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Unité de Biologie des Infections Virales Emergentes, Centre de Recherche Mérieux-Pasteur à Lyon, Laboratoire P4-Jean Mérieux, Lyon, France
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Abstract |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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-dystroglycan receptor (4, 5). Humans become infected through contact with infected excreta, tissue, or blood from the peridomestic rodent Mastomys sp., which is the reservoir host of LV (1). Human-to-human transmission of LV occurs via mucosal/cutaneous contact or nosocomial contamination (6). The severity of the disease is variable, from asymptomatic infection to fatal hemorrhagic fever. After an incubation period of 6–12 days, nonspecific signs appear with fever, headache, arthralgia, myalgia, and severe asthenia. Abdominal pain, pharyngia, cough, conjunctivitis, diarrhea, and vomiting are observed in the next few days, and the most severe cases may be associated with cervical and facial edema, hemorrhages, renal and liver failures, and encephalopathy. Death may occur in the context of hypotensive, hypovolemic, and hypoxic shock. In surviving patients, symptoms disappear 10–15 days after onset (7). However, deafness is a common complication of Lassa fever, affecting about one-third of survivors (8). Macrophages (MP) and endothelial cells are the main targets for LV (9). However, despite widespread viral replication, changes in the endothelium or other organs are not severe enough to account for terminal shock and death (10), which seem instead to depend on host response.
Little is known about the immune responses induced during LV infection. Humoral responses cannot control viral replication and the appearance of specific IgG in patients and infected monkeys is not correlated with recovery (11, 12). The induction of cellular responses specific for viral glycoproteins protects nonhuman primates from a lethal challenge (12), and memory CD4+ T cell responses against NP have been demonstrated in seropositive endemic subjects (13). In contrast, severe LV infections seem to be associated with high levels of viremia and immunosuppression. Structural changes, the cellular depletion of secondary lymphoid tissues, necrosis of the splenic marginal zone, transitory lymphopenia, and the abolition of mitogenic T cell proliferation have been described in patients and nonhuman primates (7, 14, 15). The infection of MP in vitro leads to the release of viral particles, but not to an increase in the synthesis of TNF- and IL-8 (9). Finally, significant levels of inflammatory cytokines have been detected in the plasma of patients with nonfatal Lassa fever, but not in patients who died (16).
Dendritic cells (DC) may be a crucial target of LV, as observed for other Arenaviruses, such as the Junin virus (17) and lymphocytic choriomeningitis virus (LCMV) (18). As DC and MP play a major role in the induction and regulation of innate and adaptive immune responses (19), the tropism of LV for APC may be involved in immunosuppression, as observed in LCMV infection (20). In this study, we determined and compared the susceptibility and responses to LV infection of two types of APC: human monocyte-derived DC and MP.
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Materials and Methods |
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LV (AV strain (21)) was cultured in the Vero E6 cell line at 37°C. Viruses from the serum of a patient were subjected to four passages on Vero E6 cells. The cell-free supernatant, with a viral titer of 107 focus-forming units (FFU)/ml was then used as the infectious virus stock. This supernatant was also irradiated (5 x 106 rad) and used as inactivated LV (22). We checked that cell lines and viruses were not contaminated with mycoplasmas. All experiments with infectious viruses were conducted in Biosafety Level 4 facilities.
Virus titration
Vero E6 cells were incubated for 1 h at 37°C with several dilutions of cell-free supernatants and 1.6% carboxy-methyl-cellulose (BDH Laboratory Supplies, Poole, U.K.) in DMEM (Life Technologies, Cergy-Pontoise, France) was then added. Infectious foci were detected after 5 days of culture, by incubation with a mixture of mAbs against LV NP (mAbs 52-158-3, 52-54-6, and 52-189-13 (23), generously provided by Dr P. Jahrling, U.S. Army Medical Research Institute of Infectious Diseases, Fort Detrick, MD) followed by HRP-conjugated goat polyclonal anti-mouse IgG (Sigma-Aldrich, Saint Quentin Fallavier, France) and di-amino-benzidine. Results are expressed as FFU per million cells.
Obtention of DC and MP
Fresh human peripheral blood was obtained from the Etablissement Français du Sang (Lyon, France). Mononuclear cells were isolated by density gradient centrifugation on Ficoll-Paque (Pharmacia, Uppsala, Sweden), followed by centrifugation through a 50% Percoll gradient (Pharmacia) for 20 min at 400 x g. The low-density fraction from the interface was recovered, washed twice in RPMI 1640-Glutamax I (Life Technologies) supplemented with 10 mM HEPES (Life Technologies), 1% penicillin-streptomycin (Life Technologies), 1% nonessential amino acids (Life Technologies), and 10% FCS (Life Technologies) (C-RPMI), and incubated for 15 min in C-RPMI supplemented with 5% human AB serum (Etablissement Français du Sang). Monocytes were further purified by immunomagnetic depletion (Dynabeads; Dynal, Oslo, Norway) with a mixture of mAbs against CD3 (SPV-T3b; Dynal), CD19 (AB1; Dynal) and CD56 (C218; Immunotech, Marseille, France). DC were generated by culturing monocytes at a density of 106 cells/ml in C-RPMI supplemented with 1000 U/ml recombinant human (rh) GM-CSF (PeproTech, London, U.K.) and 500 U/ml rhIL-4 (PeproTech). We replaced 40% of the culture medium every 2 days and replenished the supply of cytokines. The cells were harvested after 7 days: 70–90% were CD1a-expressing immature DC. MP were obtained by culturing monocytes in C-RPMI supplemented with 10 U/ml rhM-CSF (PeproTech) for 7 days and replacing the medium and cytokines every 2 days. Mature DC were obtained by adding 500 ng/ml LPS (Sigma-Aldrich) to cell cultures 24 h before infection.
Infection of DC and MP by LV
Cell pellets were resuspended in cell-free supernatants containing infectious or inactivated LV, or with Vero E6 supernatant for mock infection. Cells were incubated for 1 h at 37°C with gentle shaking. They were then thoroughly washed in C-RPMI, and cultured at a density of 106 cells/ml in C-RPMI supplemented with the appropriate cytokines at 37°C. Medium and cytokines were renewed every 2 days. In some cases, DC and MP were activated 2 h after infection by adding 250 ng/ml soluble rhCD40 ligand (sCD40L) and 1 µg/ml enhancer (Alexis, Carlsbad, CA) for DC and 500 ng/ml LPS for MP.
Immunofluorescence
DC and MP were harvested 48 h after infection and centrifuged onto coated slides (Shandon, Cheshire, U.K.) in a Cytospin 3 cytocentrifuge (Shandon). Cells were fixed with 3% paraformaldehyde (PFA) in PBS for 10 min at room temperature (RT) and were incubated in 5% AB+ human serum for 15 min. CD1a- or CD14-specific mAbs (BD PharMingen, San Diego, CA) diluted 1/20 in PBS were then added and the cells were incubated for 1 h at RT. They were then incubated with rhodamine-conjugated goat anti-mouse IgG (Zymed, San Francisco, CA). Cells were permeabilized by incubation with 0.5% Triton X-100 (Sigma-Aldrich) in PBS for 5 min. A mixture of two FITC-conjugated mAbs specific for LV NP (52-54-6 and 52-189-13) was then added and the cells incubated for 1 h at RT. Slides were mounted in FluoPrep mounting medium (Dako, Trappes, France), and images were captured with Qfluoro software (Leica, Cambridge, U.K.).
Detection of mRNA by RT-PCR
Total RNA was isolated from 5 x 105 cells, using the RNeasy kit (Qiagen, Courtaboeuf, France) according to the manufacturer’s instructions. Contaminating genomic DNA was digested with DNase I (Qiagen) during the extraction procedure. First-strand cDNA was synthesized from 200 ng of RNA, using Superscript II reverse transcriptase (Life Technologies) and oligo(dT)(15) (Life Technologies); 10% of the products of this reaction were used as a template for qualitative and real-time PCR. We checked for the absence of genomic DNA amplification for each gene by adding RNA directly to the amplification mixture without reverse transcription. The following primers were used for qualitative PCR (annealing temperatures are indicated for each primer pair): 
-actin, 5'-CAGGCACCAGGGCGTGAT-3' and 5'-GCCAGCCAGGTCCAGACG-3', 60°C; IL-6, 5'-AGTTGCCTTCTCCCTGG-3' and 5'-ATTTGCCGAAGAGCCCTCA-3', 55°C; IL-12p40, 5'-GGATGCCCCTGGAGAAATGG-3' and 5'-CTCCCAGCTGACCTCCACCT-3', 60°C. DNA was amplified by PCR with TaqDNA polymerase (Roche, Mannheim, Germany) for 40 cycles (cytokines) or 35 cycles (actin). PCR products were separated by electrophoresis in a 1.5% agarose gel and visualized by ethidium bromide staining (data not shown). For real-time PCR, predeveloped primers and TaqMan probes for -actin, TNF-, IL-1, IL-10, IL-12p35, TGF, IFN-
, CCR5, CCR6, CCR7, macrophage inflammatory protein (MIP)-1, MIP-1, and IL-8 were used (Applied Biosystems, Courtaboeuf, France). DNA were amplified using TaqMan Universal Master Mix (Applied Biosystems) on an ABI PRISM 7000 real-time thermocycler (Applied Biosystems), according to the manufacturer’s instructions. DNA fragments encoding cytokines were amplified in duplex with -actin and relative mRNA levels for each sample were calculated as follows: 
cycle threshold (Ct) = Ct gene X - Ct -actin. Ratio (mRNA in infected cells/mRNA in mock-infected cells) = 2-(Ct infected - Ct mock).
ELISA detection of cytokines in supernatants
Supernatants from cultures of DC and MP were harvested, centrifuged, and stored at -80°C. Commercial ELISA kits were used for TNF-, IL-10, IL-1 (CLB, Amsterdam, The Netherlands), IL-12 (Cytimmune Sciences, College Park, MA), IL-8, MIP-1, MIP-1 (R&D Systems, Abingdon, U.K.) detection, according to the manufacturer’s instructions.
Flow cytometry
MP and DC were harvested in PBS containing 0.5 mM EDTA at various times after infection, incubated for 15 min in PBS-5% AB+ human serum, and stained for 30 min at 4°C with the following mAbs: FITC-conjugated anti-human CD14 (RMO52), FITC-conjugated anti-HLA-DR (B8.12.2), PE-conjugated anti-CD1a (BL6), PE-conjugated anti-CD40 (mAb89), PE-cyanin 5 (PC5)-conjugated anti-ILT3 (ZM3.8) (Immunotech, Marseille, France), FITC-conjugated anti-CD86 (FUN-1), PE-conjugated anti-CD80 (L307.4), PE-conjugated anti-CD83 (HB15e), CyChrome-conjugated anti-HLA-ABC (G46-2.6), CyChrome-conjugated anti-CD95 (DX2), CyChrome-conjugated anti-CD54 (HA58) (BD PharMingen). Isotypic controls were performed by staining cells with FITC-conjugated mouse IgG1 (MOPC-21), PE-conjugated IgG2a (G155-178), and CyChrome-conjugated IgG1 (MOPC-21) (BD PharMingen). The purity of monocyte populations was checked by staining cells with FITC-conjugated anti-CD8 (B9.11), PE-conjugated anti-CD19 (J4.119), and PC5-conjugated anti-CD3 (UCHT1) (Immunotech) Abs. Cells were washed in 2.5% FCS in PBS, and resuspended in 3% PFA in PBS. LV NP was detected by intracellular staining as follows. Cells were fixed and permeabilized using the Cytofix/Cytoperm kit (BD PharMingen), according to the manufacturer’s instructions. Cells were incubated for 30 min at 4°C with the mixture of two FITC-conjugated anti-LV NP mAbs described above or with FITC-conjugated mouse IgG1 (MOPC-21). They were then washed twice and resuspended in 3% PFA in PBS. Double staining for Annexin VFITC binding and 7-aminoactinomycin D (7AAD) (BD PharMingen) was performed according to the manufacturer’s instructions, to assess cell viability. Absolute counts were obtained with FlowCount-calibrated fluorospheres (Beckman Coulter, Villepinte, France), used according to the manufacturer’s instructions. Flow cytometry was performed in an EPICS-XL four-color cytometer (Beckman Coulter), with EXPO 32 ADC software (Beckman Coulter).
Internalization of dextran-FITC
DC were incubated for 1 h at 37°C (or at 4°C for control) with 20 µg/ml dextran-FITC (Sigma-Aldrich) in C-RPMI. Cells were harvested in cold PBS supplemented with 0.5 mM EDTA, washed in PBS, resuspended in 3% PFA in PBS, and analyzed by flow cytometry.
Chemotaxis assay
Cell migration was performed in 24-well Transwell cell culture chamber (Costar, Corning, NY). DC were harvested 72 h after mock or LV infection, and 105 cells in 100 µl of C-RPMI were added to each transwell insert. Six hundred microliters of serum-free medium containing 100 ng/ml rh MIP-1, MIP-3, or MIP-3 (Peprotech) were added to the lower compartment. In other experiments, immature DC migration assays toward supernatants of mock and LV-infected DC harvested 3 days after infection were performed. After 2 h of incubation at 37°C, the cells that migrated through the 8.0-µm pore size polycarbonate membranes in the lower compartment were collected and counted on a flow cytometer (EPICS-XL; Beckman Coulter) with FlowCount-calibrated fluorospheres (Beckman Coulter). The lower compartment of control chambers contained medium alone, while the absence of chemokinesis was demonstrated by the absence of cell migration when both compartments contained chemokines. Each assay was performed in duplicate and two independent experiments have been done.
Statistical analysis
Student’s t test was used to analyze differences between datasets for the assays. Statistical data were obtained using Excel software (Microsoft, Redmond, WA).
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Results |
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Immature DC and MP were infected with LV (strain AV (21)) at a multiplicity of infection (MOI) of 2 FFU/cell. Cells were harvested after 2 days, centrifuged onto coated slides, and double-stained with a mixture of two NP-specific mAbs and a phenotype-specific mAb (CD1a or CD14). NP was detected in 80–95% of MP (Fig. 1, B and C), and CD14 was detected in the membrane (Fig. 1B). DC were also susceptible to LV infection, as shown by the detection of NP in 60–90% of DC, which otherwise express CD1a (Fig. 1, E and F). NP was also demonstrated to be present in DC and MP by flow cytometry. Intracellular NP production was detected as early as 1 day after infection, reaching a peak at day 3 in both DC and MP (Fig. 2, A and B), and declining thereafter (data not shown).
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Viral particles were titrated in supernatants of DC and MP infected at various MOI. LV infection is productive in both DC and MP, particularly with an MOI of 0.2 (Fig. 3A). With MOI of 2 and 10, virus production in supernatants was lower for DC, and not significant for MP, given the large number of residual viral particles found 1 h after infection. The production of viral particles was observed in supernatants of DC and MP as soon as 24 h after infection. Virus production peaked at 72 h in DC and 48 h in MP, and was still efficient at 7 days for DC (MOI = 0.2). More viral particles were produced in DC than in MP (at least 30 times more), with the level of production almost reaching that in Vero E6 cells (Fig. 3A). We assessed the susceptibility of mature DC to LV infection by activating DC with LPS 24 h before infection. LV infection of mature DC was also productive, and infection followed a similar time course to that in immature DC (data not shown). We compared viral particle production three days after infection in mature and immature DC (Fig. 3B). Mature DC produced significantly fewer viral particles than their immature counterparts (p < 0.05). Thus, mature DC are susceptible to LV infection, but to a lesser extent than immature DC.
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We investigated whether apoptosis was associated with LV infection of DC and MP by staining cells with annexin V and 7AAD various numbers of days after infection. The percentage of apoptotic cells was similar in mock- and LV-infected DC, whether or not they were activated with sCD40L 2 h after infection. (Fig. 4A) Similarly, LV infection of MP was not associated with cell death (Fig. 4B). Cell viability was similar in inactivated LV-stimulated DC and MP (data not shown). The absolute number of cells recovered in each well was determined using calibrated beads and flow cytometry, but no difference was observed between mock- and LV-infected DC and MP (data not shown). Thus, LV infection does not reduce the viability of DC and MP.
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The expression of several molecules involved in cell activation, costimulation, or Ag presentation at the surface of DC and MP was analyzed during the course of LV infection. DC expressed CD1a, whereas CD14 was detected only on the surface of MP (Fig. 5A). The expression of CD86, CD80, CD40, CD54, HLA-DR, HLA-ABC, and ILT3 did not change during the course of LV infection in DC and MP (Fig. 5B) even 7 days after infection (data not shown). Similar results were obtained if both types of cell were stimulated with inactivated LV (data not shown). Thus, LV did not activate DC or MP, and did not induce the maturation of DC, as suggested by the absence of CD83 at the surface of infected DC (Fig. 5B). The expression of these molecules was similar in LV-infected, inactivated LV-stimulated, and mock-treated cells if DC and MP were activated 2 h after infection with sCD40L and LPS, respectively, suggesting that LV infection did not interfere with these exogenous activation signals (Fig. 5B and data not shown).
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We studied the functional properties of infected DC to investigate whether DC matured in response to LV infection. As shown in Fig. 6, the infection of immature DC with LV neither modified their capacity to take up dextran-FITC (85 ± 12% dextran-positive in mock-DC and 82 ± 14% in Lassa-DC) by phagocytosis, nor changed the rate of disappearance of the phagocytic properties of these cells in response to sCD40L treatment 2 h postinfection (p.i.) (36 ± 9% dextran-positive in mock-DC and 38 ± 12% in Lassa-DC). Similar results were obtained with inactivated LV (data not shown).
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The synthesis of several pro- and anti-inflammatory cytokines and of the IL-2R (CD25) was studied at the RNA and protein levels (except for CD25). The levels of several mRNA species were assessed by real-time RT-PCR, with normalization according to -actin mRNA levels. Certain other mRNA species were detected purely by qualitative RT-PCR. Neither the infection of DC nor the stimulation of these cells with inactivated virus induced the synthesis of mRNA for TNF-, IL-1, IL-12p35, IL-10, TGF, IFN-, or CD25 (Table I). In contrast, we observed a decrease (nonsignificant in most cases) in the production of mRNAs encoding TNF- and IL-1 in response to LV. Similarly, the synthesis of mRNAs for IL-6 and IL-12p40 was not induced in response to LV infection or to stimulation with inactivated virus (data not shown). These results were confirmed by the absence of TNF-, IL-1, IL-10 (Fig. 7A), and IL-12p70 (below the detection threshold, not shown) in supernatants of infected DC. It is interesting to note that neither LV infection nor inactivated LV-stimulation of DC affected the response to sCD40L activation (2 h after infection), in terms of either mRNA (Table I) or protein (Fig. 7A) levels. Similarly, neither the infection of MP nor the stimulation of these cells with inactivated LV induced the production of mRNAs encoding TNF-, IL-1, IL-12p35 and p40, IL-10, IL-6, IFN-, TGF, or CD25 mRNA (Table II and data not shown). These results were confirmed by the lack of TNF-, IL-1, IL-10 (Fig. 7B), and IL-12p70 (undetected, not shown) proteins in the supernatants of LV-infected MP.
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To determine whether LV infection of DC lead to an induction of the expression of chemokines and their receptors, the expression of mRNA coding for MIP-1, MIP-1, IL-8, CCR5, CCR6, and CCR7 was analyzed by real-time RT-PCR. The expression of the mRNA coding for CCR6 was unchanged after infection of DC, activated or not with sCD40L, with LV or stimulation with inactivated LV. On the contrary, infection of DC with LV, and in a lesser extent with inactivated LV, led to a 2-fold drop (nonsignificant) and a 4-fold drop (p < 0.05) of the synthesis of CCR5 and CCR7 mRNA, respectively (Table III). LV infection of DC led to a 2-fold enhancement (nonsignificant) of the expression of the mRNA coding for MIP-1 and MIP-1 1 and 3 days after infection, while sCD40L-stimulated LV-infected DC expressed significantly more MIP-1 mRNA 3 days p.i than their mock-infected counterparts (Table III). Interestingly, the expression of IL-8 mRNA was significantly up-regulated (p < 0.05) following LV infection but not after stimulation with inactivated LV, and the same results have been obtained when DC are stimulated with sCD40L 2 h after infection (Table III). However, the constitutive levels of the respective chemokines in supernatants of DC were unchanged following LV infection, as observed in the Table IV.
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The capacity of LV-infected DC to migrate toward several chemokines was studied 3 days after infection. Whereas 24% of mock-infected DC migrated toward MIP-1, only 14% of LV-infected DC have kept this capacity (Fig. 8). The stimulation of DC with LPS led to the disappearance of the response to MIP-1. Mock-infected DC migrated poorly toward MIP-3 (6.5% of input cells). Interestingly, infection of DC with LV did not lead to an enhancement (5.1%) in the number of cells that have migrated toward MIP-3. Activation of DC with LPS induced a strong migration in response to MIP-3 stimulation (52 and 48% for mock- and LV-infected DC, respectively), confirming that LV infection do not interfere with exogenous activation signals. Finally, migration toward MIP-3 was negligible in all conditions tested (Fig. 8). To determine whether LV infection of DC induce the production of chemokines able to attract uninfected DC, the capacity of immature DC to migrate toward LV-infected DC culture supernatants was analyzed. However, no significant migration of immature DC has been observed in response to mock- and LV-infected DC culture supernatants (migration of immature DC toward day 3 mock-infected DC supernatant, 7.1%; toward day 3 LV-infected DC supernatant, 5.1%).
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Discussion |
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LV infection of DC and MP did not affect cell viability. This may be due to binding of the Z protein of LV with promyelocyte leukemia protein, an oncoprotein induced by and involved in type I IFN responses (28, 29). This binding leads to the redistribution of promyelocyte leukemia protein to the cytoplasm (30) and inhibition of its pro-apoptotic activity (31). Thus, the deregulation of host cell apoptotic machinery may allow Arenaviruses to establish persistent infections.
DC and MP were not activated by LV infection. Indeed, the expression of several surface molecules involved in the costimulation of T cells, adhesion, and Ag presentation was not modified during the course of LV infection, even at day 7 (data not shown). Similarly, the lack of synthesis of proinflammatory cytokines by infected DC and MP, observed at both the mRNA and protein levels, confirmed that these cells were not activated. The absence of TNF- production in LV-infected MP and DC has been reported in previous studies (9, 25). In this study, we observed that none of the proinflammatory cytokines normally induced in virus-infected DC and MP (i.e., TNF-, IL-1, IL-6, and IL-12 (32)) was up-regulated. On the contrary, LV infection may even have inhibited the constitutive synthesis of these cytokines, given the slightly, but not significantly lower levels of mRNA and protein for TNF- and IL-1 in LV-infected DC. This finding was unexpected, as most RNA viruses activate DC and MP due to the production of dsRNA, an intermediate product of viral replication that appears to trigger many of the cellular responses to virus infection (33). The recognition of dsRNA is mediated by Toll-like receptor 3, the IFN-inducible protein kinase, and 2',5' oligoadenylate synthetase, and results in NF-
B activation (33, 34). The Z protein of LV may be responsible for the lack of cell activation as this protein has been reported to interact with certain host cell translation factors in the LCMV system (35). The lack of proinflammatory cytokine production by LV-infected DC and MP suggests that these cells may not directly be involved in the increase in permeability of the endothelium observed in LV infection, in contrast to previous suggestions (36). Nevertheless, we cannot exclude the possibility that factors other than those analyzed here are produced by infected APC.
LV infection of DC is associated with a moderate induction of the expression of the mRNA coding for the chemokines IL-8, MIP-1, and MIP-1, as previously described (25). However, the expression of these mRNA does not lead to a significant release of chemokines in our hands, as we did not observe difference in the levels of the three chemokines in supernatants of mock- and LV-infected DC. Furthermore, no migration of immature DC was detected toward supernatants of LV-infected DC, confirming that MIP-1 and MIP-1 were not significantly released after LV-infection. The discrepancy between our results and the significant levels of IL-8, MIP-1, and MIP-1 observed by Mahanty et al. (25) may be related to the different strains of LV used in both studies, but more probably to the different methods used to prepare the viral stocks. Rather than obtaining LV after 14 days of culture on Vero E6 and disruption of the cell-monolayer, we harvested the LV-containing Vero E6 cell supernatant after 3 days of culture. Viral titers are maximal at this time point in Vero E6 cells. Thus, the viral preparation of Mahanty et al. (25) probably contained a significant quantity of noninfectious viral products in addition to infectious particles. Indeed, we found with the AV strain of LV that the ratio of viral RNA copies to infectious viral particles was 100 times higher 14 days after infection of Vero cells than 3 days after infection (unpublished results). These noninfectious viral products may lead to the LV replication-independent release of chemokines by DC, consistent with the finding that chemokine release was induced by both infectious and inactivated LV (25). LV-infected DC do not express CCR6 neither acquire the capacity to migrate toward MIP-3, a chemokine produced in inflamed tissues. This lack of expression is probably linked to the lack of TGF production by LV-infected DC, as monocyte-derived DC express CCR6 only when TGF is present in the culture medium (37).
DC do not mature in response to LV infection, as shown by the lack of CD83 expression and of changes in the ability of these cells to take up dextran-FITC. This lack of maturation was confirmed by the lack of induction of allogeneic lymphocyte proliferation by LV-infected DC (25). Furthermore, LV-infection of immature DC did not induce the capacity to migrate toward MIP-3, a chemokine involved in the chemotaxis of mature DC in secondary lymphoid organs (38), and CCR-7 mRNA levels were significantly decreased by the LV infection of DC. Together with the absence of maturation in LV-infected DC, these results suggest that an immature DC infected in the periphery is unlikely to migrate to the secondary lymphoid organs and induce T cell responses (32, 39), and that splenic immature DC that seem to represent the main population of splenic DC (40), will be massively infected without maturation. This hypothesis is supported by the findings that Ag presentation by immature DC results in tolerance and the induction of regulatory T cells (41, 42), and that proinflammatory cytokines are crucial to the induction of adaptive immunity (43). Similarly, the lack of activation of MP following LV infection may be involved in the pathogenicity of the disease, favoring viral spread, as the activation of MP is known to increase the microbicidal action of these phagocytes (44). This is consistent with the finding that infection of the guinea pig with a virulent strain of Pichinde virus, a new-world Arenavirus, leads to a lack of MP activation, whereas the attenuated counterpart of this virus does induce MP activation (45). Furthermore, the infection of MP with Mopeia virus, a nonpathogenic Arenavirus closely related to LV (46), induces the activation of cells (D. Pannetier, manuscript in preparation). Thus, the absence of activation and maturation in LV-infected DC and MP may be associated with viral pathogenicity and with the absence of effective inflammatory and immune responses observed in fatal LV infection (14, 16).
Interestingly, LV does not modify the responses of DC and MP to exogenous activation signals such as sCD40L and LPS. This was demonstrated by the lack of change in the modifications induced in cells by these stimuli: up-regulation of surface molecule expression and of cytokine production, and acquisition of the mature phenotype by DC including the capacity to migrate toward MIP-3. LV infection only slightly inhibits activation as this virus is able to prevent cellular responses by its own products without changing other activation pathways. Most viruses known to inhibit APC activation also affect responses to other stimuli (47, 48). In another report, LV was reported to inhibit the production of TNF- and IL-8 induced by LPS in MP (9). The reasons for these differences are unclear, but may be related to differences in the experimental conditions used in the two studies. The dose of LPS and the time after LV infection at which LPS was added to cultures differed, as well as the LV strains used. These results suggest that LV probably cannot inhibit the activation of APC by other pathogens and that mature LV-infected DC residing in secondary lymphoid organs may be able to interact with resident T cells via CD40-CD40L signals and to induce effective T cell responses.
We found that immature DC produced significantly more LV particles than MP and mature DC. We cannot yet account for immature DC being the most permissive cells for LV, but investigations are underway. This observation suggests that immature DC may be a better reservoir than MP and mature DC for LV production in vivo.
In conclusion, we have demonstrated that DC and MP are susceptible to a human isolate of LV and that these cells probably play an important role in the early burst of LV replication from the initial site of infection. LV infection did not lead to activation of DC and MP, but the virus did not interfere with exogenous activation signals or with cell viability. Thus, the tropism of LV for APC is probably crucial to the pathogenicity and immunosuppression associated with this severe disease.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Sylvain Baize, Unité de Biologie des Infections Virales Emergentes, Centre de Recherche Mérieux-Pasteur à Lyon, 21 Avenue Tony Garnier, 69365 Lyon Cedex 07 France. E-mail address: baize{at}cervi-lyon.inserm.fr
3 Abbreviations used in this paper: LV, Lassa virus; Z, zinc-binding; L, RNA polymerase; NP, nucleoprotein; DC, dendritic cell; MP, macrophage; LCMV, lymphocytic choriomeningitis virus; FFU, focus-forming unit; rh, recombinant human; PFA, paraformaldehyde; RT, room temperature; sCD40L, soluble rh CD40 ligand; CT, cycle threshold, MOI, multiplicity of infection; 7AAD, 7-aminoactinomycin D; MIP, macrophage inflammatory protein; GAM, goat anti-mouse; p.i., postinfection.
Received for publication March 31, 2003. Accepted for publication December 17, 2003.
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References |
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  B. Hemati, V. Contreras, C. Urien, M. Bonneau, H.-H. Takamatsu, P. P. C. Mertens, E. Breard, C. Sailleau, S. Zientara, and I. Schwartz-Cornil Bluetongue Virus Targets Conventional Dendritic Cells in Skin Lymph J. Virol., September 1, 2009; 83(17): 8789 - 8799. [Abstract] [Full Text] [PDF]
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S. Baize, P. Marianneau, P. Loth, S. Reynard, A. Journeaux, M. Chevallier, N. Tordo, V. Deubel, and H. Contamin Early and Strong Immune Responses Are Associated with Control of Viral Replication and Recovery in Lassa Virus-Infected Cynomolgus Monkeys J. Virol., June 1, 2009; 83(11): 5890 - 5903. [Abstract] [Full Text] [PDF]
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S. Arimilli, J. B. Johnson, K. M. Clark, A. H. Graff, M. A. Alexander-Miller, S. B. Mizel, and G. D. Parks Engineered Expression of the TLR5 Ligand Flagellin Enhances Paramyxovirus Activation of Human Dendritic Cell Function J. Virol., November 15, 2008; 82(22): 10975 - 10985. [Abstract] [Full Text] [PDF]
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 E. P. Scott and J. F. Aronson Cytokine patterns in a comparative model of arenavirus haemorrhagic fever in guinea pigs J. Gen. Virol., October 1, 2008; 89(10): 2569 - 2579. [Abstract] [Full Text] [PDF]
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A. O. Weinzierl, G. Szalay, H. Wolburg, M. Sauter, H.-G. Rammensee, R. Kandolf, S. Stevanovic, and K. Klingel Effective Chemokine Secretion by Dendritic Cells and Expansion of Cross-Presenting CD4-/CD8+ Dendritic Cells Define a Protective Phenotype in the Mouse Model of Coxsackievirus Myocarditis J. Virol., August 15, 2008; 82(16): 8149 - 8160. [Abstract] [Full Text] [PDF]
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R. Carrion Jr., K. Brasky, K. Mansfield, C. Johnson, M. Gonzales, A. Ticer, I. Lukashevich, S. Tardif, and J. Patterson Lassa Virus Infection in Experimentally Infected Marmosets: Liver Pathology and Immunophenotypic Alterations in Target Tissues J. Virol., June 15, 2007; 81(12): 6482 - 6490. [Abstract] [Full Text] [PDF]
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S. Muller, R. Geffers, and S. Gunther Analysis of gene expression in Lassa virus-infected HuH-7 cells J. Gen. Virol., May 1, 2007; 88(5): 1568 - 1575. [Abstract] [Full Text] [PDF]
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S. M. Fennewald, E. P. Scott, L. Zhang, X. Yang, J. F. Aronson, D. G. Gorenstein, B. A. Luxon, R. E. Shope, D. W. C. Beasley, A. D. T. Barrett, et al. Thioaptamer decoy targeting of AP-1 proteins influences cytokine expression and the outcome of arenavirus infections J. Gen. Virol., March 1, 2007; 88(3): 981 - 990. [Abstract] [Full Text] [PDF]
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G. C. Bowick, S. M. Fennewald, E. P. Scott, L. Zhang, B. L. Elsom, J. F. Aronson, H. M. Spratt, B. A. Luxon, D. G. Gorenstein, and N. K. Herzog Identification of Differentially Activated Cell-Signaling Networks Associated with Pichinde Virus Pathogenesis by Using Systems Kinomics J. Virol., February 15, 2007; 81(4): 1923 - 1933. [Abstract] [Full Text] [PDF]
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A. Bergthaler, D. Merkler, E. Horvath, L. Bestmann, and D. D. Pinschewer Contributions of the lymphocytic choriomeningitis virus glycoprotein and polymerase to strain-specific differences in murine liver pathogenicity J. Gen. Virol., February 1, 2007; 88(2): 592 - 603. [Abstract] [Full Text] [PDF]
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G. C. Bowick, S. M. Fennewald, B. L. Elsom, J. F. Aronson, B. A. Luxon, D. G. Gorenstein, and N. K. Herzog Differential signaling networks induced by mild and lethal hemorrhagic Fever virus infections. J. Virol., October 1, 2006; 80(20): 10248 - 10252. [Abstract] [Full Text] [PDF]
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L. Martinez-Sobrido, E. I. Zuniga, D. Rosario, A. Garcia-Sastre, and J. C. de la Torre Inhibition of the type I interferon response by the nucleoprotein of the prototypic arenavirus lymphocytic choriomeningitis virus. J. Virol., September 1, 2006; 80(18): 9192 - 9199. [Abstract] [Full Text] [PDF]
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S. Arimilli, M. A. Alexander-Miller, and G. D. Parks A Simian Virus 5 (SV5) P/V Mutant Is Less Cytopathic than Wild-Type SV5 in Human Dendritic Cells and Is a More Effective Activator of Dendritic Cell Maturation and Function. J. Virol., April 1, 2006; 80(7): 3416 - 3427. [Abstract] [Full Text] [PDF]
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 P. M. A. de Graaff, E. C. de Jong, T. M. van Capel, M. E. A. van Dijk, P. J. M. Roholl, J. Boes, W. Luytjes, J. L. L. Kimpen, and G. M. van Bleek Respiratory Syncytial Virus Infection of Monocyte-Derived Dendritic Cells Decreases Their Capacity to Activate CD4 T Cells J. Immunol., November 1, 2005; 175(9): 5904 - 5911. [Abstract] [Full Text] [PDF]
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 A. Boesen, K. Sundar, and R. Coico Lassa Fever Virus Peptides Predicted by Computational Analysis Induce Epitope-Specific Cytotoxic-T-Lymphocyte Responses in HLA-A2.1 Transgenic Mice Clin. Vaccine Immunol., October 1, 2005; 12(10): 1223 - 1230. [Abstract] [Full Text] [PDF]
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S. Kunz, J. M. Rojek, M. Perez, C. F. Spiropoulou, and M. B. A. Oldstone Characterization of the Interaction of Lassa Fever Virus with Its Cellular Receptor {alpha}-Dystroglycan J. Virol., May 15, 2005; 79(10): 5979 - 5987. [Abstract] [Full Text] [PDF]
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D. Pannetier, C. Faure, M.-C. Georges-Courbot, V. Deubel, and S. Baize Human Macrophages, but Not Dendritic Cells, Are Activated and Produce Alpha/Beta Interferons in Response to Mopeia Virus Infection J. Virol., October 1, 2004; 78(19): 10516 - 10524. [Abstract] [Full Text] [PDF]
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