Type I IFN plays an important role in the activation of NK cells. However, the mechanism underlying type I IFN-dependent NK cell activation remains largely unknown. A recent report suggested that type I IFN acted on accessory dendritic cells, leading to IL-15 production, and that subsequent trans-presentation of IL-15 was required for NK cell activation upon stimulation with synthetic TLR ligands. It is not clear how type I IFN regulates NK cell activation in response to live pathogens. Using a murine model of infection with vaccinia virus (VV), we previously demonstrated a critical role for type I IFN in the innate immune control of VV infection. In this study, we first showed that type I IFN did not directly protect L929 cells from VV infection in vitro and that type I IFN-dependent innate immune control of VV infection in vivo was mediated by activated NK cells. We further demonstrated that direct action of type I IFN on NK cells, but not on dendritic cells, is required for the activation of NK cells in response to VV infection both in vitro and in vivo, leading to efficient VV clearance. Our findings may help design effective strategies for the control of poxviral infections in vivo.

Vaccinia virus (VV)3 represents one of the medically important viruses. It is a member of the poxvirus family, which includes smallpox (variola) virus, monkeypox virus, cowpox virus, and ectromelia virus, and has a large, complex dsDNA genome that replicates exclusively in the cytoplasm (1). VV is the most studied member of the poxvirus family and is the vaccine responsible for successful eradication of smallpox worldwide in the late 1970s (2). This unparalleled success is now being threatened by bioterrorists deliberately reintroducing smallpox, against which vaccination is no longer routine (3, 4, 5). The revival of smallpox vaccination has been countermanded by the relatively high incidence of adverse events associated with the currently used live VV vaccine (6, 7, 8). Thus, to effectively control poxviral infections, it is necessary to elucidate the host’s defense mechanism(s) against poxviruses in vivo.

Recovery from viral infections depends on the host’s ability to mount effective innate antiviral responses that can eliminate, or at least control, the invading pathogen. NK cells represent an important component of the innate immune system. It has been shown that NK cells play a critical role in innate immune defense against various viral infections in vivo (9). NK cells have also been implicated in the response to poxviruses. Upon poxviral infection, NK cells are activated, expand, and accumulate at the site of infection, and the activated NK cells are important for recovery of the infection (10, 11, 12, 13). However, it remains poorly understood how NK cell activation is regulated upon poxviral infection.

Type I IFNs, produced by host cells early after viral infection, represent a key player in antiviral defense (14). They are a family of cytokines that constitute 13 and 17 IFN-α subtypes in mice and humans, respectively, and one IFN-β in both species (15). All type I IFNs signal through a heterodimeric receptor composed of two subunits, IFN-αβ receptor 1 (IFNαβR1), and IFNαβR2. Stimulation of IFNαβR with type I IFNs triggers a series of signaling cascades leading to the transcription of >100 IFN-stimulated genes (14). The serine/threonine protein kinase and the 2′-5′ oligoadenylate synthetases, both of which are activated by viral dsRNA, are among the best characterized IFN-stimulated genes with antiviral activity through suppression of viral replication in infected cells by inhibiting RNA and protein synthesis (16, 17).

In addition to the direct antiviral effects, type I IFN also mediates a variety of immunoregulatory effects (14), including regulation of NK cell activation (18). How type I IFN regulates NK cell activation is yet to be fully elucidated. A recent study suggested that action of type I IFN on accessory dendritic cells (DCs), but not on NK cells was required for NK cell activation in response to synthetic TLR ligands (19). However, it remains unknown how type I IFN regulates NK cell activation in response to live pathogens. In a murine model of VV infection, we have recently shown that type I IFN play a critical role in the innate immune control of VV infection (20). In this study, we showed that type I IFN did not directly protect cells from VV infection in vitro and that type I IFN-dependent innate immune control of VV infection in vivo was mediated through the activation of NK cells. We further demonstrated that type I IFN signaling directly on NK cells was necessary for their activation and effector function both in vitro and in vivo.

IFNαβR−/− mice on 129/Sv background (H-2b) were obtained from B&K Universal, and their wild type (WT) control 129/Sv mice were purchased from Charles River Laboratories. Groups of 6- to 8-wk-old mice were selected for this study. All experiments involving the use of mice were done in accordance with protocols approved by the Animal Care and Use Committee of Duke University.

The Western Reserve strain of VV and encephalomyocardititis virus (EMCV) were purchased from American Type Culture Collection (ATCC). VV was grown in TK-143B cells (ATCC) and purified, and the titer was determined by plaque assay on TK-143B cells and stored at −80°C until use as described (20). EMCV was grown in L929 cells (ATCC) and purified by centrifugation, and the titer was determined by plaque assay on L929 cells as described (21).

L929 cells plated at 1 × 104 cells/100 μl were pretreated with 1 × 103 U/ml recombinant murine IFN-α (R&D Systems) for 24 h. After removal of the IFNα-containing supernatant, the cells were infected for 24 h with either VV or EMCV with a multiplicity of infection (MOI) of 62.5, 12.5, 2.5, 0.5, or 0.1. Survival was determined by FACS analysis 24 h later.

For depletion of NK cells in vivo, mice received 250 μg of anti-asialo GM1 antiserum (Wako Chemicals) injected i.v. 3 days prior and the day of infection with VV. Before infections, peripheral blood and splenic cells were analyzed to confirm elimination of DX5+CD3 NK cells.

FITC-conjugated anti-IFN-γ, PE-conjugated anti-DX5, and PE-Cy5-conjugated anti-CD3ε were purchased from BD Biosciences. FITC-conjugated anti-granzyme B and FITC-conjugated anti-perforin were purchased from eBioscience. To assess production of IFN-γ, granzyme B, and perforin intracellularly, splenocytes were incubated with 100 ng/ml PMA and 250 ng/ml ionomycin and 5 μg/ml Brefeldin A containing Golgi-plug (BD Biosciences) for 4 h at 37°C. Intracellular staining was performed as previously described (20). FACS Canto (BD Biosciences) was used for flow cytometry event collection, which was analyzed using FACS DiVA software (BD Biosciences).

NK cell cytotoxicity assay was performed as previously described (22). In brief, splenocytes were enriched for DX5+ NK cells by positive selection with PE-conjugated anti-DX5 and anti-PE microbeads (Miltenyi Biotec). DX5+ splenocytes were then incubated with 51Cr-labeled NK sensitive targets, YAC-1 cells (ATCC) at different E:T for 4 h at 37°C. The specific 51Cr release was calculated as (experimentalcpm − spontaneouscpm)/ (maximumcpm − spontaneouscpm) × 100.

Viral load in the ovaries was measured by plaque-forming assay as described (20). In brief, female mice were sacrificed 2 days after infection, and ovaries were harvested and stored at −80°C. Ovaries from individual mice were homogenized and freeze-thawed three times. Serial dilutions were performed on confluent TK-143B cells, and viral titers were then determined 2 days later by crystal violet staining.

DC were generated from the bone marrow cells in the presence of GM-CSF and IL-4 as described (20). In brief, bone marrow cells were harvested from femurs and tibiae of mice and cultured in the presence of mouse GM-CSF (1,000 U/ml) and IL-4 (500 U/ml) (R&D Systems) for 5 days. GM-CSF and IL-4 were replenished on days 2 and 4. On day 5, CD11c+ DCs were harvested for NK cell stimulation.

NK-DC coculture was performed as described with some modifications (23). In brief, DX5+CD3 NK cells were purified from splenocytes of naive 129/Sv mice via flow cytometry sorting on a FACS DiVA. NK cells (5 × 105) were cocultured with CD11c+ DC (2.5 × 105) at an NK:DC ratio of 2:1. The coculture was subsequently infected with VV with MOI of 1, for 48 h at 37°C.

DX5+CD3 NK cells were purified from splenocytes of naive WT or IFNαβR−/− 129/Sv mice via flow cytometry sorting. Two × 105 DX5+CD3 WT or IFNαβR−/− NK cells were administered i.v. to IFNαβR−/− recipients, which were subsequently injected i.p. with 1 × 107 PFU VV.

Results were expressed as mean ± SD. Differences between groups were examined for statistical significance using the Student’s t test.

To understand how type I IFN confers innate immune defense against VV infection in vivo, we first examined whether IFN-α interfered with the replication of VV in a permissive cell line, L929 cells. It has been shown that IFN-α pretreatment protects L929 from RNA viruses such as EMCV-induced cell death through inhibiting viral replication (24, 25). We found that infection of the L929 monolayer with EMCV at MOI as low as of 0.1 led to cell death for the majority of cells 2 days later, suggesting that cell death was dependent on viral replication and cell-to-cell spread in the monolayer (Fig. 1,A). Indeed, pretreatment with IFN-α resulted in a significant (p < 0.01) reduction in cell death even at a MOI of 62.5, confirming that IFN-α can directly protect cells from EMCV infection. However, under similar conditions, IFN-α pretreatment did not alter VV-induced cell death, suggesting that IFN-α did not prevent VV replication in L929 cells (Fig. 1 B). Thus, IFN-α does not directly protect cells against VV infection in vitro.

FIGURE 1.

Type I IFN pretreatment protects L929 cells from EMCV, but not VV, infection. L929 cells were cultured in the presence (+IFNα) or absence (−IFNα) of IFN-α for 24 h. The supernatant was then removed, and the cells were infected with either EMCV (A) or VV (B) at different MOI for another 48 h. The mean percentage of survival ± SD is indicated. Data shown are representative of three independent experiments.

FIGURE 1.

Type I IFN pretreatment protects L929 cells from EMCV, but not VV, infection. L929 cells were cultured in the presence (+IFNα) or absence (−IFNα) of IFN-α for 24 h. The supernatant was then removed, and the cells were infected with either EMCV (A) or VV (B) at different MOI for another 48 h. The mean percentage of survival ± SD is indicated. Data shown are representative of three independent experiments.

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Our observation that IFN-α did not directly protect cells against VV infection in vitro suggests that the type I IFN-dependent innate immune control of VV infection in vivo likely operates through a different mechanism. As type I IFNs have been shown to mediate immunoregulatory effects on NK cells in other models of infection (18) and NK cells have been implicated in innate immunity against poxviruses (10, 11, 12, 13), we hypothesized that type I IFN-dependent innate immunity against VV infection in vivo was mediated by activated NK cells. We first tested whether VV infection in WT mice led to NK cell activation. Forty-eight hours after infection with 1 × 107 PFU VV i.p., splenic NK cells expanded more than twice (Fig. 2, A and B), produced significantly (p < 0.001) higher amounts of effector molecules (Fig. 2,C), and demonstrated lytic function on NK-sensitive YAC-1 cells (Fig. 2,D) compared with the naive controls, confirming that indeed NK cells are activated upon VV infection. These activated NK cells were critical for VV clearance as WT mice depleted of NK cells (Fig. 3,A) showed a defect in NK lytic activity (Fig. 3,B) and had a significantly (p < 0.001) higher viral titer than the control mice (Fig. 3,C). We next determined whether activation of NK cells upon VV infection was regulated by type I IFN signaling. IFNαβR−/− mice were infected with 1 × 107 PFU VV i.p. and splenic NK cells were analyzed 48 h later. No significant expansion of NK cells was observed in IFNαβR−/− mice upon VV infection (Fig. 4,B). In addition, IFNαβR−/− NK cells failed to produce any significant amounts of effector molecules (Fig. 4,C) or lytic activity (Fig. 4,D) over the background levels in the naive mice, indicating that type I IFN was critical for NK cell activation in response to VV infection. We further observed that IFNαβR−/− mice and WT mice depleted of NK cells displayed similar levels of viral titer that was significantly (p < 0.001) elevated compared with the WT mice (Fig. 3 C). Collectively, our data support the conclusion that type I IFN-dependent innate immune control of VV infection in vivo is mediated through regulating NK cell function.

FIGURE 2.

Type I IFN signaling is required for NK cell activation in response to VV infection in vivo. Mice were infected with VV (+VV) or left uninfected (Naive). Splenocytes were harvested 48 h later and stained for NK cells with anti-DX5 and anti-CD3 Abs. A, The percentage of NK (DX5+CD3) cells among total lymphocytes from WT mice is indicated. B, The mean percentage ± SD of NK (DX5+CD3) cells among total lymphocytes from WT and IFNαβR−/− mice is indicated. C, Splenocytes from WT and IFNαβR−/− mice were assayed for intracellular granzyme B (GRB), perforin (PFN), and IFN-γ production by NK cells. The percentage of GRB, PFN, or IFN-γ positive cells among DX5+CD3 cells is indicated. D, Splenocytes were assayed for NK lytic activity on YAC-1 cells for 4 h at different E:T ratios. Naive splenocytes (Naive) were used as a control. The percentages of specific lysis are indicated. Data shown are representative of three independent experiments.

FIGURE 2.

Type I IFN signaling is required for NK cell activation in response to VV infection in vivo. Mice were infected with VV (+VV) or left uninfected (Naive). Splenocytes were harvested 48 h later and stained for NK cells with anti-DX5 and anti-CD3 Abs. A, The percentage of NK (DX5+CD3) cells among total lymphocytes from WT mice is indicated. B, The mean percentage ± SD of NK (DX5+CD3) cells among total lymphocytes from WT and IFNαβR−/− mice is indicated. C, Splenocytes from WT and IFNαβR−/− mice were assayed for intracellular granzyme B (GRB), perforin (PFN), and IFN-γ production by NK cells. The percentage of GRB, PFN, or IFN-γ positive cells among DX5+CD3 cells is indicated. D, Splenocytes were assayed for NK lytic activity on YAC-1 cells for 4 h at different E:T ratios. Naive splenocytes (Naive) were used as a control. The percentages of specific lysis are indicated. Data shown are representative of three independent experiments.

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FIGURE 3.

NK cells are required for efficient clearance of VV in vivo. A, Mice were treated with 250 μg of anti-asialo GM1 Ab on days 0 and 3 (anti-asialo GM1) or left untreated (Control). On day 5, splenocytes were stained with anti-DX5 and anti-CD3 Abs. The percentage of DX5+CD3 cells among total lymphocytes is indicated. B, Mice were depleted of NK cells on days 0 and 3 (anti-asialo GM1) or left untreated (Control), and infected with VV. Forty-eight hours later, splenocytes were assayed for NK lytic activity on YAC-1 cells for 4 h at different E:T ratios. Naive splenocytes (Naive) were used for comparison. The percentage of specific lysis is shown. C, Female WT or IFNαβR−/− mice were depleted of NK cells on days 0 and 3 (anti-asialo GM1) or left untreated (Control), followed by VV infection. After 48 h, their ovaries were harvested for measurement of viral load. Data represents viral titer ± SD as PFU per milliliter of cell lysate. Data shown is representative of three independent experiments.

FIGURE 3.

NK cells are required for efficient clearance of VV in vivo. A, Mice were treated with 250 μg of anti-asialo GM1 Ab on days 0 and 3 (anti-asialo GM1) or left untreated (Control). On day 5, splenocytes were stained with anti-DX5 and anti-CD3 Abs. The percentage of DX5+CD3 cells among total lymphocytes is indicated. B, Mice were depleted of NK cells on days 0 and 3 (anti-asialo GM1) or left untreated (Control), and infected with VV. Forty-eight hours later, splenocytes were assayed for NK lytic activity on YAC-1 cells for 4 h at different E:T ratios. Naive splenocytes (Naive) were used for comparison. The percentage of specific lysis is shown. C, Female WT or IFNαβR−/− mice were depleted of NK cells on days 0 and 3 (anti-asialo GM1) or left untreated (Control), followed by VV infection. After 48 h, their ovaries were harvested for measurement of viral load. Data represents viral titer ± SD as PFU per milliliter of cell lysate. Data shown is representative of three independent experiments.

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FIGURE 4.

Activation of NK cells by DCs upon VV infection in vitro. A, Purified DX5+CD3 NK cells were cocultured with CD11c+ DC and infected with VV (+VV) or left uninfected (Uninfected). NK cells were assayed for intracellular granzyme B (GRB), perforin (PFN), and IFN-γ 48 h later. The percentage of GRB, PFN, or IFN-γ positive cells among DX5+CD3 cells is indicated. Data shown are representative of three independent experiments.

FIGURE 4.

Activation of NK cells by DCs upon VV infection in vitro. A, Purified DX5+CD3 NK cells were cocultured with CD11c+ DC and infected with VV (+VV) or left uninfected (Uninfected). NK cells were assayed for intracellular granzyme B (GRB), perforin (PFN), and IFN-γ 48 h later. The percentage of GRB, PFN, or IFN-γ positive cells among DX5+CD3 cells is indicated. Data shown are representative of three independent experiments.

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We next investigated how type I IFN regulated the activation of NK cells upon VV infection. Conventional CD11c+ DCs have been shown to play a critical role in NK cell activation (19, 23). A recent report has suggested that type I IFN signaling through DCs may be important in the activation of NK cells in response to stimulation with various TLRs (19). To address whether the same is true in NK cell response to VV infection, we used an in vitro DC-NK cell coculture system. Purified DX5+CD3 NK cells were cocultured in vitro with conventional CD11c+ DCs generated from bone marrow cells in the presence of GM-CSF and IL-4, followed by infection with VV. Forty-eight hours after infection, NK cells produced much higher amounts of granzyme B, perforin, and IFN-γ compared with the uninfected control (Fig. 4). No activation of NK cells was observed when NK cells were stimulated alone with VV (data not shown), suggesting that DCs were also critical for NK cell activation upon VV infection. Because infection of DC with VV in vitro also elicits type I IFN production (20), we next examined whether type I IFNs acted on DCs or NK cells for NK cell activation in response to VV infection. To address this question, purified WT or IFNαβR−/− NK cells were cocultured with WT or IFNαβR−/− DCs, followed by infection with VV. The activation of NK cells was analyzed 48 h after infection. Our data showed that IFNαβR−/− DCs elicited similar amounts of granzyme B (Fig. 5, A and B), perforin (Fig. 5,C), and IFN-γ (Fig. 5,D) production by NK cells compared with the WT counterparts, suggesting NK cell activation is independent of type I IFN signaling on DCs in response to VV infection. In contrast, NK cell activation was severely compromised when IFNαβR−/− NK cells were used for stimulation (Fig. 5), indicating that direct action of type I IFNs on NK cells is required for their activation upon VV infection.

FIGURE 5.

NK cell activation upon VV is dependent on type I IFN signaling on NK cells in vitro. WT or IFNαβR−/− DX5+CD3 NK cells were cocultured with WT or IFNαβR−/− CD11c+ DC and infected with VV (+VV) or left uninfected (Uninfected). Forty-eight hours after infection, NK cells were assayed for intracellular granzyme B (GRB), perforin (PFN), and IFN-γ. A, The FACS plots of GRB is shown with the percentage of GRB positive cells among DX5+CD3 cells indicated. B–D, The percentage ± SD of GRB (B), PFN (C), or IFN-γ (D) positive cells among DX5+CD3 cells is shown. Data shown are representative of two independent experiments.

FIGURE 5.

NK cell activation upon VV is dependent on type I IFN signaling on NK cells in vitro. WT or IFNαβR−/− DX5+CD3 NK cells were cocultured with WT or IFNαβR−/− CD11c+ DC and infected with VV (+VV) or left uninfected (Uninfected). Forty-eight hours after infection, NK cells were assayed for intracellular granzyme B (GRB), perforin (PFN), and IFN-γ. A, The FACS plots of GRB is shown with the percentage of GRB positive cells among DX5+CD3 cells indicated. B–D, The percentage ± SD of GRB (B), PFN (C), or IFN-γ (D) positive cells among DX5+CD3 cells is shown. Data shown are representative of two independent experiments.

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We next sought to examine whether direct type I IFN signaling on NK cells was also required for NK cell activation upon VV infection in vivo. We have previously shown that IFNαβR−/− mice produce large amounts of type I IFN upon VV infection (20). Thus, if type I IFN acts directly on NK cells for their activation in vivo, adoptive transfer of WT NK cells into IFNαβR−/− mice should restore NK cell activation and result in a significant reduction of viral load. To address this question, DX5+CD3 NK cells were purified from the spleens of WT or IFNαβR−/− 129s/v mice by FACS sorting. Two × 105 WT or IFNαβR−/− NK cells were transferred into IFNαβR−/− mice i.v., which were subsequently infected i.p. with 1 × 107 PFU VV. After 48 h, the spleens and ovaries from these recipient mice were analyzed for NK cell activation and viral titer. In IFNαβR−/− mice reconstituted with WT NK cells, the production of granzyme B, perforin, and IFN-γ by splenic NK cells neared that in WT mice (data not shown). Furthermore, splenic NK cells harvested from IFNαβR−/− mice reconstituted with WT, but not IFNαβR−/−, NK cells were capable of lysing YAC-1 targets to a level equivalent to that of WT mice (Fig. 6,A). These data indicated that WT NK cells were functionally activated in otherwise IFNαβR−/− mice. When VV titer in the ovaries was assessed, IFNαβR−/− mice reconstituted with WT NK cells were able to clear VV in vivo similarly to WT mice, whereas IFNαβR−/− mice or IFNαβR−/− mice reconstituted with IFNαβR−/− NK cells failed to clear the virus (Fig. 6 B). Taken together, these data support the conclusion that direct action of type I IFN on NK cells is required for activation of NK cells in response to VV infection in vivo.

FIGURE 6.

Type I IFNs act directly on NK cells for their activation in response to VV infection in vivo. IFNαβR−/− mice were reconstituted with WT NK cells (+ WT NK) or IFNαβR−/− NK cells (+ IFNαβR−/− NK) followed by infection with VV. WT and IFNαβR−/− mice infected with VV were used as controls. A, Forty-eight hours later, splenocytes were assayed for NK lytic activity on YAC-1 cells at different E:T ratios. The percentage of specific lysis is indicated. B, The ovaries of female mice were harvested for measurement of viral load. Data represents viral titer ± SD as PFU per milliliter of cell lysate. Data are representative of two independent experiments.

FIGURE 6.

Type I IFNs act directly on NK cells for their activation in response to VV infection in vivo. IFNαβR−/− mice were reconstituted with WT NK cells (+ WT NK) or IFNαβR−/− NK cells (+ IFNαβR−/− NK) followed by infection with VV. WT and IFNαβR−/− mice infected with VV were used as controls. A, Forty-eight hours later, splenocytes were assayed for NK lytic activity on YAC-1 cells at different E:T ratios. The percentage of specific lysis is indicated. B, The ovaries of female mice were harvested for measurement of viral load. Data represents viral titer ± SD as PFU per milliliter of cell lysate. Data are representative of two independent experiments.

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In this study, we have presented evidence that type I IFN does not protect L929 cells directly from VV infection in vitro. In vivo, type I IFN-dependent innate immune clearance of VV infection is mediated by regulating NK cell activation. We have further shown that type I IFN acts directly on NK cells for their activation and effector function in response to VV infection both in vitro and in vivo.

The observation that type I IFN did not directly protect L929 cells against VV infection in vitro may not rule out completely the possibility of a direct role for type I IFNs in anti-viral defense in vivo. However, our data that mice defective for type I IFN and those depleted of NK cells showed similar levels of elevated viral titer, and that WT NK cells were sufficient to clear VV infection in IFNαβR−/− mice supports our conclusion that type I IFN-dependent NK cell activation is mainly responsible for innate immune defense against VV infection in vivo. This is in contrast to a previous report that IFN-mediated prophylaxis against VV or murine cytomegalovirus (MCMV) infections is NK cell independent (26). What contributes to the differences is not entirely clear, but might be related to the timing (before vs during the infection), the dose, and/or the source (exogenously administered vs endogenously induced upon VV infection) of type I IFNs.

Previous studies have demonstrated a critical role of NK cells in innate immune defense against viral infections (9). It has been shown that NK cell activation upon MCMV infection is mediated by NK cell activation receptor, Ly49H, which specifically recognizes the m157 gene product of MCMV (27, 28). Furthermore, a recent report has suggested a role of NKG2D-activating receptor in NK cell activation in response to MCMV infection (23). Consistent with previous observations (10, 11, 12, 13), we showed in this study that NK cells are activated upon VV infection, which is critical for innate immune defense against VV infection in vivo. How VV activates NK cells remains to be defined. Thus, it will be important to identify what component of VV is responsible for NK cell activation and the corresponding NK cell activation receptor. Identification of these will help in the design of effective NK cell-based strategies to control poxviral infections in vivo.

In addition to direct stimulation through NK cell receptors, the activation of NK cells is also regulated by cytokines, particularly type I IFN (18). It has been shown that type I IFN directly enhances NK cell cytotoxicity and induces IL-15 to promote NK cell proliferation during MCMV infection (29). We showed in this study that NK cell activation in response to VV infection is also critically dependent on type I IFN. We further demonstrated that this is achieved by direct action of type I IFN on NK cells, but not on accessory DCs. Our observation is in contrast to a recent report that type I IFN signaling on DCs may be important in the activation of NK cells in response to stimulation with various synthetic TLR ligands (19). The reasons for the discrepancy are not clear, but could be related to the agents (live pathogen vs synthetic TLR ligands) used for in vitro and in vivo NK cell activation. Indeed, NK activation upon viral infections is a more complex process, which involves pathogen-derived gene products, a NKG2D-activating receptor in addition to cytokines such as type I IFNs (9). In addition, the secretion of other cytokines upon stimulation with a TLR ligand vs VV would be quite different, which could influence the dependency of DC vs NK cells on type I IFN signals for NK cell activation.

In summary, we have demonstrated that type I IFN-dependent innate immune control of VV infection in vivo is mediated through activation of NK cells. We have further shown that direct action of type I IFN on NK cells is required for their activation and function upon VV infection. These results may suggest potential strategies for the control of poxviral infections in vivo.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported by National Institutes of Health Grants CA111807 and CA047741 (to Y.Y.), and an Alliance for Cancer Gene Therapy grant (to Y.Y.).

3

Abbreviations used in this paper: VV, vaccinia virus; IFNαβR1, IFN-αβ receptor 1; DC, dendritic cell; WT, wild type; EMCV, encephalomyocardititis virus; MOI, multiplicity of infection; MCMV, murine cytomegalovirus.

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