Cross-Presentation of Human Cytomegalovirus pp65 (UL83) to CD8⁺ T Cells Is Regulated by Virus-Induced, Soluble-Mediator-Dependent Maturation of Dendritic Cells

Virus and cells.HCMV strain AD169 was propagated in MRC5 human fibroblasts (BioMérieux, Marcy l’Etoile, France). Virus was collected when cytopathic effects were >90%. Supernatants were clarified by centrifugation at 1,500 × g for 10 min at 4°C and were stored at −80°C until they were used. The virus titer was determined by PFU titration in human foreskin fibroblasts (American Type Culture Collection) according to standard procedures. MRC5 (HLA-A2) cells and PBMC from blood donors were phenotyped for major histocompatibility complex (MHC) class I and class II by the Laboratoire Central d’Immunologie-Rangueil (E. Ohayon and M. Abbal, Rangueil Hospital, Toulouse, France). Infections with HCMV were performed at a multiplicity of infection (MOI) of 1 unless otherwise stated.

Generation of DCs.DCs were prepared from PBMC using a VacCell processor as described by Goxe et al. (13). Briefly, PBMC obtained from leukapheresis were cultured for 7 days in hydrophobic bags (Stedim, Marseille, France) in AIMV serum-free medium (Life Technologies, Cergy Pontoise, France) supplemented with granulocyte-macrophage colony-stimulating factor (GM-CSF; 500 U/ml; purchased from Novartis, Ruel Malmaison, France) and interleukin-13 (IL-13; 50 ng/ml; purchased from Sanofi-Synthelabo, Labège, France). Fresh IL-13 was added again after 4 days of culture. DCs were isolated on day 7 by elutriation. The isolation procedure gave rise to immature DCs with the phenotype CD1a⁺ MHC-I⁺ MHC-II⁺ CD64⁻ CD83⁻ CD80^low CD86^low, as described by Arrode et al. (2). DC purity was around 90%, and viability was >95%. Cytofluorimetric analysis of purified DCs revealed the absence of B lymphocytes (CD19⁺), macrophages (CD84⁺), and NK cells (CD56⁺) but contamination with red blood cells.

Assessment of cell death.Mycoplasma-free cells were cultured in either 24- or 6-well culture plates in RPMI 1640 (RPMI medium; Gibco, Cergy Pontoise, France) containing 10% fetal calf serum and supplemented with Glutamax-I, sodium pyruvate, and antibiotics, including the antimycoplasma antibiotic ciprofloxacine chlorhydrate (CIPRO; Bayer Pharma, Puteaux, France) (referred to as complete medium) unless otherwise stated. MRC5 cells were either mock infected or infected with HCMV AD169 for 6, 24, 48, or 72 h as described below. After incubation at 37°C under the conditions specified above, cells were detached with trypsin and added to those recovered from culture supernatant for washing with phosphate-buffered saline (PBS). The presence of dying cells was detected by multiparameter flow cytometry (Epics; Coulter Immunotech, Marseille, France) using propidium iodide (PI) according to the manufacturer’s instructions.

Phagocytosis assay.MRC5 cells were dyed red with PKH26 (Sigma) according to the manufacturer’s procedure and then infected with HCMV for various periods of time as indicated above or not infected. Red cells were then cocultured with immature DCs (one DC for one red cell) in complete medium supplemented with GM-CSF and IL-4 (a gift from J.-P. Marolleau, Paris, France) for 10 h at either 4 or 37°C. DCs were labeled with a fluorescein isothiocyanate-labeled mouse anti-HLA-DR antibody (L243; Coulter Immunotech) or with isotype control (immunoglobulin G2a-fluorescein isothiocyanate; Coulter Immunotech). Qualitative and quantitative phagocytosis of red cells by DCs was determined by flow cytometry and fluorescence microscopy.

Expansion of anti-pp65 CD8⁺ T cells from HCMV-seropositive donor PBMC.PBMC (2 × 10⁶/ml) from an HCMV-seropositive healthy HLA-A2 donor were cultured in 24-well plates in RPMI medium containing 10% human AB serum, 1% modified Eagle medium nonessential amino acids (Gibco), and 10 mM HEPES (Gibco). pp65-derived peptide corresponding to a known CTL HLA-A2 binding epitope (NLVPMVATV [N9V]) was obtained from Neosystem (Strasbourg, France). The restimulation procedure for PBMC has previously been described in detail (2). On days 3 and 7, recombinant human IL-7 (a gift from Sanofi-Synthelabo) was added at a final concentration of 25 ng/ml, and CD8⁺ T lymphocytes were obtained by positive selection with anti-CD8 magnetic beads (Miltenyi Biotec, Paris, France) according to the manufacturer’s procedure. The purity (>90%) of the isolated CD8⁺-cell subset was determined by dual staining with anti-CD4 and anti-CD8 antibodies (Coulter-Immunotech) and flow cytometry analyses. Activation of anti-pp65 CTL was assessed through quantitation of secreted IFN-γ by enzyme-linked immunosorbent assay (ELISA) as described below.

Assay for cross-presentation.Immature DCs (5 × 10⁵/well) obtained from HLA-A2 donors were added in medium supplemented with GM-CSF (100 ng/ml; Novartis) and IL-4 (50 ng/ml) to HCMV-infected or uninfected MRC5 cells (1:1 ratio) for 24 h. Then, the DCs were gently pipetted to dissociate them from adherent fibroblasts. The cells were washed, plated in duplicate on a 96-well plate at 5,000/well, and incubated for 24 h in the presence of the anti-pp65 T-cell line at different effector-to-DC ratios in a final volume of 200 μl. Alternatively, DCs were used either unloaded or pulsed overnight with 1 μM N9V peptide in the presence of tumor necrosis factor alpha (TNF-α; 50 ng/ml; R&D Systems, Abingdon, United Kingdom) for 24 h. To exclude the possibility of direct stimulation of T cells by HLA-A2-positive MRC5 cells that could be recovered during pipetting, MRC5 cells, either uninfected or infected with HCMV, were used as additional controls.

For transwell studies, immature DCs were added to 3-μm-pore-size transwell chambers (Becton Dickinson, Le Pont de Claix, France) inserted into wells containing HCMV-infected or uninfected MRC5 cells (1:1 ratio) and then cultured for 24 h. Alternatively, immature DCs were cultured for 24 h with clarified supernatants that were obtained from infected or uninfected fibroblasts and then incubated overnight with 1 μM N9V peptide or not incubated.

ELISA for IFN-γ secretion.ELISA for IFN-γ secretion by activated CTL was performed as follows. Ninety-six-well plates (Nunc) were coated overnight at 4°C with primary anti-IFN-γ monoclonal antibody (MAb; Biosources), washed, and blocked for 2 h at 37°C with the buffers provided (Medgenix screening line). Diluted (1/10) supernatants from CTL expansion experiments were added to precoated wells in duplicate and supplemented with secondary antibody (biotin-conjugated anti-IFN-γ MAb; Biosources) for 2 h. After the plates were washed with PBS-fetal calf serum (1%), streptavidin-bound peroxidase (Coulter Immunotech) was added and left for 30 min at room temperature. The plates were then washed and incubated for 5 min in chromogen buffer (Biosources). The optical density was measured on an ELISA apparatus (Dynatech Laboratories).

Analysis of DC phenotype.Mature DCs used as a control were produced by incubation for 24 h with medium containing GM-CSF and IL-4 and supplemented with TNF-α (50 ng/ml). In the case of coincubation with MRC5 cells, immature DCs were directly added to HCMV-infected or uninfected cells (1:1 ratio) and cocultured for 24 h. Alternatively, coincubations were performed with clarified supernatants that were obtained from infected or uninfected MRC5 fibroblasts, as described below. After the treatments, the cells were harvested, washed in PBS supplemented with 5% fetal calf serum, and incubated for 30 min at 4°C with either phycoerythrin (PE)-conjugated anti-CD83 antibody (HB15A; Coulter Immunotech) or isotype control (immunoglobulin G-PE; Coulter Immunotech). The mature DC phenotype (CD83-positive cells) was detected by flow cytometric analysis (Coulter Epics) by gating the cell population of interest on forward scatter-side scatter.

Quantification of TGF-β1 in culture supernatants and neutralizing assays.MRC5 cells were either mock infected or infected at an MOI of 1 to 5 with HCMV AD169. The virus was adsorbed for 30 min to 24 h at 37°C, and the cells were washed twice with PBS. They were cultured with complete medium in coincubation experiments or with serum-free medium containing 100 μg of bovine serum albumin (Sigma) when TGF-β1 quantification was performed. Supernatants from infected cells were collected in the time ranges 0 to 6, 6 to 24, 24 to 48, and 24 to 72 h, and 0 to 72 h for uninfected cells. They were clarified at 900 × g for 10 min at 4°C and then were either stored at −80°C for ELISA or incubated with immature DCs.

Latent and active TGF-β1 protein was measured by specific ELISA in both serum-free medium and supernatants. To activate the latent form of TGF, samples were acidified for 1 h by dilution at 1/10 in 1 N HCl and then neutralized with 1 N NaOH and tested immediately by ELISA. To quantify TGF-β1, 96-well plates were coated overnight at 4°C with 1 μg of an anti-TGF MAb/ml (MAb 2G7, which recognizes both active and latent forms of TGF-β1, TGF-β2, and TGF-β3) (21). Serial dilutions of medium and supernatants, either acidified or not (100 μl/well), were added to precoated wells and incubated for 1 h at 37°C. After three washes with PBS-Tween (0.1%), biotinylated chicken anti-human TGF-β1 antibody (BAF240; R&D) was added to the wells for one additional hour at 37°C. Binding of biotinylated antibodies was revealed by incubation with streptavidin-conjugated alkaline phosphatase (Jackson). The plates were then washed with PBS-Tween (0.1%) and incubated for 30 min with nitrophenyl phosphate substrate (Sigma). The optical density was measured on an ELISA apparatus (Dynatech Laboratories), and the amounts of TGF-β1 (in picograms per milliliter) were derived from a standard curve constructed using human recombinant TGF (R&D). For neutralization assays of TGF-β1, MRC5 cultures were treated with either an anti-TGF-β1 neutralizing antibody (MAB1032; Chemicon) or an isotype control antibody (Dako), both at 10 μg/ml for 2 h at 37°C, before DCs were added. Then, cross-presentation assays were performed as described above.

DCs cocultured with HCMV AD169-infected fibroblasts stimulate pp65-specific CTL.We have previously shown that cross-presentation of HCMV antigen pp65 by DCs occurred in the presence of AD169-infected apoptotic fibroblasts (2). Moreover, we have also speculated that cell lysis due to cytopathic effects of the virus could serve as an exogenous antigen source for cross-presentation by DCs. To assess this question, immature DCs that are not permissive to HCMV AD169 (25) were added directly to wells containing mock-infected or HCMV-infected MRC5 fibroblasts (1:1 ratio) at various times (6 to 72 h) postinfection (p.i.) and maintained in coculture for 24 h. According to our previous report showing that cross-presentation occurred under conditions where most cells were in late apoptotic and necrotic stages corresponding to PI-positive populations, we first analyzed the phenotypes of infected cells by labeling the cells with propidium iodide. As shown in the histograms (Fig. 1A), infection of fibroblasts for 24 h resulted in the production of a small but significant percentage (20%) of PI-positive cells compared to the percentage with uninfected cells (3%). Microscopic analysis of coculture wells revealed behavioral and morphological changes of immature DCs, as shown in the photomicrograph in Fig. 1A. Indeed, we noted that coculture for 24 h with uninfected monolayer fibroblasts kept immature DCs in a resting state, whereas in the presence of infected cells (24 h p.i.), DCs were mainly clustered. It has been shown that such interactions facilitate antigen transfer between live DCs (14).

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FIG. 1.

DCs cocultured with HCMV-infected fibroblasts stimulate pp65-specific CTL. HLA-A2-positive DCs were added, either directly or in a pore transwell insert, to wells containing uninfected (DC+MR ni) or HCMV-infected MRC5 cells (MOI = 1) at the indicated times p.i. (DC+MR 6hpi to 72hpi), for 24 h. (A) Before DCs were added, uninfected or 24-h-p.i. MRC5 cells were labeled with PI (histograms) and detected by flow cytometry. Cocultures were also examined by optical microscopy. (B) DCs were recovered from cocultures and incubated with HLA-A2-restricted anti-pp65 CTL, raised as described in Materials and Methods, at different responder-to-stimulator ratios (R/S). DCs alone or treated with TNF-α and a relevant peptide (DC N9V TNF) were also used as stimulator cells. Additional controls consisted of uninfected or infected MRC5 cells at each time p.i. (not shown). Stimulation of CTL was assessed through quantitation of IFN-γ secretion using ELISA. The results shown are representative of two to six independent experiments.

To study whether DCs could acquire viral antigens in such cocultures, HLA-A2-positive DCs were gently pipetted to dissociate them from adherent fibroblasts. After being washed, they were incubated with HLA-A2-restricted anti-pp65 CD8⁺ CTL at different stimulator-to-responder ratios. Cross-presentation of pp65 was monitored by quantification of IFN-γ secretion using ELISA. Figure 1B shows that DCs cocultured for 24 h with 24-h-p.i. MRC5 cells specifically induced the activation of anti-pp65 CTL. We ruled out the possibility that few contaminating CD4⁺ T cells could respond, since MHC class II did not match between DCs and T-cell lines. Under these conditions, activation with 24-h-p.i MRC5 cells was higher than that following incubation with 48- or 72-h-p.i. MRC5 cells. To exclude the possibility of direct stimulation of CTL by HLA-A2-positive MRC5 cells that could have been recovered during pipetting, uninfected or HCMV-infected MRC5 cells were used as controls. Regardless of the time p.i., neither HCMV-infected fibroblasts nor DCs alone were able to induce significant CTL activation (data not shown). In contrast, TNF-α-treated DCs pulsed with N9V peptide, considered optimal for both maturation and antigen loading, led to maximal activation of CTL. These results indicate that immature DCs are able to acquire viral antigens from cocultured infected fibroblasts and to present them to CD8⁺ CTL. Although variations exist between independent experiments, globally this process was more efficient in the early stages of infection (6 to 48 h p.i.) and was always reduced when infection time was extended (72 h p.i.).

In vitro studies showed that antigens could access the exogenous pathway for class I presentation by DCs in various forms, including apoptotic and necrotic cells; exosomes; microorganisms, such as bacteria and viruses; and proteins following contact-dependent transfer from live cells (19). In the case of virus-generated damage, such as cytopathic effects caused by HCMV infection, both dead cells or their debris and viral proteins released from infected cells could be sources of viral antigens. Since DCs were not permissive to HCMV AD169, we determined by using a transwell system whether antigen loading of DCs could occur through transfer of viral proteins from infected cells. DCs were added to 3-μm-pore-size transwell chambers inserted into wells containing uninfected or infected MRC5 cells (1:1 ratio) at various times p.i., incubated for 24 h, and recovered for assay of CTL activation. Regardless of the time p.i., activation of specific CTL by DCs was completely abrogated. This indicates that cell-to-cell contact is required for DCs to acquire viral antigens from infected fibroblasts.

DCs phagocytose PKH26⁺ cells after coculture with early HCMV-infected cells and cross-present incoming pp65 to CTL.Since we had previously shown that in the very early time after infection of MRC5 fibroblasts the matrix protein pp65 from the viral inoculum contained in dead cells was available for cross-presentation, we asked whether this could occur in coculture experiments. To this end, MRC5 cells were infected at a higher MOI (MOI = 3) than that used in the experiment shown in Fig. 1 and cocultured with DCs for 24 h. Figure 2A shows that under these conditions, pp65 contained in very early infected fibroblasts (6 h p.i.) was cross-presented to CTL and that, as shown above (Fig. 1), stimulation was reduced in cocultures with late-infected cells. These data are in agreement with our previous results demonstrating that incoming pp65 was available for cross-presentation to CTL. Labeling of 6-h-infected MRC5 cells with PI revealed that 15% (Fig. 2B, left) of the cells were positive compared with 3% of uninfected cells (data not shown), suggesting that dying infected cells could be a source of antigen uptake by DCs. To clarify the mechanism by which DCs could acquire viral antigen, MRC5 cells were sequentially labeled with PKH26, infected with HCMV for 6 h, and cocultured for 12 h with DCs (1:1 ratio) at either 4 or 37°C. To quantify the uptake of PKH26-labeled material by DCs, the number of HLA-DR⁺ PKH26⁺ DCs was analyzed by flow cytometry. Figure 2B (right) shows that 30% of DCs were PKH26⁺ after being cocultured with 6-h-p.i. MRC5 cells. Flow cytometry analysis was confirmed by fluorescence microscopy as previously described (2) (data not shown). Thus, our results suggest that very early after infection of fibroblasts, incoming pp65 could participate in antigen loading of DCs for stimulation of CTL and that phagocytosis of dying cells could take part in this process.

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FIG. 2.

DCs phagocytose PKH26⁺ cells during coculture with early-infected MRC5 cells and cross-present pp65 to CTL. (A) HLA-A2-positive DCs were added to wells containing uninfected (DC+MR ni) or HCMV-infected MRC5 cells (MOI = 3) at the indicated times p.i. (DC+MR 6hpi to 72hpi) and were maintained in coculture for 24 h. The experiment was extended as described in the legend to Fig. 1B. (B) (Left) Mock-infected (not shown) or 6-h-p.i. MRC5 cells were labeled with PI. The indicated percentages of PI⁺ cells were detected by flow cytometry. (Right) PKH26-labeled MRC5 cells infected for 6 h were cocultured with immature DCs for 12 h at 4 or 37°C at a DC/MRC5 cell ratio of 1:1. The uptake of infected MRC5 cells by HLA-DR⁺ DCs was analyzed by flow cytometry; dot plots were gated on FL1 (HLA-DR) high-positive cells (DCs), thus excluding the uncleared material. Similar results were obtained in three independent experiments. R/S, responder-to-stimulator ratios.

DC maturation is regulated by supernatant of HCMV-infected fibroblasts.It has been shown that optimal cross-presentation of antigens from dead tumor cells required a maturation signal in DCs provided by necrotic cells (29). To demonstrate whether HCMV-infected fibroblasts can deliver a maturation signal to DCs, the expression of CD83, an adhesion receptor highly restricted to mature DCs (30), was determined. Immature DCs were added directly to HCMV-infected or uninfected fibroblasts (1:1 ratio) and cultured for 24 h. Coculture with 6- and 24-h-p.i. MRC5 cells induced maturation of DCs at a higher ratio than TNF-α-treated DCs, as revealed by the appearance of a CD83⁺ population. Furthermore, this maturation was reduced in coculture with fibroblasts that had been infected for 48 h and almost completely abolished in cells infected for 72 h (Fig. 3A). Uninfected fibroblasts kept DCs in an immature state. Expressions of MHC class I and class II surface molecules, which are known to be upregulated in mature DCs, were slightly regulated following coculture with infected fibroblasts (data not shown).

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FIG. 3.

DC maturation following interaction with HCMV-infected fibroblasts or their supernatant. Immature DCs (alone) were added directly to uninfected (+M ni) or infected MRC5 cells at the indicated times p.i. (+M 6h pi to M 72h pi) (A) or to supernatant from uninfected (+SN ni) or infected MRC5 cells (+SN 6h pi to 72h pi) (B). TNF-α-treated DCs (TNF) were used as a positive control for maturation. All of the cells were collected 24 h after coculture and labeled with an anti-CD83-PE antibody or isotype control. CD83 expression was detected by flow cytometry analysis by gating the DCs on forward scatter-side scatter. Four experiments were performed and gave similar results.

These data, as well as those reported above, suggest that when DCs are added to infected cells at a given time, conditions are combined to provide signals for DC maturation and sufficient pp65-positive cells for antigen loading, which are both essential for optimal activation of specific CD8⁺ T cells.

We then investigated the mechanisms responsible for regulation of DC maturation by HCMV. Contrasting results have been reported concerning interference of viruses with the maturation process of DCs. For instance, ingestion of debris from canarypox virus-infected cells by DCs partially drove their maturation (15), whereas infection by vaccinia virus induced synthesis of soluble factors that affect DC maturation (32). To establish whether soluble factors released by HCMV-infected cells were involved in modulation of their maturation, DCs were cultured for 24 h with either infected MRC5 cells or their clarified culture supernatant. As shown in Fig. 3, maturation of DCs was modulated to the same extent under both conditions, with a delayed maturation in the presence of supernatant recovered from 72-h-p.i. MRC5 cells. These results show that soluble factors released by infected cells have direct but opposite effects on DC maturation if we consider medium from early- or late-infected fibroblasts.

Supernatants from infected cells are sufficient to modulate the T-cell-stimulatory capacity of DCs.The most distinctive functional characteristic of mature DCs is their ability to potently stimulate T cells (20). Therefore, we assessed whether supernatant from infected cells could modulate the T-cell-stimulatory capacity of DCs. To answer this question, DCs were either directly cocultured as described in the legend to Fig. 1B or cultured for 24 h with clarified supernatant of infected cells supplemented or not with HLA-A2 binding N9V peptide. DCs pulsed with N9V and treated with TNF-α to stimulate their maturation or not treated were used as controls. DCs were recovered and used as stimulators in coculture with specific anti-pp65 CD8⁺ CTL. IFN-γ secretion by activated T cells was measured by ELISA. Figure 4A indicates that DCs cocultured for 24 h with 48-h-p.i. MRC5 cells induced the highest activation of HLA-A2-restricted anti-pp65 CTL. This activation was always reduced following incubation with 72-h-p.i. MRC5 cells. In experiments using supernatants that were supplemented with N9V peptide, specific activation of CTL was modulated to the same extent as in direct coculture (Fig. 4B). No activation was observed when supernatants alone were used in these assays. Moreover, immature DCs pulsed with N9V were relatively poor stimulators of CTL. Reduction of CTL response to N9V peptide in the presence of 72-h-p.i supernatant was not due to a decrease of MHC class I expression on DCs as assessed by flow cytometry (data not shown). These results demonstrate that soluble factors released from infected cells have direct effects on the T-cell-stimulatory function of DCs.

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FIG. 4.

Supernatants from infected fibroblasts are sufficient to modulate T-cell-stimulatory capacity of DCs. HLA-A2-positive DCs were directly cocultured with uninfected (DC+ MR ni) or infected fibroblasts at the indicated times p.i. (DC+MR6hpi to -72hpi) for 24 h (A) or cultured for 24 h with clarified supernatant recovered from uninfected (MR ni) or infected cells at different times of infection (MR6hpi to -72hpi) supplemented (DC+Supernatant+N9V) or not (DC+Supernatant alone) with a relevant peptide (N9V) (B). DCs pulsed with N9V and treated (DCN9V TNF) or not (DCN9V) with TNF-α were also used as controls. After coculture, recovered DCs were treated as described in the legend to Fig. 1B, and the same controls were used. One representative experiment of two is shown. R/S, responder-to-stimulator ratios.

TGF-β1 released by late-infected fibroblasts takes part in down regulation of both DC maturation and T-cell activation.The results discussed above strongly support the notion that a soluble factor contained in the supernatant of late-infected fibroblasts is responsible for the failure of CD83 induction of DCs and of their T-cell-stimulatory ability. It has been reported that several cytokines, including IL-10 and TGF-β, caused down regulation of expression of CD83 and other costimulatory molecules on DCs (33). Michelson and coworkers (22) reported induction of TGF-β1 secretion following HCMV infection of human foreskin fibroblasts, suggesting that this cytokine could modify systemic immune reactions to the advantage of the virus by both up regulating its replication and down regulating host immune responses. Accordingly, we focused our investigations on TGF-β1 as a potential mediator of DC maturation. Thus, we first examined the production of TGF-β1 in the supernatant of mock-infected and HCMV-infected MRC5 cells by ELISA. Table 1 shows that TGF-β1 was detected in increasing amounts in infected-cell supernatants. Indeed, the amount of TGF-β1 was three times higher in supernatants recovered from 24- to 72-h-p.i. cells than in supernatants from mock-infected MRC5 cells. To test whether TGF-β1 production by infected cells was at least in part responsible for impairment of DC maturation, we used neutralizing antibody against TGF-β1. Uninfected or 48- and 72-h-p.i. MRC5 cultures were incubated for 2 h at 37°C with either anti-TGF-β1 neutralizing antibody or an isotype-matched control. DCs were then added to pretreated MRC5 cells and incubated for 24 h, and their CD83 status was determined by flow cytometry. As shown in Fig. 5, down modulation of CD83 expression in DCs cultured with 72-h-p.i. MRC5 cells was abrogated by the anti-TGF-β1 neutralizing antibody but not by the isotype control antibody. These results allowed the identification of TGF-β1 as an essential mediator of the decreased expression of CD83 on DCs following incubation with late-infected fibroblasts. To further determine whether TGF-β could interfere with CTL activation, IFN-γ release by anti-pp65 CTL was assessed following neutralization of TGF-β1 in MRC5 cultures. Under these conditions, T-cell activation was not recovered to a highly significant extent (data not shown), suggesting that unknown soluble factors contained in late supernatant may down regulate costimulatory molecules other than CD83 that are critical for CTL response.

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FIG. 5.

TGF-β1 released by late-infected fibroblasts is essential in failure to induce CD83 expression on DCs. Cultures of uninfected (MR ni) or infected MRC5 cells at 48 and 72 h p.i. were incubated with a neutralizing anti-TGF-β antibody at 10 μg/ml or an isotype control for 2 h at 37°C. Immature DCs were then added to neutralized conditioned medium and incubated for 24 h. The cells were collected and labeled with an anti-CD83-PE antibody. The flow cytometry histograms show percentages (± standard deviations) of CD83-positive cells representative of three independent experiments.

Altogether, our data suggest that cross-presentation of pp65 to CTL is regulated by soluble-mediator-dependent maturation of DCs.