Interaction of classical swine fever virus with dendritic cells

C. P. Carrasco1, R. C. Rigden1, I. E. Vincent1, C. Balmelli1, M. Ceppi1, O. Bauhofer1, V. Tâche1, B. Hjertner2, F. McNeilly2, H. G. van Gennip3, K. C. McCullough1 and A. Summerfield1

1 Institute of Virology and Immunoprophylaxis, Sensemattstrasse 293, CH-3147 Mittelhäusern, Switzerland
2 Department of Agriculture for Northern Ireland, Veterinary Sciences Division, Belfast, UK
3 Animal Sciences Group, 8200 AB Lelystad, The Netherlands

Correspondence
A. Summerfield
artur.summerfield{at}ivi.admin.ch


   ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Functional disruption of dendritic cells (DCs) is an important strategy for viral pathogens to evade host defences. Monocytotropic viruses such as classical swine fever virus (CSFV) could employ such a mechanism, since the virus can suppress immune responses and induce apoptosis without infecting lymphocytes. Here, CSFV was shown to infect and efficiently replicate in monocyte- and in bone marrow-derived DCs. Interestingly, the infected DCs displayed neither modulated MHC nor CD80/86 expression. Stimulation of DCs with IFN-{alpha}/TNF-{alpha} or polyinosinic–polycytidylic acid (pIC) induced phenotypic maturation with increased MHC and CD80/86 expression, both with mock-treated and infected DCs. In addition, the T cell stimulatory capacity of CSFV-infected DCs was maintained both in a polyclonal T cell stimulation and in specific antigen-presentation assays, requiring antigen uptake and processing. Interestingly, similar to macrophages, CSFV did not induce IFN-{alpha} responses in these DCs and even suppressed pIC-induced IFN-{alpha} induction. Other cytokines including interleukin (IL)-6, IL-10, IL-12 and TNF-{alpha} were not modulated. Taken together, these results demonstrated that CSFV can replicate in DCs and control IFN type I responses, without interfering with the immune reactivity. These results are interesting considering that DC infection with RNA viruses usually results in DC activation.


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Classical swine fever (CSF) is a highly contagious disease of pigs caused by CSF virus (CSFV) and leads to important economic losses worldwide. CSFV together with bovine viral diarrhoea virus (BVDV) and border disease virus (BDV) form the genus Pestivirus within the family Flaviviridae.

CSFV is a monocytotropic viral pathogen, which can efficiently evade and compromise the host's immune system. The virus has a high affinity for reticulo-endothelial cells (Cheville & Mengeling, 1969; Ressang, 1973; Susa et al., 1992) causing lymphopenia, thrombocytopenia, coagulation disorders and atrophy of the thymus and bone marrow (Gomez-Villamandos et al., 2003; Pauly et al., 1998; Sanchez-Cordon et al., 2002; Summerfield et al., 2000, 2001). Lymphopenia is caused, at least in part, by apoptosis detectable in uninfected lymphocytes (Sanchez-Cordon et al., 2003; Summerfield et al., 1998b). In addition, viable lymphocytes isolated from CSFV-infected pigs do not respond to mitogen stimulation (Pauly et al., 1998; Summerfield et al., 1998b; Van Oirschot et al., 1983). These modulated cells are not infected. Instead, it is the myeloid population, particularly monocytes (Mo) and macrophages (M{phi}), that contains the early target cell for infection and replication, both in vivo (Ressang, 1973; Gomez-Villamandos et al., 2001; Sanchez-Cordon et al., 2003; Summerfield et al., 2000; Trautwein, 1988) and in vitro (Knoetig et al., 1999). Despite this clear targeting and tropism, no direct evidence has been found of a role for infected Mo and M{phi} in the observed immunosuppression and death of T lymphocytes (Knoetig et al., 1999).

Dendritic cells (DCs) are one of the primary immunological sentinels of the immune system (Banchereau et al., 2000; Steinman, 1991). They can efficiently sense invading pathogens by a set of pattern recognition receptors. Their strategic localization at the site of pathogen entry – mucosal surfaces and dermal layers – makes them a particularly early target for virus contact (MacPherson & Liu, 1999). After detection, uptake and degradative processing of the virus, DCs should mature and subsequently promote effective immune responses by migrating into lymphoid tissue to present the processed viral antigens to T lymphocytes (Pulendran et al., 2001).

Interestingly, certain viruses are tropic for DCs. Human immunodeficiency virus, measles virus (MV), cytomegalovirus and herpes simplex virus type 1 infect, survive and even replicate in these potent antigen-presenting cells, inducing immunosuppression, mediated in part through the infected DCs (Klagge & Schneider-Schaulies, 1999). A problem for the virus could arise if the infection results in activation of receptors for double-stranded (ds)RNA such as toll-like receptor 3 and protein kinase R, which would activate and mature the DCs. Such an observation has been made particularly with certain DC-tropic RNA viruses including MV (Fugier-Vivier et al., 1997), influenza virus (Cella et al., 1999), Semliki Forest virus (Johnston et al., 1996), dengue virus (Ho et al., 2001) and Sindbis virus adapted to human DCs (Gardner et al., 2000). As a consequence, type I IFN production is induced and the virus-infected DCs mature to acquire the capacity to induce potent MHC class I- and II-restricted antiviral immune responses. In the context of CSFV, the results with dengue virus, also a member of the Flaviviridae, are interesting, yet nothing is known about the interaction of CSFV with DCs.

Considering these fundamental elements in virus–host interactions, particularly of monocytotropic viruses, the objective of the present study was to determine the affinity of CSFV for DCs and the functional consequences of that infection.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Media and reagents.
DCs were cultured in Dulbecco's modified Eagle's medium (DMEM; Invitrogen) supplemented with 2 mM glutamine, 100 U penicillin ml–1, 100 µg streptomycin ml–1 and 50 µM 2-ME (all from Invitrogen). Serum supplements for DC cultures employed 10 % (v/v) porcine serum (Sigma Chemicals). Recombinant porcine (rp) cytokines utilized for DC generation were as follows: rpGM-CSF (kindly provided by Dr S. Inumaru, Institute for Animal Health, Ibaraki, Japan; Inumaru et al., 1998) and rpTNF-{alpha} (kindly provided by Dr G. Bertoni, Institute of Veterinary Virology, Berne, Switzerland; Von Niederhausern et al., 1993). Recombinant porcine IL-4 was prepared in our laboratory, as described previously (Carrasco et al., 2001). MHC class I was detected with mAb 74-11-10 (Pescovitz et al., 1984) (kindly donated by Dr J. K. Lunney, USDA, Beltsville, USA) and MHC class II with mAb MSA3 (Hammerberg & Schurig, 1986) (from VMRD, Pullman, WA, USA). CD80/86 expression was measured using a huCTLA4–mouse Ig fusion protein (Alexis, Lausen, Switzerland).

Generation of porcine DCs.
Bone marrow (BM)-derived and blood Mo-derived DCs were generated as described previously (Carrasco et al., 2001). Briefly, low-density bone marrow haematopoietic cells (BMHCs) and PBMCs were obtained by density-gradient centrifugation over Ficoll-Paque (1·077 g l–1; Amersham Pharmacia Biotech). BM-DCs were generated from BMHCs using rpGM-CSF (25 ng ml–1) and rpTNF-{alpha} (30 U ml–1), cultured for 8 days at 39 °C. At days 3 and 6, the BM-DC culture was fed with the cytokines. Mo-DCs were generated from blood Mo isolated by plastic adherence, using rpGM-CSF (150 ng ml–1) and rpIL-4 (100 U ml–1), cultured for 7 days at 39 °C. At days 2, 4 and 6, the Mo-DC culture was fed with the cytokines. Maturation of DCs was induced by treatment with a cocktail of recombinant porcine IFN-{alpha} (1000 U ml–1; R&D Systems) and TNF-{alpha} (10 ng ml–1; Endogen) for 48 h. Alternatively, maturation was induced with polyinosinic–polycytidylic acid (pIC; Sigma) transfection to mimic a virus infection. DCs were washed twice with serum-free DMEM at 37 °C and resuspended at a concentration of 4x106 cells ml–1 in 500 µl Opti-MEM (Invitrogen). pIC was first incubated for 30 min with 2 µl of a single-stranded RNA-specific RNase cocktail composed of 500 U RNase A ml–1 and 20 000 U RNase T1 ml–1 (both from Ambion) in 100 µl at 37 °C. This digestion improved the stimulatory activity of pIC for DCs, whereas treatment with dsRNA-specific RNase abolished the capacity of pIC to induce DC maturation (M. Ceppi, unpublished results). This mixture was allowed to form complexes with 24 µl TransFast (Promega) in 500 µl Opti-MEM for 15 min at room temperature, before adding to the DCs and incubating for 1 h at 39 °C. After lipofection, cells were washed twice with serum-free DMEM at 37 °C before use in T cell stimulation assays.

CSFV infection.
The virulent Brescia strain (Summerfield et al., 1998b) was used if not otherwise indicated. For certain other experiments, the virulent Eystrup strain (Mayer et al., 2003), the moderately virulent Alfort/187 strain (Ruggli et al., 1996) and the avirulent C-strain (vflc2 corresponding to Flc 133 described by Moormann et al., 1996) were employed. These viruses and mock controls were prepared in swine kidney SK-6 cells as described (Knoetig et al., 1999). In some experiments, UV-inactivated virus and preparations (Knoetig et al., 1999) were used as additional controls. DCs (BMHCs or Mo) were mock-treated or infected with CSFV at the m.o.i.s specified in the results. The presence of viral antigen in the DCs was quantified by immunofluorescent analysis using microscopic or flow cytometric detection of the viral E2 structural glycoprotein with mAb HC/TC26 (kindly provided by Dr Bommeli, Diagnostics AG, Berne, Switzerland; Greiser-Wilke et al., 1990). The viral non-structural NS3 protein was detected with mAb C16 (kindly provided by Dr I. Greiser-Wilke, Hannover Veterinary School, Germany; Greiser-Wilke et al., 1992). Due to the internal expression of E2 and NS3, the DCs were fixed and permeabilized (Cell Permeabilization Kit; Harlan Sera-Lab) before labelling with the mAbs.

Analysis of CSFV replication in BM- and Mo-DCs was assessed by incubating the non-adherent DCs with CSFV at an m.o.i. of 10 TCID50 per cell for 1 h at 39 °C. The cells were then washed a total of ten times by centrifugation. Cell-associated virus (CAV) from these DCs was obtained by two cycles of freeze–thawing of the cell pellets, which were then clarified by centrifugation at 10 000 g for 20 min and resuspended to the original culture volume. The measurement of virus titres from extracellular virus (ECV) and CAV were quantified by end-point titration on PK-15 cells and immunofluorescent detection.

DC-dependent T lymphocyte stimulation assays.
T cell purification for all assays (SEB-, FMDV- and CSFV-dependent) used MACS (magnetic activated cell sorting), wherein either CD6+ T cell enrichment or SWC3+ Mo depletion was used. T cells were cultured at 2x105 cells per well. The microbial superantigen staphylococcal enterotoxin B (SEB) (Toxin Technology) was used to measure the potency of DCs at inducing T cell responses dependent on MHC class II–T cell receptor cross-linking (Bhardwaj et al., 1992). BM-DCs were incubated simultaneously with mitomycin C (10 µg ml–1) and SEB (100 ng ml–1) for 1 h at 39 °C, washed four times and titrated in a 96-well flat bottom microtitre plate as described. After 2 days, 1 µCi (37 kBq) [3H]thymidine was added for another 18 h to quantify proliferation.

The processing and presentation of antigen was assessed in two antigen-specific assays. For the CSFV-specific antigen-presentation assays, PBMCs for both Mo-DC generation and lymphocyte isolation were obtained from pigs 4–8 months after infection with CSFV Alfort187 (Ruggli et al., 1996). CSFV was added to the DC/lymphocyte co-cultures at an m.o.i. of 0·01 TCID50 per cell. The foot-and-mouth disease virus (FMDV)-specific restimulation assay employed T cells and Mo-DCs isolated from pigs vaccinated against FMDV (three booster injections, serotype C1 Oberbayern; vaccine kindly provided by Dr P. Barnett, Institute for Animal Health, Pirbright, UK). The DCs and T cells were co-cultured in the presence of {beta}-propiolactone-inactivated FMDV antigen as described previously (Carrasco et al., 2001). After 4 days for the FMDV and 5 days for the CSFV restimulation, 1 µCi [3H]thymidine was added for another 18 h to quantify proliferation. T cell activation was only observed with immune animals.

Cell viability and apoptosis analysis.
For quantification of apoptotic cells expressing phosphatidylserine on their surface and dead cells permeable to propidium iodide (PI; Sigma), dual parameter analysis of AnnexinV-FITC (Bender Med Systems) and PI were performed (Vermes et al., 1995). To this end, 5x105 cells were labelled with 2 µg AnnexinV-FITC ml–1 in 140 mM NaCl, 2·5 mM CaCl2, 10 mM HEPES (pH 7·4) buffer for 10 min. After FL1/FL2 compensation, PI (100 ng ml–1) was added, to discriminate between apoptotic and dead cells and the sample analysed by flow cytometry.

Cytokine responses.
Cytokines produced by DCs were induced with either pIC or lipopolysaccharide (LPS, Escherichia coli-derived; Sigma). The concentration of secreted IFN type I in the supernatants was quantified using a bioassay based on the antiviral effect of IFN type I against vesicular stomatitis virus (VSV). Reduction of VSV-induced cytopathic effect in PK-15 cells induced by the test samples or by recombinant porcine IFN-{alpha} (R&D Systems) as a standard was quantified as described previously (Ruggli et al., 2003). The presence of IFN-{alpha} was controlled by addition of a neutralizing anti-porcine IFN-{alpha} polyclonal antiserum (R&D Systems).

Other cytokines produced by DCs were measured using commercially available ELISAs designed for porcine IL-6, IL-10, IFN-{gamma} (Biosource) and TNF-{alpha} (Perbio Science).

For quantification of IL-10 and IL-12 mRNA levels, RNA was isolated from DCs using the RNeasy Kit (Qiagen). This RNA was reverse-transcribed to cDNA using the Omniscript RT Kit (Qiagen). The cDNA levels of cyclophilin (housekeeping gene), IL-10 and IL-12 (subunits p35 and p40) were quantified on the Icycler IQ Multicolour Real-Time PCR Detection System (Bio-Rad) using the Lightcycler Faststart DNA Master SYBR Green kit (Roche Molecular Diagnostics). Duplicate samples as well as serial dilutions of each target amplicon were analysed using the following optimized protocol. A 10 min incubation at 95 °C was employed to activate the polymerase and denature the targets. This was followed by 50 cycles of 30 s at 95 °C, 30 s at 55 °C (cyclophilin), 59 °C (IL-10, IL-12p40) or 61 °C (IL-12p35), and 30 s at 72 °C. After a final extension for 10 min at 72 °C, the programme was ended with a melt curve programme. All reactions were performed in 20 µl (1 µl sample added), the MgCl2 concentration was 3 mM in all cases and the primer concentrations were 0·25 µM. The following forward and reverse primers, respectively, were used for PCR: cyclophilin: 5'-TAACCCCACCGTCTTCTT-3' and 5'-TGCCATCCAACCACTCAG-3'; IL-10: 5'-GCATCCACTTCCCAACCA-3' and 5'-CTTCCTCATCTTCATCGTCAT-3'; IL-12p35: 5'-GCCTCAACCACTCCCAAAATC-3' and 5'-CTTCCAGGGAGGGCTTCTGT-3'; and IL-12p40: 5'-GCAAAACCCTGACCATCCAC-3' and 5'-TGCCCTCCTGACACTCCACT-3'. Serial dilutions were used to generate standard curves from which the concentration of each sample was deduced. All cytokine data were normalized to the expression of cyclophilin.


   RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
CSFV infection of DCs
It is known that CSFV has an affinity for myeloid cells (Knoetig et al., 1999; Ressang, 1973; Summerfield et al., 1998a, b, 2000, 2001). However, there is no information on the susceptibility of DCs to CSFV infection. Considering the central role that DCs play in the stimulation and control of immune defence development, this is a subject of particular importance. As a cell culture model, Mo- and BM-DCs, both representing myeloid DCs, were used. Similar to other species, these cells are characterized as DCs based on their dendritic morphology, non-adherence, their potent T cell stimulatory capacity and their expression of CD1, CD80/86 and MHC class II (Carrasco et al., 2001). Consequently, BM- or Mo-DCs were infected with CSFV at an m.o.i. of 1 TCID50 per cell. At 48 h post-infection, the majority of the infected BM- and Mo-DCs were seen to express viral E2 and NS3 proteins (Fig. 1). The detection method for these viral proteins did not permit the detection of virus entry, but only de novo protein synthesis (Knoetig et al., 1999). These results suggested a productive infection of the DCs, which was confirmed by the observation of a 3–4 log10-fold increase in the virus titres of ECV and CAV between 24 and 72 h post-infection (Fig. 2). Interestingly, the CAV titres were invariably lower than the ECV titres, contrasting with infected SK-6 cells, where the majority of de novo-synthesized virus was cell-associated (data not shown). Similar elevated ECV titres compared with CAV titres have been observed with CSFV-infected Mo (S. M. Knoetig and A. Summerfield, unpublished data).



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Fig. 1. Viral protein expression in BM-DCs and Mo-DCs at 48 h after CSFV infection at an m.o.i. of 1 TCID50 per cell. (a) Structural glycoprotein E2 expression. (b) Non-structural protein NS3 expression. The filled histograms represent the staining of mock-treated cells used to determine the non-specific staining. Data shown in (a) and (b) are representative of 11 independent experiments.

 


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Fig. 2. Replication kinetics of CSFV in BM-DCs (a) and Mo-DCs (b), infected at an m.o.i. of 10 TCID50 per cell for 1 h at 39 °C. At the indicated time points, the culture supernatant medium was harvested for ECV and the cells for CAV, as described in Methods. Data shown in (a) and (b) are representative of three independent experiments.

 
DC morphology and maturation after CSFV infection
No morphological difference between CSFV-infected and mock-treated DCs was noted (data not shown). This correlated with the observation that CSFV is non-cytopathic for Mo and M{phi} (Knoetig et al., 1999). In addition, CSFV infection of DCs had no apparent influence on their ‘maturation status' with respect to the expression of MHC class I, II and CD80/86 (Fig. 3). CD80/86 was already expressed without addition of maturation signals, but could be induced to be further upregulated by in vitro stimulation. This is characteristic of porcine Mo-DCs (Carrasco et al., 2001) and also porcine blood DCs (Summerfield et al., 2003). Consequently, it was investigated whether CSFV-infected DCs were altered in their response to maturation signals. When CSFV-infected or mock-treated DCs were stimulated either with a TNF-{alpha}/IFN-{alpha} cocktail or with pIC, upregulation of molecules involved in antigen presentation, typical of DC maturation, was observed in both mock-treated and CSFV-infected DCs (Fig. 3 shows the results for the IFN-{alpha}/TNF-{alpha}-induced maturation). The observation that CSFV did not influence the expression of maturation markers such as CD80/86 was also observed with other strains of CSFV, including the avirulent C-strain, the moderately virulent Alfort strain and the virulent Eystrup and Brescia strains. In addition, no modulation was observed using UV-inactivated virus as an additional control (Fig. 3b).



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Fig. 3. CD80/86, MHC class I and MHC class II expression on Mo-DCs after infection with CSFV. (a) The cells were mock-treated (light-line histograms) or infected with CSFV (dark-line histograms) at an m.o.i. of 1 TCID50 per cell and cultured for 48 h. DC maturation was induced with IFN-{alpha}/TNF-{alpha} for another 48 h in some cultures as indicated. The dotted-line histograms represent the conjugate controls. For each histogram plot, the geometric mean fluorescence intensity (MFI) for the mock controls and the virus-infected DCs is indicated. The data shown are representative of five independent experiments. (b) Influence of various CSFV strains and UV-inactivated virus on CD80/86 expression. The geometric means determined as described in (a) for unstimulated DCs (white bars) and for IFN-{alpha}/TNF-{alpha}-stimulated DCs (grey bars) are shown. Experimental conditions were as described in (a).

 
Functional activity of DCs infected with CSFV
Having observed efficient CSFV replication in DCs without detectable alteration in cell morphology or consistent modulation of MHC and CD80/86 expression, the influence of infection on the functional activity of the DCs was assessed. To this end, DCs were pulsed either with the microbial superantigen SEB or with viral antigens, and their capacity to stimulate proliferative T cell responses was quantified. Although CSFV-infected DCs were seen to be more stimulatory for SEB-induced T cell proliferation compared with mock-treated DCs (Fig. 4), the difference was not always reproducible. A statistical analysis (Student's t-test) of 11 SEB-dependent proliferation assays demonstrated no significant differences between mock-treated and CSFV-infected DCs in this functional test (P=0·2).



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Fig. 4. Antigen-presentation capacity of CSFV-infected DCs, influenced by maturation signals. Mo-DCs were mock-treated ({blacktriangleup}, {triangleup}) or CSFV-infected (m.o.i. of 1 TCID50 per cell; {bullet}, {circ}) for 48 h. Some cultures were further stimulated (filled symbols) with either IFN-{alpha}/TNF-{alpha} for 48 h (a) or pIC (transfection) for 24 h (b). The capacity of the Mo-DCs to present cell-bound SEB to T lymphocytes and to induce T cell proliferation at different DC : lymphocyte ratios is shown. Data shown are representative of 11 experiments, displaying the mean±SD of triplicate values.

 
When the same experiments were performed with IFN-{alpha}/TNF-{alpha}-stimulated infected or mock-treated DCs, it was often observed that the CSFV-infected cells promoted a higher lymphoproliferation compared with the mock controls (Fig. 4a). Nevertheless, this observation was also not seen in all experiments and a statistical analysis of the results indicated no significant difference (P=0·23). When the DCs were stimulated with lipofected pIC as the maturation signal, both the mock-treated and the CSFV-infected DCs upregulated their capacity to stimulate T cell proliferation to a similar degree (Fig. 4b).

To investigate further any possible functional modulation of CSFV-infected DCs, the influence of virus infection on an MHC-restricted antigen-specific T cell response, which requires antigen uptake, processing and presentation, was tested. CSFV-infected DCs were able to stimulate T cell proliferation in an FMDV-specific antigen-presentation assay and this was of a similar intensity to that obtained with mock-treated Mo-DCs (Fig. 5a). Another important observation was that IFN-{gamma} protein levels were not altered when supernatants from T lymphocytes activated by mock-treated or CSFV-infected DCs presenting FMDV antigen were analysed (Fig. 5b). This indicated that CSFV did not shift the T cell response towards a Th1 or a Th2 pattern. Finally, CSFV-infected DCs could also restimulate CSFV-specific T cell proliferation (Fig. 5c). Taken together, these results demonstrated that CSFV-infected DCs were certainly functionally intact with respect to their capacity to induce T cell activation.



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Fig. 5. Capacity of CSFV-infected DCs to process and present antigen. Mo-DCs were exposed to mock ({blacktriangleup}, {triangleup}) or CSFV (m.o.i. of 1 TCID50 per cell; {bullet}, {circ}) for 48 h. For (a) and (b), these DCs were then incubated with FMDV antigen (filled symbols) or mock BHK-cell antigen (open symbols) for another 24 h, before co-culturing with autologous T lymphocytes from an FMDV-immune animal. The T cell responses were measured in a proliferation test (a) and by quantification of IFN-{gamma} protein in supernatants after 48 h co-culture (b). The data shown are representative of four (a) or two (b) independent experiments. (c) The CSFV-infected DCs ({bullet}) or mock-treated controls ({blacktriangleup}) were co-cultured with autologous T lymphocytes from a CSFV-immune animal. The antigen specificity of these responses was controlled by performing the same experiments with non-immune animals (data not shown). DCs were titrated to obtain different DC : T lymphocyte ratios and the mean±SD of triplicate c.p.m. values are displayed.

 
Cell viability following CSFV infection
In the acute phase of severe CSF, death of uninfected lymphocytes can be detected in the animals. Consequently, even though CSFV was not apparently cytopathic for DCs in terms of morphological changes, it was necessary to determine whether an augmented rate of cell death was present in the infected DC/lymphocyte co-cultures. Using lymphocyte co-cultures with DCs at a ratio of 10 : 1, viability analysis at 24, 48 and 72 h showed no significant difference in the number of AnnexinV-positive and PI-positive cells between mock controls and CSFV-infected cultures (data not shown).

Cytokine responses
In a final series of analyses, the capacity of CSFV to modulate the cytokine response of virus-infected DCs was characterized. Similar to infection of Mo and M{phi} (Knoetig et al., 1999; Ruggli et al., 2003), CSFV did not induce type I IFN responses in Mo-DCs. On the contrary, CSFV-infected DCs had a defective type I IFN response to pIC stimulation (Table 1). As for other cytokines, the protein levels of IL-6, IL-10 and TNF-{alpha} measured by ELISA, as well as the mRNA levels of IL-10 and IL-12 (p35 and p40), were not modulated by CSFV infection (Table 1). No modulation of spontaneously induced TNF-{alpha}, IL-6 or IL-10 protein was detectable. The same was found for IL-10 and IL-12 p40 mRNA levels (Table 1). IL-12 p35 mRNA was not detectable in mock- or in CSFV-infected DCs (data not shown). Furthermore, the induction of TNF-{alpha} by LPS and of IL-6 by LPS and pIC were not modified by CSFV (Table 1).


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Table 1. Cytokine responses of CSFV-infected DCs

 

   DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
CSFV is a haemorrhagic and immunosuppressive viral pathogen that has been seen in histological sections associated with dendritic-like cells (Susa et al., 1992). Non-infected T lymphocytes from animals with CSF do not respond to antigen or mitogen stimulation and rapidly undergo apoptosis in vivo and after ex vivo culture. Yet, CSFV is well documented as being non-cytopathogenic. Furthermore, in vitro infection of Mo or M{phi} with this monocytotropic virus did not induce detectable lymphocyte immunosuppression nor was apoptosis discernible (Knoetig et al., 1999). This would indicate that infected M{phi} are not directly responsible for the reported lymphocyte death and anergy in CSF (Pauly et al., 1998; Van Oirschot et al., 1983). Consequently, the present work sought to determine whether DCs, the major myeloid cell type interacting with lymphocytes, are involved in CSFV immunopathogenesis.

CSFV was seen to be highly efficient at infecting and productively replicating in DCs, both Mo-DCs and BM-DCs. These observations that CSFV is tropic for DCs indicated that these critically important promoters and controllers of immune response development would serve as a main target cell for CSFV replication. Considering the non-cytopathogenic nature of CSFV and the migratory characteristics of DCs, infected DCs would present a reservoir for the efficient dissemination of the virus. The latter aspect is particularly important due to the trafficking of DCs from peripheral sites, such as the mucosal surfaces at which CSFV would invade the host, to the secondary lymphoid tissue where close contact to lymphocytes could have detrimental consequences.

Despite this tropism of CSFV for DCs, there was no clear morphological, phenotypic or functional modulation. This contrasts with dengue virus, another flavivirus, which induced DC maturation (Ho et al., 2001; Libraty et al., 2001; Wu et al., 2000). It has been reported that DCs can be activated through dsRNA-dependent stimulation (Cella et al., 1999). Considering the fact that many RNA viruses forming dsRNA intermediates during their replicative cycle activate DCs (for example, see Introduction), it would appear that CSFV has developed mechanisms to control or limit DC activation. Both the non-cytopathogenic and the cytopathogenic biotypes of another pestivirus, BVDV, do not activate DCs (Glew et al., 2003), suggesting that such common means for evading immune recognition may be a characteristic of all pestiviruses. This would relate to their capacity to interfere with pIC-induced IFN responses in M{phi} (Ruggli et al., 2003; Schweizer & Peterhans, 2001). Indeed, CSFV impairs pIC-induction of type I IFN in DCs and the recent report of Glew et al. (2003) has shown a similar effect of BVDV in bovine DCs. However, the current analyses demonstrating that the response of CSFV-infected DCs to maturation signals for DCs – including pIC – was not impaired contrasted with this effect on type I IFN inducibility. It would therefore appear that the DC maturation induced by pIC was IFN-independent. In fact, the independence of virus-induced DC maturation from secreted IFN-{alpha} has been reported recently (Lopez et al., 2003).

Considering this inability of CSFV to interfere with DC maturation, the question arose concerning the functionality of the DCs as antigen-presenting cells. Using antigen-presentation assays for polyclonal SEB stimulation, as well as FMDV- and CSFV-specific restimulation assays, it was observed that CSFV did not impair the capacity of DCs to process and present antigens. The recent report of Glew et al. (2003) has also shown that BVDV-infected bovine DCs retained their capacity to function as antigen-presenting cells. With the CSFV infections in the present report, it was particularly interesting to note that the infected DCs could still present CSFV antigen and stimulate CSFV-specific lymphocyte responses. Furthermore, apart from suppressed IFN type I responses, the cytokine profile of the DCs was not modified. For the interpretation of these results and particularly the cytokine responses, it is important to note that in vitro-derived DCs represent a cell culture model for myeloid DCs, which does not reflect the complexity of the DC system. Consequently, further studies investigating the interaction of CSFV with, for instance, plasmacytoid DCs are required in the future.

The interaction of CSFV with DCs can be seen as a double-edged sword. On one side, CSFV has developed a mechanism to prevent antiviral responses, permitting the virus to replicate in DCs and to use these highly migrating cells as a ‘taxi’ for transport to different sites in the host body. This would enable virus ‘delivery’ to lymphoid tissue, where it is know that considerable lymphocyte destruction occurs during CSF. The simple interaction between CSFV-infected DCs and lymphocytes is inadequate for the induction of such cell death. It would therefore appear that the environment particular to lymphoid follicles has a major role to play. On the other side, the present report demonstrates that CSFV-infected DCs do remain functional and are able to induce T cell responses against the virus.


   ACKNOWLEDGEMENTS
 
We thank Viviane Neuhaus for preparation of recombinant proteins, Heidi Gerber for mAb preparation and Drs Michael Horn and Nicolas Ruggli for critical discussions of the results. For animal care, we are grateful to D. Brechbühl and H.-P. Lüthi. This work was supported by the Swiss Federal Office for Education and Science (EU projects PL97-3732 and QLRT-2000-01374).


   REFERENCES
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Banchereau, J., Briere, F., Caux, C., Davoust, J., Lebecque, S., Liu, Y. J., Pulendran, B. & Palucka, K. (2000). Immunobiology of dendritic cells. Annu Rev Immunol 18, 767–811.[CrossRef][Medline]

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Received 14 October 2003; accepted 19 January 2004.