The Absence of IDO Upregulates Type I IFN Production, Resulting in Suppression of Viral Replication in the Retrovirus-Infected Mouse

Masato Hoshi, Kuniaki Saito, Akira Hara, Ayako Taguchi, Hirofumi Ohtaki, Ryo Tanaka, Hidetsugu Fujigaki, Yosuke Osawa, Masao Takemura, Hidetoshi Matsunami, Hiroyasu Ito and Mitsuru Seishima
J Immunol September 15, 2010, 185 (6) 3305-3312; DOI: https://doi.org/10.4049/jimmunol.0901150

Abstract

Indoleamine 2,3-dioxygenase, the l-tryptophan–degrading enzyme, plays a key role in the powerful immunomodulatory effects on several different types of cells. Because modulation of IDO activities after viral infection may have great impact on disease progression, we investigated the role of IDO following infection with LP-BM5 murine leukemia virus. We found suppressed BM5 provirus copies and increased type I IFNs in the spleen from IDO knockout (IDO−/−) and 1-methyl-d-l-tryptophan–treated mice compared with those from wild-type (WT) mice. Additionally, the number of plasmacytoid dendritic cells in IDO−/− mice was higher in the former than in the WT mice. In addition, neutralization of type I IFNs in IDO−/− mice resulted in an increase in LP-BM5 viral replication. Moreover, the survival rate of IDO−/− mice or 1-methyl-d-l-tryptophan–treated mice infected with LP-BM5 alone or with both Toxoplasma gondii and LP-BM5 was clearly greater than the survival rate of WT mice. To our knowledge, the present study is the first report to observe suppressed virus replication with upregulated type I IFN in IDO−/− mice, suggesting that modulation of the IDO pathway may be an effective strategy for treatment of virus infection.

Indoleamine 2,3-dioxygenase has been identified as an enzyme endowed with powerful immunomodulatory effects, resulting from its enzymatic activity that leads to catabolism of the essential amino acid l-tryptophan (L-TRP) (1, 2). This enzyme is expressed in epithelial, macrophage, and dendritic cells (DCs) induced by proinflammatory cytokines, including type I and type II IFN (3). In addition, engagement of CTLA-4 with CD80/CD86 on the membrane of DCs stimulates IDO transcriptional expression and activity (4, 5). Further, IDO is not a mechanism of regulatory T cell (Treg) function, but is rather induced by Tregs. Increased IDO activity provokes tolerogenicity of APCs and deprives T cells of tryptophan, leading to proliferation arrest and T cell apoptosis (6). Further, l-tryptophan-kynurenine pathway metabolites have been shown to act as immunoregulatory molecules that mediate immunosuppressive effects in the tissue microenvironment (7–9). Although IDO activities in various tissues are induced by several cytokines after viral infections, the role of IDO in vivo after viral infections is not fully understood. The role of IDO in chronic viral infectious diseases was investigated in mice infected with LP-BM5 murine leukemia virus (MuLV), including both replication-competent and replication-defective viruses, which resulted in the development of a fatal immunodeficiency syndrome in mice known as murine AIDS (10). Murine AIDS is characterized by activation and proliferation of T and B cells, impaired T and B cell function, an aberrant regulation of cytokine production, hypergammaglobulinemia, decreased NK cell function, the development of B cell lymphoma, and susceptibility to opportunistic infections (11). In the current study, we used IDO−/− mice to examine whether IDO is an important factor for immune regulation against LP-BM5 infection and especially whether the presence of IDO is necessary for the induction of cytokines and IDO-related molecules, which are important for viral clearance. Remarkably, we demonstrated upregulated type I IFNs and downregulated virus replication in IDO−/− mice with LP-BM5 infection. To our knowledge, these findings provide the first evidence that the absence of IDO is involved in the clearance of murine retroviral infection via upregulated type I IFNs, and they offer insights into the role of innate and adaptive immune-response cytokines in control of viral dissemination from the site of entry into the host.

Materials and Methods

Mice

Four- to 6-wk-old male mice were used in this study. IDO1 gene-deficient (IDO−/−) mice of a C57BL/6J background were obtained from The Jackson Laboratory (Bar Harbor, ME). Mice that were wild-type (WT) (+/+) or homozygous null (−/−) for targeted disruption of the IDO gene were selected from offspring of heterozygous/homozygous matings by PCR tail DNA. The mice were divided into four groups: infected mice with or without 1-methyl-d-l-tryptophan (1-MT) and uninfected mice with or without 1-MT. The mice were administrated 1-MT in drinking water (5 mg/ml) postinfection, and they ingested an average of 3.5 ml/d. The experiments were performed according to the Guidelines for Animal Care of Gifu University (Gifu City, Japan).

LP-BM5 infection

Mice were i.p. infected with 0.2 ml MuLV mixture prepared from the G6 clone of chronically infected SC-1 cells obtained through the National Institutes of Health AIDS Research & Reference Reagent Program (Germantown, MD), as described previously (12). Control groups of mice were infected with 0.2 ml culture media from uninfected SC-1 cells.

Toxoplasma gondii infection

Parasites were harvested from the brains of mice chronically infected with T. gondii cysts (“Fukaya” strain) as described in a previous report (13). Brain tissue was dispersed in saline. The final concentration of the infectious agent was adjusted to a dose of 20 cysts/0.2 ml, which was injected i.p. into the mice.

Measurement of l-tryptophan and l-kynurenine

L-TRP and l-kynurenine (L-KYN) were measured by using HPLC with a spectrophotometric detector (Tosoh ultraviolet-8000, Tosoh, Tokyo, Japan) or fluorescence spectrometric detector (Hitachi, Tokyo, Japan) as described in a previous report (14).

Enzymatic assay for IDO activity

IDO activity was assayed as described in a previous report (13). In brief, tissues were homogenized with a Polytron homogenizer (Kinematica, Lucerne, Switzerland) in 1.5 volumes of ice-cold 0.14 M KCl-20 mM potassium phosphate buffer (pH 7). The homogenate samples were centrifuged at 7000 × g and 4°C for 10 min. An aliquot of supernatant was taken for the measurement of IDO activity. The reaction mixture contained 50 μl enzyme preparation and 50 μl substrate solution. The composition of the substrate solution was 100 mM potassium phosphate buffer (pH 6.5), 50 μM methylene blue, 20 μg catalase, 50 mM ascorbate, and 0.4 mM L-TRP. Postincubation of the reaction mixture at 37°C, samples were acidified with 3% perchloric acid and centrifuged at 7000 × g and 4°C for 10 min. The concentrations of the enzymatic products were measured by using HPLC. Enzyme activity was expressed as the product content per hour per gram of tissue protein.

Histopathology

Tissues were fixed in 10% formalin in PBS overnight. Specimens were then embedded in paraffin. Sections (thickness: 4 μm) were used for H&E staining and immunofluorescence as described in a previous report (15). For the immunofluorescence analysis, the deparaffinized sections were heated in 0.1 M citrate buffer (pH 6) using the Pascal heat-induced target retrieval system (DakoCytomation, Carpinteria, CA). The sections were subsequently incubated with rabbit anti-IDO polyclonal Ab (anti-mouse IDO Ab was generated by the peptide H-CMKPSKKKPTDGDKS-OH) and biotin anti-CD11c Ab (1:400) in 2% BSA/PBS and incubated overnight at 4°C. After primary Ab incubation, the sections were rinsed with PBS and incubated with secondary Abs for 30 min at room temperature to visualize the cell signals. The secondary Abs used were FITC-conjugated streptavidin (1:200; DakoCytomation), rhodamine-conjugated anti-rabbit IgG (1:300; DakoCytomation), and 4,6-diamino-2-phenylindole for nuclear staining (Dojindo, Tokyo, Japan). Immunostained slides were visualized using the fluorescence microscope BX51 equipped with a DP70 digital camera (Olympus, Tokyo, Japan).

RNA extraction, RT-PCR, and real-time PCR

Total RNA was rapidly isolated using the Isogen RNA isolation kit (Nippon Gene, Tokyo, Japan). Total RNA (1 μg) was used for the synthesis of the first strand of cDNA. RT-PCR was performed using the Takara mRNA selective PCR kit (Takara Biomedicals, Tokyo, Japan). cDNA quantification for IDO, β-actin, IL-4, IL-10, TGF-β, 18s, IFN-α, -β, and Foxp3 was performed using real-time PCR conducted with LightCycler (Roche, Meylan, France). IDO, IFN-α, and β-actin reactions were performed using a SYBR green PCR mix (Qiagen, Hilden, Germany) as described in a previous report (13). IL-4, IL-10, TGF-β, 18s, IFN-β, and Foxp3 reactions were performed using a TaqMan probe. The TaqMan Universal Master Mix (Applied Biosystems, Foster City, CA) was used, and PCR cycling conditions were 95°C for 10 min and 50 cycles of 95°C for 15 s and 60°C for 1 min. The reference cDNA sample used to make standard curves was obtained from the spleen at 8 wk after LP-BM5 infection. Data analysis was performed with the LightCycler software (version 5.32; Roche).

Viral load measurement

We quantified BM5d RNA in the spleen with real-time PCR as described in a previous report (16). In brief, RNA was extracted from the spleen and amplified using BM5d-specific primers. The amount of viral RNA in each sample was quantified against a standard curve constructed using RNA transcripts of LP-BM5def. Quantification was made using the LightCycler software (version 5.32; Roche). The copy number was expressed per 1 μg RNA.

Splenocyte subpopulation preparations, culture condition, and proliferation

Splenocyte suspensions were labeled with various Ab-coupled paramagnetic beads (MACS; Miltenyi Biotec, Auburn, CA) and subjected to column purification according to the manufacturer’s protocol. In brief, the spleen cell suspensions were labeled with anti-CD4 and CD11c beads, and positive selection yielded cell preparations that were ≥98%, as detected by flow cytometric analysis. Isolated CD11c-positive cells were plated in 96-well culture plates (5 × 105 cells/ml; 200 μl/well) in RPMI 1640 (Nikken Bio Medical Laboratory, Kyoto, Japan) supplemented with 10% heat-inactivated FBS (Boehringer Mannheim Biochemica, Mannheim, Germany), 100 U/ml penicillin, and 100 mg/ml streptomycin (Invitrogen, Carlsbad, CA) at 37°C in 5% CO2. At 72 h of the culture, an aliquot of culture media was used for the ELISA analysis. The splenocytes (2 × 105 cells/ml; 200 μl/well) from individual normal and LP-BM5–infected mice (WT, 1-MT–treated, and IDO−/−) were plated in triplicate onto 96-well plates with media to obtain a final concentration of 1 μg/ml Con A. After 48 h, the cells were terminally pulsed with1 μCi [3H]thymidine (GE Healthcare, Tokyo, Japan) and harvested after 6 h for assessment of thymidine incorporation by scintillation counting (Beckman Coulter LS-6500, Beckman Coulter).

Flow cytometric analyses

The splenocytes were washed twice in PBS. Membrane Ag expression was determined by incubating cells for 20 min at 4°C with anti-mouse CD4, CD8, CD3, DX-5, CD11c, B220, and murine plasmacytoid DC (pDC) Ag-1 mAb (BD Biosciences, Franklin Lakes, NJ). Cells were analyzed using FACScan (BD Biosciences). Samples were gated on viable cells by forward and side light scatter, and a minimum of 10,000 live cell events was acquired for each sample.

Western blot analysis

Total spleens were lysed by the addition of ice-cold lysis buffer containing 0.5% Triton X-100, 50 mM Tris (pH 8), 150 mM NaCl, 5 mM EDTA, 5 mM NaF, 1 mM Na3VO4, 10 μg/ml aprotinin, pepstatin A, and 5 μg/ml leupeptin. Lysates were placed on ice for 20 min and then centrifuged for 30 min at 12,000 × g in a microcentrifuge at 4°C to remove the nuclei. Proteins (20 μg) were separated on 10% SDS-polyacrylamide gels and transferred to polyvinylidene difluoride membranes. The membranes were blocked with 5% nonfat dried milk in T-PBS (0.2% Tween 20 in PBS) and incubated with Ab against IDO or GAPDH for 1 h. The membranes were subsequently washed and incubated for 1 h with secondary Ab conjugated to HRP. Immunolabeling was performed using an ECL detection system (Millipore, Billerica, MA).

Neutralization and quantification of type I IFNs in ex vivo cultures

IFN-α and IFN-β in supernatants from the isolated CD11c-positive cell cultures were analyzed using an ELISA kit (R&D Systems). To assess the effect of neutralization of type I IFNs, the splenocytes obtained from LP-BM5–infected WT mice or IDO−/− mice were cultured for 72 h in a medium containing 5 × 106 cells/ml at 37°C in the presence or absence of rabbit anti-mouse IFN-α Ab (1000 neutralization unit/ml; Merck, Darmstadt, Germany), rabbit anti-mouse IFN-β Ab (1000 neutralization unit/ml; PBL Biochemical Laboratories, Piscataway, NJ), or isotype control mAb (Santa Cruz Biotechnology, Santa Cruz, CA). In addition, LP-BM5 replication was measured in the collected supernatant and splenocytes by using real-time PCR.

Statistical analyses

Results are expressed as mean ± SDs. Survival rates of mice were analyzed by the Kaplan-Meier method. Comparisons between two groups were made using the Student t test, and those among three groups were made with a one-way ANOVA. Stat View 4.5 software was used for statistical analyses on a Windows XP operating system (Microsoft, Redmond, WA). A p value of <0.05 was considered statistically significant.

Results

Changes in serum L-TRP catabolism and increased IDO following LP-BM5 infection

To assess the changes in IDO activity following LP-BM5 infection, we first addressed the time course of L-TRP catabolism in blood at 1, 2, 4, and 8 wk postinfection (wpi). As shown in Fig. 1A, LP-BM5 virus infection resulted in a marked increase in serum L-KYN concentration, the ratio of L-KYN/L-TRP, and a decrease in serum L-TRP concentration at 8 wpi. We next examined both IDO mRNA and enzymatic activities in the lung, liver, brain, and spleen. The level of IDO mRNA, protein, and enzymatic activities in the spleen was maximally upregulated in infected mice at 8 wpi (Fig. 1B–D). In the lung, IDO mRNA was also significantly upregulated at 4 wpi, as was IDO enzymatic activity (uninfected 6.8 ± 0.9 nmol/g/h versus infected 29.4 ± 5.9 nmol/g/h, mean ± SD). Although the levels of IDO mRNA and enzymatic activities in the liver and brain did not increase to significant levels until 8 wpi, they were significantly enhanced compared with uninfected mice at 12 wpi (data not shown). Histopathologic examination of the spleens of LP-BM5–infected mice at 8 wpi by H&E staining showed loss of lymphoid follicle structure (Fig. 1E). This structural change in the spleen indicates the preneoplastic condition of lymphoid tissues that were affected by LP-BM5 retrovirus infection. Immunofluorescent staining of CD11c (green) and IDO (red) indicates coexpression of CD11c and IDO (yellow) in the DCs in the spleens of the LP-BM5–infected mice (merge image, Fig. 1F).

FIGURE 1.
FIGURE 1.

Time course of IDO expression and serum L-TRP catabolism in LP-BM5–infected mice. WT mice were inoculated with LP-BM5 virus; spleen and serum were taken at the indicated weeks. A, The levels of serum l-tryptophan and l-kynurenine and the ratio of L-KYN/L-TRP were analyzed using HPLC (n = 6). Uninfected mice, opened bars; infected mice, closed bars. B, The upregulation of IDO mRNA in spleen from uninfected (opened squares) or infected (closed squares) mice was quantified using RT- qPCR (n = 6). The levels of target mRNA were normalized to β-actin mRNA as an endogenous control. C, The IDO enzymatic activity was dramatically enhanced time dependently (n = 6). Uninfected mice, opened bars; infected mice, closed bars. Results are presented as the mean ± SD. D, The protein levels of IDO were analyzed by Western blotting. IDO protein was increased time dependently. E, H&E staining of the normal spleen (week 0) and LP-BM5-infected spleen (week 8) (original low magnification ×10; 120 mM; original high magnification ×40; 30 mM). F, Double immunofluorescence labeling was performed with biotin anti-mouse CD11c Ab (green), rabbit anti-mouse IDO Ab (red), and DAPI nuclei staining; the merged image shows staining with both Abs and DAPI. The section shows the image of the spleen 8 wk after LP-BM5 infection (original high magnification ×40; 30 mM). E and F, Scale bars, 120 μm (low magnification ×10; 120 mM); scale bar, 30 μm (high magnification). The results of one of the five analyses with similar results are shown. *p < 0.05; **p < 0.001, as determined using one-way ANOVA. qPCR, quantitative PCR.

Effect of IDO gene deficiency on IFNs and various immunoregulatory factors after LP-BM5 infection

We investigated the time courses of the gated percentage of CD4+CD25+ cells (Fig. 2A) and the expression of Foxp3 mRNA (Fig. 2B) as a specific marker of Tregs in the whole spleen from LP-BM5–infected WT and IDO−/− mice. Expression of Foxp3 mRNA and the number of CD4+CD25+ T cells in WT and IDO−/− mice at 2 wpi were significantly upregulated compared with those of uninfected mice. It is noteworthy that there was no change in the levels of IL-4, IL-10, and TGF-β mRNA in the spleens of IDO−/− mice at 8 wpi compared with those of WT mice (Fig. 2C–E). Surprisingly, the expressions of both IFN-α and IFN-β mRNA in the spleens of IDO−/− mice at 8 wpi were dramatically upregulated compared with those of WT mice (Fig. 2F, 2G). Next, we isolated CD11c+ and CD4+ cells from the spleens of the mice at 8 wpi to identify type I IFN-generated cells. The level of IFN-α in CD11c+ cells from IDO−/− mice tended to be higher than that in CD11c+ cells from WT mice (Fig. 2H). Moreover, the level of IFN-β in CD11c+ cells from IDO−/− mice was significantly upregulated compared with that in CD11c+ cells from WT mice (Fig. 2I). To confirm the benefit of IDO deficiency without the potential confounding effects of long-term compensation in the knockout mouse system, we examined type I IFN production in the culture supernatant of CD11c+ cells isolated from the spleens of 1-MT–treated mice after LP-BM5 infection (Fig. 3). Consistent with the results obtained for IDO−/− mice, the level of type I IFN in 1-MT–treated mice at 8 wpi was clearly upregulated compared with that in WT mice.

FIGURE 2.
FIGURE 2.

Type I IFNs and immunosuppressive cytokine production in the spleens from WT and IDO−/− mice after LP-BM5 infection. Spleens from LP-BM5–infected WT (n = 5, opened bars) and LP-BM5–infected IDO−/− mice (n = 5, closed bars) were resected at the indicated weeks. CD4+CD25+ cells (shown in gated percentages) obtained from the total spleen were analyzed using FACScan (A). Total RNA was purified from unfractionated splenocytes, and Foxp3 (B), IL-4 (C), IL-10 (D), TGF-β (E), IFN-α (F), and IFN-β (G) mRNA were quantified using RT-qPCR. The levels of target mRNAs were normalized to those of 18s mRNA as an endogenous control. The levels of these mRNA were obtained from total spleen. Splenocytes at 8 wk after LP-BM5 infection were fractionated to yield MACS-enriched CD11c+ and CD4+ cells. The levels of type I IFN mRNA in CD11c+ cells and CD4+ cells were determined (H, I). These results are presented as the mean ± SD obtained from five mice at each time point. *p < 0.05 versus uninfected control group. qPCR, quantitative PCR.

FIGURE 3.
FIGURE 3.

Effect of IDO gene deficiency or inhibition increases type I IFN level in mice after LP-BM5 infection. WT, IDO−/−, and 1-MT–treated mice were killed at 8 wpi, and isolated CD11c-positive cells obtained from spleen of those mice were cultured at 72 h. IFN-α and -β detection was measured by using ELISA. Data are presented as mean ± SD (n = 5 in each group).

Effect of IDO gene deficiency on the phenotype of splenocytes after LP-BM5 infection

To identify the cells producing type I IFNs for virus clearance in IDO−/− mice infected with LP-BM5, we used flow cytometric analysis to quantify the number of NK cells (DX5+CD3−), NKT cells (NKTs) (DX5+CD3+), CD4+ cells, CD8+ cells, total DCs (CD11c+), conventional DCs (CD11chighB220−), and pDCs (CD11clowpDC Ag-1+) in spleen from both WT and IDO−/− mice at 8 wpi (Fig. 4). Interestingly, the number of CD11c+ cells in IDO−/− mice was increased compared with that in WT mice, and the number and gated percentage of pDCs in IDO−/− mice, which are the main source of type I IFNs, were also significantly increased. In addition, to address the other antiviral effects, we further assessed whether the number of NK, NKTs, CD4+, and CD8+ cells contributed to defense against LP-BM5 infection. The numbers of NKT, CD4+, and CD8+ cells in IDO−/− mice were significantly higher than those in WT mice, and the number of NK cells in IDO−/− mice tended to increase compared with that in WT mice. Further, the proliferation of Con A-treated splenocytes from 1-MT–treated or IDO−/− mice was dramatically enhanced compared with the proliferation of these splenocytes from WT mice in an ex vivo model (Fig. 5).

FIGURE 4.
FIGURE 4.

Increased number of pDCs in spleen from IDO−/− mice after LP-BM5 infection. Splenocytes from uninfected WT (white bars) and IDO−/− (black bars) or LP-BM5–infected WT (white striped bars) and IDO−/− (black striped bars) mice were resected at 8 wk postinfection. The number (mean × 107) and gated percentage of CD4+ cells, CD8+ cells, NK cells, NKTs, CD11c+ cells cDCs, and pDCs were quantified using FACScan analysis (BD Biosciences). Results are presented as the mean ± SD, the number, and the percentage of each cell type from spleen from at least three mice per group. KO, knockout.

FIGURE 5.
FIGURE 5.

Proliferation of splenocytes from WT, 1-MT–treated, and IDO−/− mice after LP-BM5 infection. WT mice, IDO−/− mice, and 1-MT–treated mice were killed at 8 wpi, and the isolated splenocytes obtained from these mice were cultured for 48 h. Con A response was set up as described in Materials and Methods. Data are presented as mean ± SD (n = 5 in each group). *p < 0.001 determined using one-way ANOVA for LP-BM5–infected WT mice versus LP-BM5–infected IDO−/− or 1-MT–treated mice.

Effect of IDO gene deficiency on viral copies and survival after LP-BM5 infection

The time course of the virus copies following LP-BM5 infection in spleen from WT and IDO−/− mice is shown in Fig. 6A. The number of BM5 provirus copies in the spleen from IDO−/− mice was significantly lower than that from WT mice at 8 wpi and 12 wpi, although the number of virus copies in the spleen from IDO−/− mice did not change compared with that from WT mice at 1, 2, and 4 wk after LP-BM5 infection. In addition, the number of virus copies in 1-MT–treated mice was significantly suppressed compared with nontreated mice at 8 wpi (Fig. 6B). We also assessed the role of upregulated type I IFN in the antiviral effect (Fig. 7). LP-BM5 viral replication was significantly increased in anti-type I IFN Ab-treated IDO−/− mice compared with control IgG-treated IDO−/− mice. The results obtained showed that upregulated type I IFNs mediated the inhibition of LP-BM5 viral replication in IDO−/− mice.

FIGURE 6.
FIGURE 6.

Comparison of LP-BM5 viral copies in spleen from WT, 1-MT–treated, and IDO−/− mice. A, WT (open bars) and IDO−/− mice (striped bars) were infected with MuLV and sacrificed at the indicated weeks. B, The viral copies in spleen from WT, IDO−/−, and 1-MT–treated mice were determined at 8 wpi. Virus copies shown were quantified using RT-qPCR. These results are presented as the mean ± SD obtained from five mice at each time point. qPCR, quantitative PCR.

FIGURE 7.
FIGURE 7.

Effect of anti-type I IFN Ab treatment on LP-BM5 viral copy in splenocytes from LP-BM5–infected mice. The splenocytes from LP-BM5–infected WT mice or from IDO−/− mice were treated with anti–IFN-α Ab (1000 neutralizing units/ml) alone or anti–IFN-β Ab alone (1000 neutralizing units/ml), with a combination of anti–IFN-α and -β (1000 neutralizing units/ml, respectively) or with control IgG for 72 h. LP-BM5 viral copies were analyzed by RT-PCR. Data are presented as mean ± SD (n = 5 in each group). *p < 0.05; **p < 0.01 versus control IgG-treated splenocytes from IDO−/− mice.

To determine if IDO would regulate virus infection lethality, we administered the LP-BM5 virus or T. gondii to compare both IDO−/− mice and WT mice (Fig. 8A). In addition, to assess the influence of IDO in the opportunistic infection model, we administered both T. gondii and the LP-BM5 virus to IDO−/− mice versus WT mice (Fig. 8B) or 1-MT–treated mice versus WT mice (Fig. 8C). T. gondii, an intracellular protozoan, is a major pathogen of the opportunistic infectious disease toxoplasmosis in infants and in immunocompromised hosts, such as patients with AIDS. The survival rate in IDO−/− mice after LP-BM5 virus single infection tended to increase compared with that in WT mice (p = 0.07); however, a single T. gondii infection was not potentially lethal. Importantly, although all WT mice died by 12 wpi, all of the IDO−/− mice survived considerably longer, up to 15 wpi. Moreover, the survival rates of 1-MT–treated mice as well as that of the IDO−/− mice were significantly greater than that of WT mice. These results indicate that the inhibition of IDO may slow the virus infection process, at least in this model system.

FIGURE 8.
FIGURE 8.

Reduced survival rates in the deficiency and blockade of IDO after LP-BM5 infection. A, WT (n = 15 in each group) and IDO−/− (n = 15 in each group) mice were inoculated with LP-BM5 or T. gondii alone. B, WT (n = 15) and IDO−/− (n = 15) mice were inoculated with both LP-BM5 and T. gondii. C, WT (n = 15) and 1-MT–treated (n = 15) mice were also infected with both LP-BM5 and T. gondii. The survival rate was monitored daily for 150 d. The difference in survival rates was confirmed using the Wilcox log-rank test.

Discussion

The present study demonstrates that upregulated type I IFNs suppressed virus replication after LP-BM5 infection in both IDO−/− and 1-MT–treated mice compared with WT mice.

Although the role of IDO after viral infection is not fully understood, our present study demonstrated that IDO−/− mice or IDO inhibitor-administrated mice clearly suppressed the LP-BM5 viral replication (Fig. 6). Further, recent studies have demonstrated that CTLA-4 blockade decreases TGF-β, IDO, and viral RNA expression in tissues of SIVmac251-infected macaques (17), and inhibition of IDO with the competitive molecule 1-MT may be a promi-sing candidate for enhancing immune responses, including anti-HIV immunity, in HIV-infected patients (18). Taken together, these results perhaps demonstrated that the inhibition of IDO is a potential therapeutic strategy for the treatment of retrovirus infection.

The role of IDO may depend on the difference of stimulus or tissue. For example, in the CNS, it has been reported that IDO activity in astrocytes is induced by TLR3 ligand polyinosinic-polycytidylic acid and causes an antivirus response (19). Additionally, both the abrogation of IDO activity by 1-MT and the supplementation of cultures with L-TRP result in the inhibition of IFN-γ–induced antimicrobial effects mediated by lung cells (20). However, it is notable that these findings were shown in an in vitro model.

Dramatic changes in serum tryptophan catabolism and IDO activity were seen in the spleen after LP-BM5 infection (Fig. 1). Further, we observed the enhanced expression of IDO in the spleen, particularly in the CD11c+ cells, after LP-BM5 infection. IDO is induced by proinflammatory cytokines, including type II IFN (3), and we have also demonstrated that a significant synergistic IDO induction occurring in THP-1 cells that were cultured in the presence of a combination of TNF-α, IL-6, or the β form of pro-IL-1, although TNF-α, IL-6, or the β form of pro-IL-1 alone did not induce IDO in THP-1 cells (21). Further, engagement of CTLA-4 with CD80/CD86 on the membrane of DCs stimulates IDO transcriptional expression and activity (4, 22). A previous study demonstrated that the spleens from LP-BM5–infected mice produced several different cytokines, such as TNF-α and IFN-γ (23). Further, it has been shown that IDO is expressed in human and mouse tumor cells (24), DCs, and macrophages following retrovirus infection, such as human T cell leukemia virus-1 and HIV (25). In particular, a recent study has demonstrated that monocyte-derived DC is clearly enhanced IDO via TLR7, which recognizes ssRNA viruses (26).

Importantly, our results demonstrated the induction of markedly increased type I IFNs in spleen from IDO−/− and 1-MT–treated mice after LP-BM5 infection (Figs. 2, 3), and we observed suppressed LP-BM5 viral replication and increased mortality (Figs. 6, 8). Innate defense occurs when pathogens contact or invade host cells and elicit the production of cytokines and chemokines, which in turn induce an influx of immune cells that affect pathogen clearance. In particular, type I IFNs are critical mediators of innate immunity and limit disease caused by many viruses (27). Indeed, it has been suggested that IFN-α/β considerably slows the development of LP-BM5 infection (28). In contrast, a recent study has shown that although IDO inhibition increases proinflammatory cytokine production in mice with fungi, it greatly exacerbates infection and associated inflammatory pathology as a result of deregulated innate and adaptive/regulatory immune responses (29). These findings also demonstrated that tryptophan metabolites were capable of inducing the Foxp3-encoding gene transcriptionally and suppressing the gene encoding RORγt, a Th17 lineage specification factor, in the fungal infection model (30).

Therefore, the role of IDO may be complex and may depend on the difference of disease stages (e.g., acute/chronic disease) or the stimulus pathogens. In fact, prevention of efficient immune responses may also favor chronic pathologic disorders, such as persistent infections and cancer. In this context, IDO has been suggested to play a role in tumor immune escape and pathogen immune evasion, mainly through the generation of a tolerogenic environment limiting effective immunity (31, 32). Such an immunosuppressive environment likely occurs in HIV-1 infection, in which prolonged activation of pDC by the virus, through a CD4/gp120 interaction, might result in long-term suppression of T cell responses and immune exhaustion through activation of IDO (33).

We showed that upregulated type I IFNs in 1-MT–treated mice or IDO−/− mice suppress LP-BM5 virus replication (Fig. 7), and these findings resulted in an increased survival rate postinfection with LP-BM5 alone or infection with LP-BM5 and T. gondii (Fig. 8). The survival rate in IDO−/− mice after LP-BM5 single infection tended to increase compared with that in WT mice, although the difference was not statistically significant (Fig. 8A). It has been shown that LP-BM5–infected mice developed a B cell lymphoma at late phase (34). Although we do not exclude the possibility that type I IFNs may be not effective against LP-BM5–induced lymphoma, at least in this model, we speculate that upregulated type I IFNs are effective against pathogens, particularly viruses.

Because the source of upregulated type I IFNs in IDO−/− mice is still unknown at present, our results as one important mechanism may support that upregulated type I IFNs were due to increased pDCs (Fig. 4). pDCs are immature DCs that participate in both innate and adaptive immunity (35), and they are specialized cells located in blood and lymphoid organs that produce up to 1000-fold more type I IFNs than other cell types in response to virus exposure (36). In addition, a strong increase in pDC density in the T cell zone in lymph nodes of asymptomatic HIV-infected patients was reported in an earlier study (37). Furthermore, it has been recently shown that pDCs are indeed chronically stimulated in HIV-infected patients and produce type I IFNs, which in turn contributes to their hyporesponsiveness to subsequent in vitro stimulation, probably through a negative regulatory feedback mechanism (38).

Viral activation of pDCs can be regulated by either of the two TLRs, TLR7 or TLR9, which are considered to be the receptors that human and mouse pDCs use for recognition of RNA/retroviruses (39) and DNA (40), respectively. A recent study showed that ablation of IDO did not block IFN-α production by CD19+ DC after TLR9 ligation, but ablation of IDO completely abrogated IFN-α production by CD19+ DCs after B7 ligation (41). Therefore, it is possible that induction of type I IFNs in pDCs via TLR–LP-BM5 viral RNA interactions may be induced by the ablation of IDO. Indeed, it has been suggested that retrovirus activates pDCs via TLR–viral RNA interactions (42).

As shown in Fig. 4, although it is unknown why the number of pDCs in IDO−/− mice increases after LP-BM5 infection, we also showed that there was a dramatic increase in the proliferation of splenocytes from 1-MT–treated WT mice or IDO−/− mice with LP-BM5 infection compared with the proliferation of splenocytes from untreated WT mice (Fig. 5). Therefore, we speculate that this increase indicates an escape from IDO-mediated immunosuppression. In fact, two major theories of L-TRP catabolism have recently been proposed to account for tolerance induction via L-TRP catabolism. According to one theory, downstream metabolites of L-TRP (collectively known as L-KYN) act to suppress immune reactivity through direct interactions with effector T lymphocytes and other cell types (7–9). According to another theory, L-TRP itself plays a key role, and breakdown of this molecule suppresses T cell proliferation by critically reducing the availability of this indispensable amino acid under local tissue microenvironments (6).

Although one study reported that the cytotoxic action of tryptophan downstream metabolites affected preferentially activated T cells, B cells, and NK cells but not DCs themselves in vitro (9), it is possible that our present results indicate that tryptophan downstream metabolites affect T cells, B cells, NK cells, and DCs in the in vivo model. In fact, the total numbers of splenocytes from IDO−/− mice were clearly increased compared with those from WT mice (Fig. 4).

In conclusion, the results of the current study provide, to our knowledge, the first evidence that the absence and inhibition of IDO is critical for the suppression of murine retroviral infection and upregulated type I IFNs. Our study offers insights into the role of innate and adaptive immune response cytokines in control of viral dissemination in the host.

Acknowledgments

We thank Drs. Yoshitatsu Sei, Shizuko Sei, and Suwako Fujigaki for excellent advice.

Disclosures The authors have no financial conflicts of interest.

Footnotes

  • This work was supported by Grant-in-Aid for Scientific Research 20390167 from the Ministry for Education, Culture, Sports, Science and Technology of Japan.

  • Abbreviations used in this paper:

    DC
    dendritic cell
    KO
    knockout
    L-TRP
    l-tryptophan
    1-MT
    1-methyl-d-l-tryptophan
    MuLV
    murine leukemia virus
    NKT
    NKT cell
    pDC
    plasmacytoid dendritic cell
    qPCR
    quantitative PCR
    Treg
    regulatory T cell
    wpi
    wk postinfection
    WT
    wild-type.

  • Received April 10, 2009.
  • Accepted June 24, 2010.

References

  1. ↵
    1. Munn D. H.,
    2. E. Shafizadeh,
    3. J. T. Attwood,
    4. I. Bondarev,
    5. A. Pashine,
    6. A. L. Mellor
    . 1999. Inhibition of T cell proliferation by macrophage tryptophan catabolism. J. Exp. Med. 189: 1363–1372.
    OpenUrlAbstract/FREE Full Text
  2. ↵
    1. Mellor A. L.,
    2. D. H. Munn
    . 1999. Tryptophan catabolism and T-cell tolerance: immunosuppression by starvation? Immunol. Today 20: 469–473.
    OpenUrlCrossRefPubMed
  3. ↵
    1. Taylor M. W.,
    2. G. S. Feng
    . 1991. Relationship between interferon-gamma, indoleamine 2,3-dioxygenase, and tryptophan catabolism. FASEB J. 5: 2516–2522.
    OpenUrlAbstract
  4. ↵
    1. Fallarino F.,
    2. U. Grohmann,
    3. K. W. Hwang,
    4. C. Orabona,
    5. C. Vacca,
    6. R. Bianchi,
    7. M. L. Belladonna,
    8. M. C. Fioretti,
    9. M. L. Alegre,
    10. P. Puccetti
    . 2003. Modulation of tryptophan catabolism by regulatory T cells. Nat. Immunol. 4: 1206–1212.
    OpenUrlCrossRefPubMed
  5. ↵
    1. Fallarino F.,
    2. R. Bianchi,
    3. C. Orabona,
    4. C. Vacca,
    5. M. L. Belladonna,
    6. M. C. Fioretti,
    7. D. V. Serreze,
    8. U. Grohmann,
    9. P. Puccetti
    . 2004. CTLA-4-Ig activates forkhead transcription factors and protects dendritic cells from oxidative stress in nonobese diabetic mice. J. Exp. Med. 200: 1051–1062.
    OpenUrlAbstract/FREE Full Text
  6. ↵
    1. Lee G. K.,
    2. H. J. Park,
    3. M. Macleod,
    4. P. Chandler,
    5. D. H. Munn,
    6. A. L. Mellor
    . 2002. Tryptophan deprivation sensitizes activated T cells to apoptosis prior to cell division. Immunology 107: 452–460.
    OpenUrlCrossRefPubMed
  7. ↵
    1. Frumento G.,
    2. R. Rotondo,
    3. M. Tonetti,
    4. G. Damonte,
    5. U. Benatti,
    6. G. B. Ferrara
    . 2002. Tryptophan-derived catabolites are responsible for inhibition of T and natural killer cell proliferation induced by indoleamine 2,3-dioxygenase. J. Exp. Med. 196: 459–468.
    OpenUrlAbstract/FREE Full Text
    1. Platten M.,
    2. P. P. Ho,
    3. S. Youssef,
    4. P. Fontoura,
    5. H. Garren,
    6. E. M. Hur,
    7. R. Gupta,
    8. L. Y. Lee,
    9. B. A. Kidd,
    10. W. H. Robinson,
    11. et al
    . 2005. Treatment of autoimmune neuroinflammation with a synthetic tryptophan metabolite. Science 310: 850–855.
    OpenUrlAbstract/FREE Full Text
  9. ↵
    1. Terness P.,
    2. T. M. Bauer,
    3. L. Röse,
    4. C. Dufter,
    5. A. Watzlik,
    6. H. Simon,
    7. G. Opelz
    . 2002. Inhibition of allogeneic T cell proliferation by indoleamine 2,3-dioxygenase-expressing dendritic cells: mediation of suppression by tryptophan metabolites. J. Exp. Med. 196: 447–457.
    OpenUrlAbstract/FREE Full Text
  10. ↵
    1. Aziz D. C.,
    2. Z. Hanna,
    3. P. Jolicoeur
    . 1989. Severe immunodeficiency disease induced by a defective murine leukaemia virus. Nature 338: 505–508.
    OpenUrlCrossRefPubMed
  11. ↵
    1. Mosier D. E.,
    2. R. A. Yetter,
    3. H. C. Morse III.
    . 1985. Retroviral induction of acute lymphoproliferative disease and profound immunosuppression in adult C57BL/6 mice. J. Exp. Med. 161: 766–784.
    OpenUrlAbstract/FREE Full Text
  12. ↵
    1. Iida R.,
    2. K. Saito,
    3. K. Yamada,
    4. A. S. Basile,
    5. K. Sekikawa,
    6. M. Takemura,
    7. H. Fujii,
    8. H. Wada,
    9. M. Seishima,
    10. T. Nabeshima
    . 2000. Suppression of neurocognitive damage in LP-BM5-infected mice with a targeted deletion of the TNF-alpha gene. FASEB J. 14: 1023–1031.
    OpenUrlAbstract/FREE Full Text
  13. ↵
    1. Fujigaki S.,
    2. K. Saito,
    3. M. Takemura,
    4. N. Maekawa,
    5. Y. Yamada,
    6. H. Wada,
    7. M. Seishima
    . 2002. L-tryptophan-L-kynurenine pathway metabolism accelerated by Toxoplasma gondii infection is abolished in gamma interferon-gene-deficient mice: cross-regulation between inducible nitric oxide synthase and indoleamine-2,3-dioxygenase. Infect. Immun. 70: 779–786.
    OpenUrlAbstract/FREE Full Text
  14. ↵
    1. Hoshi M.,
    2. H. Ito,
    3. H. Fujigaki,
    4. M. Takemura,
    5. T. Takahashi,
    6. E. Tomita,
    7. M. Ohyama,
    8. R. Tanaka,
    9. K. Saito,
    10. M. Seishima
    . 2009. Indoleamine 2,3-dioxygenase is highly expressed in human adult T-cell leukemia/lymphoma and chemotherapy changes tryptophan catabolism in serum and reduced activity. Leuk. Res. 33: 39–45.
    OpenUrlCrossRefPubMed
  15. ↵
    1. Hoshi M.,
    2. K. Saito,
    3. Y. Murakami,
    4. A. Taguchi,
    5. H. Fujigaki,
    6. R. Tanaka,
    7. M. Takemura,
    8. H. Ito,
    9. A. Hara,
    10. M. Seishima
    . 2009. Marked increases in hippocampal neuron indoleamine 2, 3-dioxygenase via IFN-gamma-independent pathway following transient global ischemia in mouse. Neurosci. Res. 63: 194–198.
    OpenUrlCrossRefPubMed
  16. ↵
    1. Casabianca A.,
    2. C. Orlandi,
    3. A. Fraternale,
    4. M. Magnani
    . 2004. Development of a real-time PCR assay using SYBR Green I for provirus load quantification in a murine model of AIDS. J. Clin. Microbiol. 42: 4361–4364.
    OpenUrlAbstract/FREE Full Text
  17. ↵
    1. Hryniewicz A.,
    2. A. Boasso,
    3. Y. Edghill-Smith,
    4. M. Vaccari,
    5. D. Fuchs,
    6. D. Venzon,
    7. J. Nacsa,
    8. M. R. Betts,
    9. W. P. Tsai,
    10. J. M. Heraud,
    11. et al
    . 2006. CTLA-4 blockade decreases TGF-beta, IDO, and viral RNA expression in tissues of SIVmac251-infected macaques. Blood 108: 3834–3842.
    OpenUrlAbstract/FREE Full Text
  18. ↵
    1. Boasso A.,
    2. J. P. Herbeuval,
    3. A. W. Hardy,
    4. S. A. Anderson,
    5. M. J. Dolan,
    6. D. Fuchs,
    7. G. M. Shearer
    . 2007. HIV inhibits CD4+ T-cell proliferation by inducing indoleamine 2,3-dioxygenase in plasmacytoid dendritic cells. Blood 109: 3351–3359.
    OpenUrlAbstract/FREE Full Text
  19. ↵
    1. Suh H. S.,
    2. M. L. Zhao,
    3. M. Rivieccio,
    4. S. Choi,
    5. E. Connolly,
    6. Y. Zhao,
    7. O. Takikawa,
    8. C. F. Brosnan,
    9. S. C. Lee
    . 2007. Astrocyte indoleamine 2,3-dioxygenase is induced by the TLR3 ligand poly(I:C): mechanism of induction and role in antiviral response. J. Virol. 81: 9838–9850.
    OpenUrlAbstract/FREE Full Text
  20. ↵
    1. Heseler K.,
    2. K. Spekker,
    3. S. K. Schmidt,
    4. C. R. MacKenzie,
    5. W. Däubener
    . 2008. Antimicrobial and immunoregulatory effects mediated by human lung cells: role of IFN-gamma-induced tryptophan degradation. FEMS Immunol. Med. Microbiol. 52: 273–281.
    OpenUrlAbstract/FREE Full Text
  21. ↵
    1. Fujigaki H.,
    2. K. Saito,
    3. S. Fujigaki,
    4. M. Takemura,
    5. K. Sudo,
    6. H. Ishiguro,
    7. M. Seishima
    . 2006. The signal transducer and activator of transcription 1alpha and interferon regulatory factor 1 are not essential for the induction of indoleamine 2,3-dioxygenase by lipopolysaccharide: involvement of p38 mitogen-activated protein kinase and nuclear factor-kappaB pathways, and synergistic effect of several proinflammatory cytokines. J. Biochem. 139: 655–662.
    OpenUrlAbstract/FREE Full Text
  22. ↵
    1. Grohmann U.,
    2. F. Fallarino,
    3. R. Bianchi,
    4. C. Orabona,
    5. C. Vacca,
    6. M. C. Fioretti,
    7. P. Puccetti
    . 2003. A defect in tryptophan catabolism impairs tolerance in nonobese diabetic mice. J. Exp. Med. 198: 153–160.
    OpenUrlAbstract/FREE Full Text
  23. ↵
    1. Cheung S. C.,
    2. S. K. Chattopadhyay,
    3. H. C. Morse III.,
    4. P. M. Pitha
    . 1991. Expression of defective virus and cytokine genes in murine AIDS. J. Virol. 65: 823–828.
    OpenUrlAbstract/FREE Full Text
  24. ↵
    1. Uyttenhove C.,
    2. L. Pilotte,
    3. I. Théate,
    4. V. Stroobant,
    5. D. Colau,
    6. N. Parmentier,
    7. T. Boon,
    8. B. J. Van den Eynde
    . 2003. Evidence for a tumoral immune resistance mechanism based on tryptophan degradation by indoleamine 2,3-dioxygenase. Nat. Med. 9: 1269–1274.
    OpenUrlCrossRefPubMed
  25. ↵
    1. Grant R. S.,
    2. H. Naif,
    3. S. J. Thuruthyil,
    4. N. Nasr,
    5. T. Littlejohn,
    6. O. Takikawa,
    7. V. Kapoor
    . 2000. Induction of indolamine 2,3-dioxygenase in primary human macrophages by human immunodeficiency virus type 1 is strain dependent. J. Virol. 74: 4110–4115.
    OpenUrlAbstract/FREE Full Text
  26. ↵
    1. Furset G.,
    2. Y. Fløisand,
    3. M. Sioud
    . 2008. Impaired expression of indoleamine 2, 3-dioxygenase in monocyte-derived dendritic cells in response to Toll-like receptor-7/8 ligands. Immunology 123: 263–271.
    OpenUrlPubMed
  27. ↵
    1. Hwang S. Y.,
    2. P. J. Hertzog,
    3. K. A. Holland,
    4. S. H. Sumarsono,
    5. M. J. Tymms,
    6. J. A. Hamilton,
    7. G. Whitty,
    8. I. Bertoncello,
    9. I. Kola
    . 1995. A null mutation in the gene encoding a type I interferon receptor component eliminates antiproliferative and antiviral responses to interferons alpha and beta and alters macrophage responses. Proc. Natl. Acad. Sci. USA 92: 11284–11288.
    OpenUrlAbstract/FREE Full Text
  28. ↵
    1. Heng J. K.,
    2. P. Price,
    3. C. M. Lai,
    4. M. W. Beilharz
    . 1996. Alpha/beta interferons increase host resistance to murine AIDS. J. Virol. 70: 4517–4522.
    OpenUrlAbstract/FREE Full Text
  29. ↵
    1. Bozza S.,
    2. F. Fallarino,
    3. L. Pitzurra,
    4. T. Zelante,
    5. C. Montagnoli,
    6. S. Bellocchio,
    7. P. Mosci,
    8. C. Vacca,
    9. P. Puccetti,
    10. L. Romani
    . 2005. A crucial role for tryptophan catabolism at the host/Candida albicans interface. J. Immunol. 174: 2910–2918.
    OpenUrlAbstract/FREE Full Text
  30. ↵
    1. De Luca A.,
    2. C. Montagnoli,
    3. T. Zelante,
    4. P. Bonifazi,
    5. S. Bozza,
    6. S. Moretti,
    7. C. D’Angelo,
    8. C. Vacca,
    9. L. Boon,
    10. F. Bistoni,
    11. et al
    . 2007. Functional yet balanced reactivity to Candida albicans requires TRIF, MyD88, and IDO-dependent inhibition of Rorc. J. Immunol. 179: 5999–6008.
    OpenUrlAbstract/FREE Full Text
  31. ↵
    1. Puccetti P.,
    2. U. Grohmann
    . 2007. IDO and regulatory T cells: a role for reverse signalling and non-canonical NF-kappaB activation. Nat. Rev. Immunol. 7: 817–823.
    OpenUrlCrossRefPubMed
  32. ↵
    1. Mellor A. L.,
    2. D. H. Munn
    . 2008. Creating immune privilege: active local suppression that benefits friends, but protects foes. Nat. Rev. Immunol. 8: 74–80.
    OpenUrlCrossRefPubMed
  33. ↵
    1. Boasso A.,
    2. G. M. Shearer
    . 2008. Chronic innate immune activation as a cause of HIV-1 immunopathogenesis. Clin. Immunol. 126: 235–242.
    OpenUrlCrossRefPubMed
  34. ↵
    1. Klinken S. P.,
    2. T. N. Fredrickson,
    3. J. W. Hartley,
    4. R. A. Yetter,
    5. H. C. Morse III.
    . 1988. Evolution of B cell lineage lymphomas in mice with a retrovirus-induced immunodeficiency syndrome, MAIDS. J. Immunol. 140: 1123–1131.
    OpenUrlAbstract
  35. ↵
    1. Siegal F. P.,
    2. N. Kadowaki,
    3. M. Shodell,
    4. P. A. Fitzgerald-Bocarsly,
    5. K. Shah,
    6. S. Ho,
    7. S. Antonenko,
    8. Y. J. Liu
    . 1999. The nature of the principal type 1 interferon-producing cells in human blood. Science 284: 1835–1837.
    OpenUrlAbstract/FREE Full Text
  36. ↵
    1. Kadowaki N.,
    2. S. Antonenko,
    3. J. Y. Lau,
    4. Y. J. Liu
    . 2000. Natural interferon alpha/beta-producing cells link innate and adaptive immunity. J. Exp. Med. 192: 219–226.
    OpenUrlAbstract/FREE Full Text
  37. ↵
    1. Foussat A.,
    2. L. Bouchet-Delbos,
    3. D. Berrebi,
    4. I. Durand-Gasselin,
    5. A. Coulomb-L’Hermine,
    6. R. Krzysiek,
    7. P. Galanaud,
    8. Y. Levy,
    9. D. Emilie
    . 2001. Deregulation of the expression of the fractalkine/fractalkine receptor complex in HIV-1-infected patients. Blood 98: 1678–1686.
    OpenUrlAbstract/FREE Full Text
  38. ↵
    1. Tilton J. C.,
    2. M. M. Manion,
    3. M. R. Luskin,
    4. A. J. Johnson,
    5. A. A. Patamawenu,
    6. C. W. Hallahan,
    7. N. A. Cogliano-Shutta,
    8. J. M. Mican,
    9. R. T. Davey Jr..,
    10. S. Kottilil,
    11. et al
    . 2008. Human immunodeficiency virus viremia induces plasmacytoid dendritic cell activation in vivo and diminished alpha interferon production in vitro. J. Virol. 82: 3997–4006.
    OpenUrlAbstract/FREE Full Text
  39. ↵
    1. Diebold S. S.,
    2. T. Kaisho,
    3. H. Hemmi,
    4. S. Akira,
    5. C. Reis e Sousa
    . 2004. Innate antiviral responses by means of TLR7-mediated recognition of single-stranded RNA. Science 303: 1529–1531.
    OpenUrlAbstract/FREE Full Text
  40. ↵
    1. Coccia E. M.,
    2. M. Severa,
    3. E. Giacomini,
    4. D. Monneron,
    5. M. E. Remoli,
    6. I. Julkunen,
    7. M. Cella,
    8. R. Lande,
    9. G. Uzé
    . 2004. Viral infection and Toll-like receptor agonists induce a differential expression of type I and lambda interferons in human plasmacytoid and monocyte-derived dendritic cells. Eur. J. Immunol. 34: 796–805.
    OpenUrlCrossRefPubMed
  41. ↵
    1. Manlapat A. K.,
    2. D. J. Kahler,
    3. P. R. Chandler,
    4. D. H. Munn,
    5. A. L. Mellor
    . 2007. Cell-autonomous control of interferon type I expression by indoleamine 2,3-dioxygenase in regulatory CD19+ dendritic cells. Eur. J. Immunol. 37: 1064–1071.
    OpenUrlCrossRefPubMed
  42. ↵
    1. Beignon A. S.,
    2. K. McKenna,
    3. M. Skoberne,
    4. O. Manches,
    5. I. DaSilva,
    6. D. G. Kavanagh,
    7. M. Larsson,
    8. R. J. Gorelick,
    9. J. D. Lifson,
    10. N. Bhardwaj
    . 2005. Endocytosis of HIV-1 activates plasmacytoid dendritic cells via Toll-like receptor-viral RNA interactions. J. Clin. Invest. 115: 3265–3275.
    OpenUrlCrossRefPubMed
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