To study whether murine DCs are susceptible to MVA infection, naïve BALB/c splenocytes were infected in vitro with rMVA-GFP at a MOI of 10 for 8 h followed by flow cytometric detection of EGFP expression in different cell populations. As shown in Fig. 1A, despite the presumed broad cellular tropism of VV and related viruses, rMVA-GFP did not infect all subsets of splenocytes to an equivalent extent, but rather demonstrated a striking preference for professional APC populations, with CD11c⁺ DCs being the most susceptible targets for MVA infection in the spleen (31.2 ± 2.9% GFP⁺), followed by CD11b⁺ CD11c^- macrophages (20.1 ± 2.8% GFP⁺) and CD19⁺ B cells (8.5 ± 2.1% GFP⁺). Very few GFP⁺ cells were detected in CD4⁺ (6.9 ± 1.2%) and CD8⁺ (3.6 ± 1.5%) T cell populations. Further phenotypic analysis revealed that over 50% of all GFP⁺ cells were CD11c⁺ DCs (data not shown). Therefore, professional APCs, especially DCs and macrophages, seem to be preferentially targeted by MVA among all the splenocytes subsets in the in vitro infection setting. This was held true when splenocytes were infected at a range of MOIs (from 0.3 to 10) (data not shown). At all the MOIs tested, DCs were by far the most susceptible targets among all the splenocytes subsets, followed by macrophages and B cells.

To confirm the cellular tropism of MVA infection in vivo, we infected BALB/c mice intravenously with 3 × 10⁸ PFU rMVA-GFP. At various time points post infection, spleens were harvested and MVA infected GFP⁺ splenocytes were identified by cell lineage marker staining and flow cytometry. The number of GFP⁺ cells in spleen peaked at 9 h post infection and rapidly declined to undetectable level by 24 h post infection (data not shown). At 9 h post infection, GFP expression was primarily detected in CD11c⁺ DC population (12.8 ± 2.1% GFP⁺), but not any other cell types including macrophages, as shown in Fig. 1B. This result suggests that MVA preferentially infects DCs in secondary lymphoid organ in vivo. Since DCs are vastly outnumbered by B cells, T cells in spleen, this result, together with the in vitro infection data (Fig. 1A), establish that among various subsets of hematolymphoid cells, DCs are preferentially targeted by MVA in a highly effective manner.

MVA has lost its ability to replicate in most primary mammalian cells. The defect in the viral life cycle has been shown to be the final steps of viral morphogenesis, with no alteration in early or late virus gene expression 6. To determine whether both early and late MVA genes were expressed from the infected murine DCs, we measured the expression of GFP (driven by an early promoter) and a late viral gene A56R from the in vitro MVA-infected BMDCs as well as purified CD11c⁺ splenic DCs, as described in Methods. The permissive chicken embryonic fibroblasts DF-1 cells were infected in parallel as positive controls for MVA gene expression. Cells were infected with rMVA-GFP at a MOI of 10 for 20 h. The cells were then harvested and subjected to intracellular staining for anti-A56R mAb, as described in Methods. As shown in Fig. 2, the expression of the early gene GFP was easily detected in over 60% of cultured BMDCs and 40% of splenic DCs. However, no A56R (late gene) expression was detected in either DC preparations (Fig. 2A&B), while almost 100% DF-1 cells expressed both GFP and A56R. Therefore, the MVA life cycle is blocked in murine DC at an earlier stage than the virion assembly defect reported for other non-permissive mammalian cell lines 612. These data are consistent with the report that only early but not late MVA genes are expressed from the infected human DCs 1224.

Targeted infection of MoDC by VV has been suggested as a viral immune evasion strategy, as DC maturation was shown to be blocked by VV infection 2024. To study whether MVA infection affects the maturation process of murine DC, we monitored the expression of maturation markers by DCs following rMVA-GFP infection. Initially splenic DCs were used in the experiments. However, these cells underwent rapid apoptosis within 12 h following in vitro isolation, even without infection (data not shown). BMDCs were then used to study the effect of MVA infection of DC maturation. BMDCs were infected with rMVA-GFP at a MOI of 10, mock-treated or stimulated with LPS. Mock-treated BMDCs were used to reflect the spontaneous BMDC maturation induced by the "mechanical" manipulation. LPS treatment was included to induce maximal BMDC maturation. At 0, 12 and 18 h following infection, cells were harvested and stained with mAbs specific for CD40, CD86, CD80, and MHC class II I-A^d. As shown in Fig. 3, as early as 12 h post infection there was a significant upregulation of all the maturation markers on the majority of infected BMDCs. By 18 h post infection, the virus-infected DCs further upregulated the maturation markers. The phenotype of the MVA-infected DCs was comparable to that of LPS-stimulated DCs at both 12 h and 18 h post infection. These data suggest that MVA infection induced rapid BMDC maturation. This is in direct contrast to the reported blockage of human DC maturation by VV infection 192024.

Activated DCs secrete an array of cytokines and chemokines that are crucial for amplification of both innate and adaptive immune responses 17. To test whether MVA infection of murine DCs could induce cytokine production, we measured the secretion of the pro-inflammatory cytokine IFN-α from DCs following MVA infection. BMDCs and splenic CD11c⁺ DCs were either mock-treated or infected with rMVA-GFP at a MOI of 10. Supernatants were harvested at 10 and 24 h post infection, and IFN-α levels were measured by ELISA. As shown in Fig. 4, substantial amount of IFN-α was readily detected from both BMDC and splenic DCs as early as 10 h after rMVA-GFP infection. The amount of IFN-α detected in the supernatant samples was further increased by 24 h post infection. These results suggest MVA-infected DCs not only undergo rapid phenotypic maturation, but also are capable of prompting host immune responses via pro-inflammatory cytokine production. Noticeably, endogenous splenic DCs produced significantly higher amount of IFN-α upon MVA infection compared to BMDCs (5-fold and 3-fold difference at 10 h and 24 h, respectively).

Studies of VV infection of human MoDCs have suggested that DC undergo substantial apoptosis rapidly following the infection 1220. To study whether this is also true for MVA infected murine DC, we next assessed the viability of BMDCs at different time points following MVA infection. Immature BMDCs were either infected with rMVA-GFP at a MOI of 10 or mock treated. Cells were harvested at various time points following the treatment for viability count. As shown in Fig. 5A, immature BMDCs underwent substantial apoptosis starting 12 h following MVA infection, earlier than VV-infected BMDCs which did not undergo significant apoptosis until 18 h post infection (unpublished results). By 24 h following infection, only 50% of the MVA-infected cells remained viable, while 90% of mock-treated cells were still alive. Over 70% of the infected cell had been dead by 42 h following MVA infection. We further compared the viability of immature and mature BMDCs following rMVA-GFP infection. BMDCs were stimulated with LPS for 24 h to reach maturation. Both immature and mature BMDC were then infected with rMVA-GFP or mock-treated for 24 h and the cell viability was determined. As shown in Fig. 5B, MVA infection reduced the viability of both immature and mature BMDC compared to mock treatment. Interestingly, mature BMDCs displayed higher viability than immature BMDCs at this time point. This data suggests that upon maturation, DCs become somewhat more resistant to MVA-induced cell death. This may be of particular importance for direct antigen presentation.

The rapid apoptosis of MVA-infected DCs indicates substantial proportion of infected DCs may not present Ags to T cells directly, but rather be uptaken by uninfected bystander DCs for cross-presentation of the viral Ags. We then investigated whether the MVA- infected DCs undergoing apoptosis could be phagocytosed by uninfected DCs. Green fluorescence (PKH-67)-labeled BMDCs were first infected with rMVA for 18 h, then incubated with red fluorescence (PKH-26)-labeled uninfected BMDCs at 1:1 ratio at either 37°C or 4°C for 4 h. Phagocytosis were monitored by flow cytometry. Following incubation at 37°C, one third of the uninfected DCs (30.1 ± 5.3%) became PKH-67⁺ PKH-26⁺, while few (2.1 ± 1.9%) uninfected DCs picked up PKH-67⁺ cells at 4°C (Fig. 6). These data suggest that following MVA infection, the infected DCs can be engulfed by uninfected DCs efficiently and rapidly.

Having shown that MVA-infected DCs were readily phagocytosed by uninfected bystander DCs, we then hypothesized that MVA Ags can be presented to T cells through cross-presentation. To test this hypothesis, rMVA expressing LCMV NP Ag (rMVA-NP) or rMVA-GFP was used to infect BMDCs from either WT C57BL/6 mice or MHC class I deficient β2m^-/- mice. DCs were infected at a MOI of 10 for 6 h before intravenously injected into LCMV immune mice. To prevent direct infection of host APCs by the infectious viruses co-injected with BMDCs, DCs were extensively washed and irradiated prior to the injection. Five days following DC injection, spleens were harvested and single cell suspensions were prepared and CTL activity specific to the dominant LCMV epitope NP_396–404 was determined by fluorescent cellular cytotoxicity assay, as described in Methods. Strong NP_396–404 – specific CTL activity was detected in mice immunized by rMVA-NP-infected WT DCs, but not in mice immunized with rMVA-GFP-infected BMDCs (Fig. 7). Interestingly, NP_396–404-specific CTL activity was also detectable, although to a lesser degree, in mice immunized with rMVA-NP-infectedβ2m^-/- DCs. Since β2m^-/- DCs cannot directly present MHC class I-restricted Ags to T cells, these data suggest that the detected CTL activity is due to cross-presentation of NP Ags by the endogenous APCs that have phagocytosed the injected β2m^-/- DCs.

To definitively answer the question whether DCs are responsible and required for the in vivo T cell activation following MVA infection, we took advantage of the CD11c-DTR mice that express human DT receptor under the control of the CD11c 25. DT injection has been shown to effectively deplete CD11c⁺ DCs within 24 h. We depleted DCs from CD11c-DTR mice by i.p. injection of DT (12 ng/g body weight) 2526 and infected the mice with 5 × 10⁶ PFU MVA intraperitoneally 18 h later. DT injection was repeated on day 2 after infection to ensure continuous absence of DC. On day 6 after MVA infection, spleens were harvested and the efficiency of DC depletion was determined by flow cytometry. Over 95% of CD11c⁺ DCs were depleted and the residual DCs had CD11c^low B220⁺ phenotype, similar to that of plasmacytoid DCs (Fig. 8A). Single cell suspensions prepared from the spleens were re-stimulated with C57BL/6 splenocytes infected with MVA for 6 h in the presence of Brefeldin A. Following the re-stimulation, the frequencies of IFN-γ-producing effector CD3⁺ T cells specific for MVA antigens were determined by intracellular IFN-γ-staining, as described in Methods. As shown in Fig. 8B, MVA-specific effector T cells producing IFN-γ were detected in the spleens of WT littermates on day 6 after MVA infection. These cells accounted for 1.75 ± 0.05% of total CD3⁺ T cells (Fig. 8C). However, in DC-depleted CD11c-DTR mice, we failed to detect any IFN-γ-production from CD3⁺ T cells (Fig. 8B and 8C). These data suggest that DC-mediated Ag presentation is required for the generation of IFN-γ producing effector T cells in vivo following MVA infection.

All animal work has been reviewed and approved by the Center for Animal Resources and Comparative Medicine at Emory University and Harvard Medical School. 6- to 8-week-old female BALB/c mice and C57BL/6 mice were purchased from the Jackson Laboratories (Bar Harbor, ME). MHC class I-deficient B6.129P2-B2mtm1Unc (β2m^-/-) mice and CD11c-DTR-EGFP (CD11c-DTR) breeders 25 were purchased from the Jackson Laboratories and bred in a biosafety level-1 facility at Harvard Medical School. Virus-infected mice were housed in a biosafety level -2 facility.

rMVA strains were constructed to express GFP-ZEOCIN (rMVA-GFP) or lymphocytic choriomeningitis virus (LCMV) nuclear protein (NP) -ZEOCIN (rMVA-NP) 2852 under the control of the early H5 promoter and were plaque-purified at least 5 times as previously described using standard protocol 53. Non-recombinant MVA stock used for the construction of rMVA was kindly provided by Dr. Bernard Moss (National Institutes of Health, Bethesda, MD). MVA for the phagocytosis experiment and for the infection of CD11c-DTR mice were kindly provided by Dr. Michael S. Seaman (Beth Israel Deaconess Medical Center, Boston, MA). rMVA-GFP and rMVA-NP stocks were prepared in and titered on the chicken embryo fibroblast cell line DF-1 (gift of Dr. Harold Varmus, National Institutes of Health, Bethesda, MD). Viruses were concentrated by ultracentrifugation over 36% (weight/volume) sucrose, and titers were determined using standard plaque assays. LCMV stocks (Armstrong strain) were kindly provided by Dr. Rafi Ahmed (Emory Vaccine Research Center, Atlanta, GA).

DF-1 and HeLa cells are described above. These cell lines were maintained in complete DMEM supplemented with 10% FBS, 2 mM L-glutamine and antibiotics. EL4 cells (H-2^b) (ATCC) were maintained in RPMI supplemented with 10% FBS, 2 mM L-glutamine and antibiotics. Splenic DCs were isolated from the spleens of BALB/c mice that received daily injections of 20 μg of recombinant flt3L-IgG₂ for 9 days 5455. Briefly, spleen fragments were digested with 1 μg/ml collagenase (Worthington, Lakewood, NJ) for 30 min, followed by erythrocyte depletion with RBC lysing buffer (Sigma-Aldrich Co., St. Louis, MO). DCs were then isolated from this single cell suspension using CD11c magnetic beads (Miltenyi Biotec, Auburn, CA) according to the manufacturer's instructions, and the purity of DCs was greater than 90% as evaluated by flow cytometry (data not shown).

Bone marrow-derived DCs (BMDCs) were generated from mouse bone marrow cultures as described previously with minor modifications 56. Briefly, bone marrow was flushed from femurs and tibias and depleted of erythrocytes with RBC lysing buffer. Cells were subsequently plated on 6-well tissue culture plates at 4 × 10⁶/well in 3 ml complete RPMI 1640 medium (CM, contains 10% FBS, 2 mM L-glutamine, and 10 μg/ml gentamicin sulfate), incubated at 37°C for 3 h, and then the non-adherent cells were removed. CM containing 10 ng/ml rmGM-CSF and 3 ng/ml rmIL-4 (both from R&D Systems, Minneapolis, MN) was added at 3 ml per well and the plates were incubated at 37°C for 1 week. Cultures were fed by gently aspirating 2 ml/well of medium and adding back fresh CM containing cytokines on days 2, 4 and 6 of the culture. When mature DCs were required, 10 μg/ml LPS (Sigma-Aldrich Co.) was added to the culture on day 6 for 24 h. On day 7, non-adherent cells were harvested, and the purity of the DCs culture was evaluated by cell surface staining with fluorescence-conjugated mAbs to CD11b (M1/70), CD11c (HL3), CD3 (145-2C11), and CD45R/B220 (RA3-6B2) (BD Biosciences Pharmingen, San Diego, CA). Approximally 70–90% of the cells were CD11c⁺, as determined by flow cytometry (data not shown).

For in vitro infections, cells were resuspended at 1 × 10⁷/ml CM in 15 ml polypropylene tubes (Becton Dickinson Labware). An appropriate volume of the indicated virus stock was added to reach the multiplicity of infection (MOI) required for each experiment. Following 1 h-incubation at 37°C, cells were washed three times with medium, resuspended at 1 × 10⁶/ml in CM on 24-well plates and cultured at 37°C until harvesting. Mock-treated cells were manipulated in the same way except that no virus was added. For in vivo i.v. infection, BALB/c mice were i.v. injected with 15 μg DNA plasmid pNGVL3-hFLex 57 (obtained from National Gene Vector Laboratory, University of Michigan, and provided by Dr. Abdul M. Jabbar, Emory Vaccine Center) in 1.6 ml PBS to expand DC population in vivo. Five days later, mice were infected i.v. with 3 × 10⁸ plaque forming units (PFU) of rMVA-GFP in 0.5 ml PBS. Uninfected mice were injected with 0.5 ml PBS as controls. For intraperitoneal (i.p.) infection, CD11c-DTR mice and wild type (WT) littermates were infected with 2 × 10⁶ PFU MVA.

At the indicated time points post viral infection, splenocytes were stained with mAbs against CD3, B220, CD11b, CD11c, CD8a (Ly-2) (53-6.7), and CD4 (L3T4) (RM4-5) (BD Biosciences Pharmingen). At the indicated time points following in vitro infection, DF-1 cells, BMDCs, and splenic DCs were stained with purified anti-vaccinia late surface protein A56R using mAb VV1-4G9 (kindly provided by Dr. Alan Schmaljohn, USAMRIID, Frederick, MD) and secondary PE-conjugated goat anti-mouse IgG (Biosource International, Camarillo, CA). BMDCs and splenic DCs were also stained with mAbs against DC maturation markers CD80 (16-10A1), CD86 (GL1), CD40 (5C3), and I-A^d (AMS-32.1) (BD Biosciences Pharmingen). All staining steps were performed by incubating cells and Abs at 4°C for 20 min followed by three washes with staining buffer. Samples were then processed using a FACSCalibur flow cytometer (Becton Dickinson Labware) and the data were analyzed using Flowjo software (Tree Star, Inc., Ashland, OR).

BMDCs or splenic DCs were mock- treated or infected with rMVA-GFP. Supernatant samples were harvested at 10 or 24 h following the treatment and stored at -80°C if not used immediately. IFN-α in the supernatant was detected using a commercial colorimetric sandwich ELISA kit (R&D Systems) according to manufacturer's instructions.

Viability of BMDCs was determined by trypan blue exclusion at various time points following infection.

BMDCs were labeled with the green fluorescent dye PKH-67 (Sigma-Aldrich Co.) according to the manufacturer's instructions and infected with MVA at a MOI of 10 for 18 h. They were then mixed with uninfected immature BMDCs labeled with the red fluorescent dye PKH-26 (Sigma-Aldrich Co.) for 4 h at either 37°C or 4°C. Phagocytosis of infected DCs by uninfected DCs (PKH-67⁺ PKH-26⁺ population) was visualized by flow cytometry.

BMDCs derived from either WT C57BL/6 mice (B6.DCs) or β2m^-/-mice (β2m^-/-DCs) were infected with rMVA-NP or control rMVA-GFP at a MOI of 10 for 6 h. The cells were then extensively washed with PBS and UV-irradiated to inactivate any residual viruses and prevent co-injection of infectious MVA. They were then injected (1 × 10⁶) into B6 mice that had been i.p.-infected with 2 × 10⁵ PFU LCMV 30 days earlier. Five days after the DC injection, spleens were harvested and NP_396–404-specific CTL activity was assessed by fluorescent cellular cytotoxicity assay as previously described 58. Briefly, target EL4 cells were incubated at 37°C 5% CO₂ for 1 h in the presence of 1 μM NP_396–404 peptide and 3 μM cell tracker orange (CTO, Invitrogen, Carlsbad, California). Cells were washed once with PBS and resuspended in CM at 1 × 10⁶/ml. Target cells and effector splenocytes from BMDC-immunized mice (100 μl) were co-cultured on 96-well U-bottom plates for 4 h at 37°C 5% CO₂. The cells were then pelleted and incubated in 75 μl/well fluorogenic caspase 3 substrate R2D2 (10 μM, kindly provided by Dr. Beverly Packard, OncoImmunin Inc., Gaithersburg, MD) for 30 min followed by two washes with cold PBS. Sample acquisition and analysis are as described above. The cleaved caspase substrate has the following fluorescence peak characteristics: λ_ex = 505 nm and λ_em = 530 nm, and can be detected in the FL1 channel. CTO can be detected in the FL2 channel. The percentage of specific killing = [% caspase⁺ CTO⁺ cells/(% caspase⁺ CTO⁺ cells + % caspase^- CTO⁺ cells)] × 100%.

All statistical analyses were made with Microsoft Excel software (Microsoft). Statistical comparisons between data sets were made with a two-tailed homoscedastic Student's t-test.