Coculture of DC and DV-specific T cells resulted in IFN-{gamma} production by T cells (Fig. 3c). We therefore reasoned that the large increase in chemokine production seen with the coculture of infected DC and DV-specific T cells may be mediated by IFN-{gamma}, which reached levels of 3.4 ± 1.2 ng/ml in the coculture experiemnts. To test this, we first stimulated DV-infected and mock-infected DC with 1–100 ng/ml (10–1000 U/ml) IFN-{gamma} and measured production of CXCL-9, 10, and 11 (Fig. 4). Production of IFN-{gamma}-inducible chemokines was considerably greater in DV-infected DC compared with mock-infected cells, implying enhanced responses to IFN-{gamma} stimulation. Similar results were obtained when we cultured DV-infected or mock-infected DC in the presence or absence of culture supernatants from PMA/ionomycin-activated CD14– PBMC (data not shown). Furthermore, we found a similar response in K562 (human leukemic) cells stably expressing DV replicons when compared with parental K562 cells, showing that the presence of DV RNA replication and expression of DV nonstructural proteins reset the sensitivity of cells to IFN-{gamma}-mediated stimulation of chemokine production (data not shown).


Figure 4
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  		FIGURE 4. DV infection resets the sensitivity of DC to IFN-{gamma} stimulation. DC were treated with mock or infected with DV for 12 h. Increasing amounts of IFN-{gamma} were then added, and the culture was incubated for an additional 24 h. Culture supernatants were harvested and measured for CXCL9, CXCL10, and CXCL11 levels by ELISA. The results are shown as mean ± SE from three independent experiments.

 
DV-infected DC enhance endothelial cell production of CXCL10

The hallmark of DHF is increased vascular permeability. Immunohistochemistry of infected tissues has demonstrated monocytes and lymphocytes in close proximity to the vascular endothelium (24, 25). To investigate the complex interplay between cell types implicated in the pathogenesis of DHF, we cocultured human vascular endothelial cells (HUVEC) in the presence or absence of DC or PBMC, and measured CXCL10 production by ELISA (Fig. 5a). HUVEC, in common with resting or activated PBMC, produced little CXCL10, but coculture of HUVEC with activated PBMC resulted in large amounts of CXCL10 production (Fig. 5a). A similar result was found when HUVEC were cocultured with DV-infected DC. To determine which cells were producing CXCL10, we performed intracellular staining. As before, DV-infected DC produced CXCL10, but in the coculture experiments CXCL10 production was greater in HUVEC than DC (Fig. 5b). To exclude the possibility that HUVEC production of CXCL10 was a consequence of infection, we stained the cells for DV and showed that only the DC were infected (Fig. 5b). These data show that HUVEC produce CXCL10 when cocultured with DV-infected DC, and this is not attributable to direct infection of HUVEC by DV.


Figure 5
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  		FIGURE 5. DV-infected DC and activated T cells enhance CXCL10 production by endothelial cells. A, HUVEC were cocultured with mock-treated DC, DV-infected DC, resting, or PMA/ionomycin-activated CD14– PMBC (HUVEC:DC or HUVEC:PBMC = 1:5 or 1:10, respectively). Cell supernatants were collected after 24 h and assayed for CXCL10 production by ELISA. The results are shown as mean ± SE from three independent experiments. B, HLA-A*2-negative HUVEC were cocultured with mock-treated or DV-infected HLA-A*2-positive DC for 24 h in the presence of BFA. Anti-HLA-A*2 was used to identify DC and HUVEC populations. CXCL10 production and DV infection were then detected on each cell type. These are representative of three independent experiments. C, HUVEC were cultured in the presence of the indicated concentrations of IFN-{gamma}, IFN-{alpha}, and/or TNF-{alpha}. Culture supernatants were harvested, and CXCL10 levels were measured. The results are shown as mean ± SE from three independent experiments. D, HUVEC were cocultured with DV-infected DC in the presence or absence of anti-TNF-{alpha}, anti-IFN-{alpha}/βR2, or anti-TNF-{alpha} plus anti-IFN-{alpha}/βR2. Cell supernatants were collected at 24 h and assayed for CXCL10 production by ELISA.

 
We hypothesized that CXCL10 production in cocultures of HUVEC with either DV- infected DC or activated PBMC is mediated by cytokines. To determine whether HUVEC are capable of producing CXCL10 in response to cytokines, we treated HUVEC with IFN-{gamma}, TNF-{alpha}, or IFN-{alpha} (Fig. 5c). We found that IFN-{gamma} induced CXCL10 production in HUVEC in a dose-responsive manner. The combination of IFN-{gamma} and TNF-{alpha} was strongly synergistic (Fig. 5c). IFN-{alpha} alone induced a low level of CXCL10 production by HUVEC, but this was enhanced by addition of TNF-{alpha}. When DV-infected DC were cocultured with HUVEC in the presence of anti-TNF-{alpha} and/or anti-IFN-{alpha}/βR2 blocking Abs, production of CXCL10 was diminished (Fig. 5d). This suggests that TNF-{alpha} and type I IFN are important in the interaction between DC and HUVEC that results in CXCL10 production.

IFN {gamma}-induced chemokines in patients with DHF

Our in vitro experiments that studied the interaction between DV-infected DC and other cells relevant to human DV infection suggest that CXCL10 production may be up-regulated in both infected DC and endothelial cells through cytokine-mediated pathways. The presence of activated T cells may greatly enhance CXCL10 production. To test the biological significance of these observations, we first measured IFN-{gamma}-inducible chemokines in 50 children with DV (14 DF, 36 DHF). CXCL10 was significantly raised in DHF vs DF at days –3, –2, and –1 (in which day 0 is defined as the day of defervescence; p = 0.0176, 0.0003, and 0.0075, respectively), with peak levels in plasma up to 254 ng/ml (Fig. 6a). The levels of CXCL10 peaked at day –1, and from day 0 onward there was no significant difference between patients with DF and DHF. CXCL9 and CXCL11 were also significantly higher in the DHF vs DF groups at day –2 (p = 0.005 and 0.0154, respectively). We also measured levels of MCP-1, another IFN-{gamma}-inducible chemokine from a different family, and found no significant difference between the DF and DHF groups (Fig. 6a). Five further blood samples were available from children who died from DHF. CXCL9 levels were significantly higher in these samples than the peak plasma levels in survivors from DHF (Fig. 6b).


Figure 6
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  		FIGURE 6. IFN-{gamma}-inducible chemokines in DV-infected patients. A, Levels of CXCL9, CXCL10, CXCL11, and MCP-1 in DF and DHF. B, Levels of CXCL9 in plasma taken from DHF cases who survived or died. Horizontal lines indicated mean values. Statistical values were determined by Mann-Whitney U test (*, p < 0.05; **, p < 0.01; ***, p < 0.001).

 
Finally, we went on to look at CXCR3, the receptor for CXCL9, 10, and 11 in T cells in a patient with DHF across the course of infection. We detected a large number of T cells that express CXCR3 around the time of defervescence, constituting up to 69% of all T cells. Of these, 81% were activated, as evidenced by CD38 expression. At convalescence day 60, 32% of all T cells were CXCR3 positive, and only 10% of these were CD38 positive (Fig. 7).


Figure 7
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  		FIGURE 7. CXCR3 expressions on CD8+ T cells from DV-infected patients. PBMC samples were taken from DV-infected patients during the course of acute infection and at 2 mo follow-up. CD8+ T cells were gated, and surface expression of CXCR3 and CD38 was determined. The levels of CXCL10 in plasma are also shown. These are representative of three patients.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
DC are targets of infection by DV and a number of unrelated viruses, including HIV, members of the herpes virus family HSV and CMV, and Lassa and Ebola viruses (26, 27, 28, 29, 30). It has been shown that immature DC can be readily infected by DV through DC-SIGN, whereas mature DC, expressing low DC-SIGN, can be infected by DV via an Ab-dependent enhancement process (10, 11, 31). Because DC are central to the development of the acquired T cell response, pathogens have evolved mechanisms to reduce the capacity of infected DC to prime the adaptive immune response. One mechanism to achieve this is to maintain infected DC in an immature state with relatively low level Ag presentation and costimulatory molecule expression. DV infection inhibits DC maturation, so that infected DC are potentially less able to stimulate T cell responses, and, in the absence of costimulation, may even induce anergy in a subset of DV responsive T cells (12, 13). This effect on DC maturation can also be observed with other viruses, including HSV, vaccinia virus, CMV, hepatitis C virus, and HIV (32, 33, 34, 35, 36). The mechanism whereby DV prevents DC maturation has not been fully elucidated, but some recent observations are likely to be important. First, DV infection induces production of IFN-{alpha}/β, which we have shown in this study contributes to bystander DC maturation. In contrast, the effect of IFN-{alpha}/β in infected cells is specifically inhibited by DV-mediated inhibition of type I IFN signal transduction (37, 38). Second, some recent observations with HIV may have relevance to DV infection. Both HIV and DV use the C-type lectin DC-SIGN to gain entry into DC (10, 11, 39). Cross-linking of DC-SIGN by HIV or by mAb sets in train a signaling cascade that maintains DC in an immature state refractory to stimulation with maturation signals such as LPS (40).

Failure of DV-infected DC to mature fully implies that T cell responses to DV infection may be inefficiently raised by direct priming and may instead rely on cross-priming. We and others have shown that, in contrast to infected DC, bystander DC mature in response to DV infection. Our data show that TNF-{alpha} and type I IFN play important roles in maturation of bystander DC, and hence, may play a crucial role in cross-priming of T cell responses to DV infection in primary infections. Importantly, we have shown in this study that the block to maturation in DV-infected DC is incomplete and can be overcome by activated T cells via the production of soluble factors such as type 1 IFN and TNF-{alpha}.

In secondary infection, cross-reactive memory T cell populations will already be present. Interaction of these cells with immature DV-infected DC will now lead to a different outcome. Memory cells have a lower threshold for activation and can respond and proliferate in the absence of costimulation. Soluble mediators such as TNF-{alpha} and IFN-{gamma} together with stimulation via CD40L can mature the DV-infected DC and can also change their secretion of proinflammatory cytokines and chemokines such as CXCL10 (14).

CXCL9, CXCL10, and CXCL11 all interact with CXCR3 found on memory and activated T cells, especially IFN-{gamma}-producing T cells, and promote their chemotaxis (17, 18, 20, 21, 41, 42). CXCR3 expression on dengue-infected T cells is high during acute infection and reduces upon convalescence. One of the interesting features of DV and several other acute severe viral infections is the relative lymphopenia often seen during the acute viremic phase (43). We speculate that part of this may be due to margination of CXCR3-expressing circulating lymphocytes to sites of DV infection, such as infected monocytes/DC in the tissues driven through the production of CXCR3-reactive chemokines, such as CXCL10. As levels of these chemoattractants fall, activated T cells expressing CXCR3 can then spill out into the general circulation.

We show in this study that the presence of activated T cells greatly increases production of CXCL10 and other IFN-{gamma}-inducible chemokines, CXCL9 and CXCL11, from DV-infected DC. This is probably a consequence of enhanced sensitivity of DV-infected cells to IFN-{gamma}-mediated stimulation of chemokine production. The mechanism for the increased effect of IFN-{gamma} on DV-infected DC may in part be due to an enhanced ability of DV-infected cells to transduce signals via the type II IFN receptor. It has been shown that DV infection leads to degradation of STAT-2, leading to a block in type I IFN receptor signaling that requires STAT-1/STAT-2 heterodimers to transduce the signal. STAT-1 is, however, not degraded, and, in fact, the levels of STAT-1 and STAT-1 phosphorylation seem to be increased in cells transduced with a DV replicon and DV-infected DC (37, 38).

The response seen in secondary infection will thus be further amplified by DV-specific T cells interacting with DC, leading to increased CXCL9, 10, and 11 production, allowing more T cells to be drawn to the site of infection. In addition, we have shown in this study that the production of CXCL10 is greatly amplified when DV-infected DC are cocultured in the presence of endothelial cells, potentially further promoting T cell recruitment to perivascular areas and contributing to the immunopathology of vascular leak in DHF. Finally, it has been shown previously that IFN-{gamma} can increase Ab-dependent enhancement of infection of Fc-bearing cells perhaps by increasing FcR expression (44). We have observed a similar phenomenon using activated T cells, which can enhance viral replication in the presence of enhancing Ab by over 100% (data not shown). Thus, in secondary infection, T cells may drive increased Ag production as well as Ag presentation by DC, further stoking a bonfire of T cell activation and cytokine production.

It recently has been shown that the levels of CXCL10 and 11 are elevated in cases of DF (45). In the present study, we set out to determine whether the levels of CXCL9, CXCL10, and CXCL11 were predictors of disease severity. Significant differences in chemokine levels were apparent 3 days before defervescence, and, hence, preceded the onset of severe clinical disease. From our data, we were able to calculate the value of chemokine levels in predicting subsequent clinical disease severity. For this analysis, we redefined the day of illness as days after onset of fever, because the day of defervescence would not be known at the time of sampling; we also performed an analysis based on chemokine levels on the day of admission to hospital. Using a cutoff of 50,000 pg/ml, CXCL10 gave sensitivities/specificities for predicting subsequent DHF, as follows: fever day 3, 100/66; day 4, 82/63; day 5, 60/70; day 6, 61/51; and on the day of admission, 76/50. CXCL9 was not measured at all time points, but a cutoff of 200 pg/ml at the day of admission gave values of sensitivity/specificity for predicting DHF of 77/75. Using both parameters together (either CXCL9 >200 or CXCL10 >50,000, or both) gave values of 90/50 on the admission day.

The role of CXCR3 in immune responses to infectious pathogens is complex; in some models blockade leads to lethal uncontrolled infections, whereas in other models CXCR3 has been associated with immunopathology. In the former group, neutralization of CXCL10 with mAb or mice deficient in CXCR3 led to increased pathogen replication and mortality in murine infection with Toxoplasma gondii and primary DV infection, although the latter is a poor model of the human disease (46, 47). In other models, lack of CXCR3 signaling is associated with a decrease in immunopathology; these include lymphocytic choriomeningitis virus infection in which wild-type mice die from a T cell-mediated immunopathology (48). CXCL9 and CXCL10 are expressed in demyelinating lesions in multiple sclerosis, in which they are proposed to promote the ingress of T cells, and there is now much interest in the use of anti-CXCL10 mAb to prevent disease (49, 50).

In our studies, levels of CXCL9, CXCL10, and CXCL11 were associated with disease severity, and because DHF most likely represents a T cell-driven immunopathology, blockade of CXCR3 may modulate disease severity in secondary infections. However, in the absence of good animal models of severe secondary DV infection, this will be difficult to exemplify.

In summary, the pathogenesis of DV infection is multifactorial, involving the complex interplay of a number of viral and host factors. In secondary infection, the combination of increased viral replication induced by both Ab enhancement and T cells can lead to a situation of high viral burden. It has been demonstrated that there is a correlation between the peak viral load and disease severity (51, 52). This is then combined with high amplitude T cell response, and failure to mount a suppressive T regulatory response, massive T cell activation, and secretion of proinflammatory cytokines/chemokines and mediators will occur in concert with viral clearance (53). Many cytokines, such as TNF-{alpha}, IFN-{gamma}, IL-2, IL-6, IL-8, IL-10, MCP-1, and MIP-1β, have been reported to be significantly elevated in patients with DHF in comparison with DF or healthy controls (54, 55, 56, 57, 58, 59, 60, 61, 62) (our unpublished data). Some of these cytokines have been further demonstrated to have an effect on endothelial cells, leading to increased vascular permeability. In the present study, we found a complex interplay between DV, DC, T cells, and endothelial cells, whereby in combination there is a dramatic up-regulation of CXCR3-reactive chemokine production. These chemokines will draw more activated/memory T and NK cells and other CXCR3-expressing cells to the site of infection, leading to massive immune responses and cytokine production, as mentioned above, which result in disease severity.


   Acknowledgments
 
We thank S. Noisakran, P. Chotiyarnwong, A. Chairunsri, staff at KhonKhaen Hospital in Thailand, and V. Chan for technical assistances and sample collection. Special thanks to N. Tangthawornchaikul for her roles in data, specimen, and project management to enable us to make sense out of scientific data.


   Disclosures
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by the Medical Research Council, U.K.; the Wellcome Trust, U.K.; the National Institute for Health Research Biomedical Research Centre funding scheme; the Thailand Tropical Disease Research Program T2; and the Thailand National Centre for Genetic Engineering and Biotechnology. Back

2 Address correspondence and reprint requests to Dr. Gavin Screaton, Department of Immunology, Division of Medicine, Hammersmith Hospital, Imperial College, London W12 0NN, U.K.; E-mail address: g.screaton{at}imperial.ac.uk or Dr. Juthathip Mongkolsapaya, Department of Immunology, Division of Medicine, Hammersmith Hospital, Imperial College, London W12 0NN, United Kingdom; E-mail address: j.mongkolsapaya{at}imperial.ac.uk Back

3 Abbreviations used in this paper: DV, dengue virus; BFA, brefeldin A; DC, dendritic cell; DC-SIGN, DC-specific ICAM-grabbing nonintegrin; DF, dengue fever; DHF, dengue hemorrhagic fever; MFI, mean fluorescence intensity; NS1, nonstructural protein 1. Back

Received for publication February 20, 2008. Accepted for publication August 19, 2008.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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