Amplifying IFN-γ Signaling in Dendritic Cells by CD11c-Specific Loss of SOCS1 Increases Innate Immunity to Infection while Decreasing Adaptive Immunity

Listeria monocytogenes is a ubiquitous Gram-positive facultative intracellular pathogen that is typically found in soil and food. We and other investigators have been developing live-attenuated L. monocytogenes–based vaccine platforms for application to cancer and infectious disease. The systemic infection model of listeriosis in mice has provided important insights into host–pathogen interactions and the adaptive immune response. A functional innate response and an adaptive immune response are critical for eradicating the pathogen (1–3). L. monocytogenes elicits a potent CD8⁺ T cell response in mice that is attributed to direct infection of dendritic cells (DCs) in the spleen and delivery of L. monocytogenes–associated Ag directly to the host cell cytosol (4, 5). CD8α⁺ DCs are the primary reservoir for live bacteria within the first few hours of systemic infection (6), and these cells play a critical role in priming L. monocytogenes–specific T cells (6, 7). These DCs are an early source of IL-12, which, in turn, induces IFN-γ release by NK, NKT, and T cells (8). Importantly, these inflammatory cytokines also elicit negative-feedback loops through regulatory proteins that limit cellular activation by these potent cytokines.

The suppressor of cytokine signaling (SOCS) family proteins (SOCS1–7 and CIS) are a group of structurally related proteins characterized by a central SH2 docking motif for interaction with tyrosine-phosphorylated proteins. SOCS1 is induced by cellular activation and serves as a negative-feedback mechanism for cytokines sharing the common γ-chain (IL-2, IL-4, IL-7, IL-15), IFN-α, IFN-γ, and IL-12. Although the SH2 domain targets the SOCS proteins to specific molecules within the JAK–STAT pathway, the SOCS-box functions as an E3 ubiquitin ligase, promoting degradation of the cytokine receptor complex. SOCS1-knockout (SOCS1^−/−) mice are normal at birth; however, they exhibit slow growth and die within 3 wk of birth, with activation of peripheral T cells, necrosis of the liver, and macrophage infiltration of major organs (9, 10). The neonatal defects exhibited by SOCS1^−/− mice appear to occur primarily as a result of unchecked IFN-γ signaling, because SOCS1^−/− mice that also lack the IFN-γ gene or the IFN-γ receptor gene avoid neonatal lethality (11). The major source of this IFN-γ has been shown to be T cells, because Rag^−/− mice do not display SOCS1^−/− lethality (12). SOCS1 is involved in the suppression of inflammation by regulating cytokine signaling in innate immune cells, including macrophages and DCs, as well as nonimmune cells. Deficiency of SOCS1 in macrophages was shown to result in hyperresponsiveness to LPS (13–16), and silencing SOCS1 in DCs was shown to enhance Ag presentation, T cell priming, lupus-like autoimmune diseases, and antitumor immunity (17, 18).

For these reasons, we hypothesized that SOCS1 knockout in DCs would be a means to increase Ag-specific T cell responses to L. monocytogenes–based vaccines and, therefore, potentiate their ability to generate therapeutic T cells targeting infectious diseases or cancer. We tested our primary hypothesis using Socs1^fl/fl mice crossed with mice expressing Cre recombinase under the control of the CD11c promoter. This resulted in a strain in which CD11c⁺ cells selectively lack SOCS1 expression and activity. DCs demonstrated prolonged signal transduction following IFN-γ stimulation; however, surprisingly, vaccination of mice in which DCs lack SOCS1 resulted in a deficient CD8⁺ T cell response to bacterial Ags. Although NK cells were also negatively affected in these mice, they were not responsible for the poor CD8⁺ T cell responses generated in these animals. Instead, infection in these mice led to an increase in the number of TNF-α and inducible NO synthase (iNOS)-producing DC (TipDCs) in the spleen, resulting in increased innate control of the bacterium. Our data demonstrate that blocking negative feedback of cytokine signaling via deletion of SOCS1 in DCs, rather than increasing CD8 T cell responses to a pathogen, instead suppressed T cell responses and increased innate control of bacterial infection.

To generate mice in which SOCS1 was specifically deleted in CD11c⁺ cells, CD11c-Cre-GFP–transgenic mice (19) were obtained from The Jackson Laboratory (stock number 007567; The Jackson Laboratory, Bar Harbor, ME) and bred with SOCS1^fl/fl mice (15) (generously provided by Dr. Yoshimura, Keio University). Five- to ten-week-old sex-matched Cre⁻SOCS1^fl/fl and CD11c⁻Cre⁺SOCS1^fl/fl littermates were used for all experiments. C57BL/6 mice, OT-1 mice bearing a TCR specific for the SIINFEKL epitope of OVA on a Rag1^−/− background, and Rag1^−/− mice were purchased from The Jackson Laboratory. Animal protocols were approved by the Earle A. Chiles Research Institute Institutional Animal Care and Use Committee (Animal Welfare Assurance Number A3913-01).

L. monocytogenes strains used for these studies, wild-type (wt) and ΔactA ActA-QV (ΔactA-QV, expressing the class I–restricted vaccinia virus–derived epitopes B8R_20–27, C4L_125–132, A42R_88–96, and K3L_6–15, in addition to OVA_257–264) (20), were grown to stationary phase in brain–heart infusion (BHI) broth, washed in PBS, and injected i.v. (retro-orbital route) in 200 μl total volume. Unless otherwise noted, the following infectious doses were used: for wt L. monocytogenes, 1 × 10⁴ for survival and infectious studies and 1 × 10⁵ for challenge and cell sorting, and for ΔactA-QV L. monocytogenes, 1 × 10⁵ for infection and immune response. When bacterial counts were determined in various cell types, mice were infected with 1 × 10⁵ CFU wt L. monocytogenes. At 15 h postinfection, spleens were harvested; half of each spleen was directly homogenized, and the other half was dissociated, and flow sorted for specific cell subpopulations. All spleen samples were lysed and plated on BHI plates to calculate bacterial CFU in the source material. Vaccinia virus WR expressing full-length chicken OVA was grown in HeLa cells and frozen. Thawed cell lysates were treated for 30 min with 1.25 μg/ml trypsin at 37°C. Virus was diluted in HBSS and injected i.p. as 1 × 10⁶ PFU in 200 μl. For noninfectious vaccination, mice were immunized i.p. with 5 μg of anti–DEC-205–OVA (generously provided by Celldex Therapeutics, Hampton, NJ) together with 25 μg of anti-CD40 (clone FGK4.5; Bio X Cell, West Lebanon, NH) in a total volume of 200 μl.

Bone marrow–derived DCs (BMDCs) were generated according to a standard protocol (21). Briefly, 2 × 10⁶ bone marrow cells were seeded per 100-mm Petri dish in RPMI 1640 supplemented with 10% FBS and 20 ng/ml recombinant murine GM-CSF (R&D Systems, Minneapolis, MN), with or without 10 ng/ml IL-4 (PeproTech, Rocky Hill, NJ). On day 3, 10 ml of RPMI 1640 medium containing 20 ng/ml murine GM-CSF was added to the plates. On day 6, half of the culture supernatant was collected and centrifuged, and the cell pellet was resuspended in 10 ml of fresh RPMI 1640 medium containing 20 ng/ml murine GM-CSF and returned to the original plate. When needed, 10 ng/ml IL-4 was added at the same time. In general, 7–8-d cultured BMDCs were used for the experiments, unless otherwise specified. Flow cytometric analysis showed that these DC subsets contained >90% CD11c⁺ cells (data not shown).

BMDCs (5 × 10⁴) were stimulated with 100 ng/ml LPS (InvivoGen, San Diego, CA), 10 ng/ml IFN-γ (R&D Systems), or a combination of both reagents for 18 h at 37°C. Supernatants were removed and used for cytokine analysis, and cells were washed and stained for flow cytometric analysis, as described. For Western blot analysis and RNA extraction, cells were lysed and processed as described below.

For in vitro Ag-presentation studies, BMDCs were plated in 96-well plates (Costar-Corning) at 5 × 10³ cells per well with anti–DEC-205–OVA, soluble Endo-Free OVA (InvivoGen), or OVA_257–264 (SIINFEKL) synthetic peptide for 45 min at 37°C in complete medium. BMDCs were washed three times and resuspended in 200 μl of complete medium containing 5 × 10⁴ CFSE-labeled OT-1 CD8⁺ T cells. Proliferation was analyzed after 65–72 h of culture by flow cytometry (22). For isolation of splenic CD11c⁺ cells, spleens were dissociated, and CD11c⁺ cells were purified by positive selection (EasySep Mouse CD11c Positive selection isolation kit; STEMCELL Technologies, Vancouver, BC, Canada) and purity check for flow cytometry. Each determination was performed in triplicate.

For RNA extraction and quantitative real-time PCR (qRT-PCR), BMDCs were plated in a six-well plate (2 × 10⁶ cells per well) and stimulated as described above. At 18 h, cells were harvested, and RNA was purified using QIAzol and an RNeasy Mini Kit (QIAGEN, Valencia, CA). DNase-treated RNA was used as template for cDNA synthesis using SuperScript III Reverse Transcriptase (Invitrogen, Carlsbad, CA), and qRT-PCR was performed using PowerUp SYBR Green Master Mix (Applied Biosystems, Foster City, CA) and the following primers: β-actin forward, 5′-CCCTGTGCTGCTCACCGA-3′ and β-actin reverse, 5′-ACAGTGTGGGTGACCCCGTC-3′ and SOCS1 forward, 5′-CACCTTCTTGGTGCGCG-3′ and SOCS1 reverse, 5′-AAGCCATCTTCACGCTGAGC-3′. Reactions were carried out and analyzed in a StepOnePlus Real-Time PCR system (Applied Biosystems). Fold change was expressed as 2^−ΔΔCt, where the internal control is the β-actin gene, and the gene of interest is SOCS1.

For Western blot analysis, cells were lysed in RIPA buffer in the presence of protease and phosphatase inhibitor (Thermo Fisher Scientific, Waltham, MA), denatured in SDS loading buffer containing 2-ME, electrophoresed on 10% SDS-PAGE gels, and transferred to a PVDF Membrane (EMD Millipore, Billerica, MA). Blocked blots were probed overnight at 4°C with anti-Stat1, anti–phospho-Stat1 (Tyr701) (#9171; both from Cell Signaling Technology, Danvers, MA), or anti–β-actin (A2228; Sigma-Aldrich, St. Louis, MO) primary Ab, followed by goat anti-rabbit HRP-conjugated secondary Ab (1:20,000; Sigma-Aldrich). Binding was detected using SuperSignal West Pico Chemiluminescent Substrate (Thermo Fisher Scientific), and images acquired with a FluorChem E System (ProteinSimple, San Jose, CA).

Fluorochrome-conjugated Abs specific for CD11c (clone N418), CD11b (clone M1/70), Ly-6C (clone HK1.4), Ly-6G (clone 1A8), MHC class II (MHCII) I-A/I-E (clone M5/114.15.2), CD90.1 (clone HIS51), CD3 (clone 17A2), iNOS (clone CXNFT), IL-12–p40 (clone C17.8), CD19 (clone eBio1D3), IL-2 (clone JES6-5H6), CD86 (clone GL1), CD27 (clone LG.7F9), NK1.1 (clone PK136), CD49b (clone DX5), NKp46 (clone 29A1.4), CD45.1 (clone A20), CD45.2 (clone 104), IFN-γ (clone XMG1.2), (eBioscience, San Diego, CA), CD4 (clone RM4-4), CD8α (clone 53-6.7), TNF (clone MP6-XT22) (BD Biosciences, San Jose, CA), and XCR1 (clone ZET) (BioLegend, San Diego, CA) were used at optimal titers.

Serum cytokines were determined using a BD Cytometric Bead Array (CBA) Mouse Inflammation Kit (BD Biosciences). Samples were acquired on an LSR II flow cytometer, and the exported data were analyzed using the CBA Analysis Add-in for Excel.

For analysis of T cell responses, spleens were dissociated and filtered through a 70-μm cell strainer. RBCs were lysed with RBC Lysing Buffer (Sigma-Aldrich). For peptide-stimulation assays, splenocytes were stimulated for 4 h with 1 μM OVA_257–264 (SIINFEKL), B8R_20–27, A42R_88–96, or LLO_190–201 peptide in the presence of brefeldin A (GolgiPlug; BD Biosciences). Peptides for stimulation were obtained from A&A Labs (San Diego, CA) and reconstituted in DMSO. Unstimulated controls (DMSO only) were used to assess nonspecific protein production for each animal. Cells were stained for surface Ags, fixed (Cytofix/Cytoperm buffer; BD Biosciences), and stored at −80°C (in Cytofix/Cytoperm buffer) until further analysis. For intracellular cytokine staining (ICS), frozen cells were thawed, permeabilized (Perm/Wash Buffer; BD Biosciences), and stained for intracellular IFN-γ. To assess TipDC/DC activation, splenocytes were processed as described above and incubated for 4 h at 37°C, 5% CO₂, with or without 10⁷ CFU/ml heat-killed L. monocytogenes. Cells were stained, and iNOS intracellular staining was performed as described above. Samples were acquired on an LSR II flow cytometer (BD Biosciences) and analyzed using FlowJo 10.2 (TreeStar, Ashland, OR).

Spleens were harvested from donor mice, and CD8⁺ or total T cells were purified by negative selection (EasySep Mouse CD8⁺ T Cell Isolation Kit and EasySep Mouse T Cell Isolation Kit; STEMCELL Technologies). Prior to adoptive transfer, cells were stained to confirm purity of CD8⁺ T cells and total T cells (>90%).

For in vivo Ag-presentation experiments, single-cell suspensions of purified OT-1 CD8⁺ T cells were stained with CFSE (Molecular Probes, Eugene, OR) for 10 min at 37°C. Reactions were stopped with cold PBS and resuspended in the desired volume. Mice were injected with CFSE-CD8⁺ T cells, and spleens were removed and processed after 3 d. Staining was performed as described, and analysis was conducted with FlowJo 10.2. Mitotic events were determined as described (22).

To evaluate the immune response, 10,000 purified OT-1 CD8⁺ T cells were transferred into Cre⁻SOCS1^fl/fl and CD11c⁻Cre⁺SOCS1^fl/fl mice 1 d prior to immunization with ΔactA-QV. Spleens were harvested and processed for flow cytometry 7 d later.

For reconstitution, 2 × 10⁷ purified total T cells from Cre⁻SOCS1^fl/fl or CD11c⁻Cre⁺SOCS1^fl/fl spleens were transfered to RAG1^−/− hosts and the following day they were infected with ΔactA-QV. Seven days after the immunization, spleens were removed and processed for staining.

NK cells were purified from mouse splenocytes using an EasySep Mouse NK Cell Isolation Kit (STEMCELL Technologies), as described by the manufacturer. Cell purity was >90%, as confirmed by flow cytometry. Cells were plated and stimulated with IL-18 (R&D Systems) and/or IL-12 (PeproTech) for 6 h before removing the supernatant for cytometric bead array analysis and staining the cells for ICS, as described above.

For in vivo NK cell depletion, 100 μg of anti-NK1.1 Ab (clone PK136; Bio X Cell) was injected i.p., and NK cell depletion was confirmed by analyzing blood samples 24 h later. Mice were primed with ΔactA-QV, serum IFN-γ levels were measured at 24 h postinfection, and spleens were removed and processed for ICS 7 d later, as described.

Data were analyzed and graphed using Prism (GraphPad, La Jolla, CA). Individual data sets were compared using the Student t test, and analysis across multiple groups was performed using ANOVA, with individual groups assessed using the Tukey comparison.

To achieve loss of SOCS1 in DCs, mice expressing Cre recombinase under the control of the CD11c promoter were bred with a floxed socs1 gene (SOCS1^fl/fl mice) to generate Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates for study. These mice were healthy, avoiding the postnatal lethality of SOCS1^−/− mice (9, 10); however, Cre⁺SOCS1^fl/fl mice exhibited some degree of splenomegaly, and females and males developed psoriatic symptoms at 2–3 mo and after 4–5 mo, respectively. To confirm cell-specific expression of Cre, we took advantage of bicistronic expression of GFP along with Cre under control of the CD11c promoter (19). Consistent with prior reports, GFP was clearly detected in CD11c⁺MHCII⁺ cells in the spleen, including the CD8α⁻ and CD8α⁺ subsets (Fig. 1A). CD11b⁺ cells lacking CD11c were GFP⁻, as were CD19⁺ B cells and CD3⁺ T cells of the CD4 and CD8 compartments. NK cell subsets can express CD11c; however, we could detect only weak GFP expression in these cells, primarily in the NK1.1⁺CD49b⁺ subset (Fig. 1A). To confirm loss of SOCS1, BMDCs derived from Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates were stimulated with IFN-γ, and the socs1 transcript was detected by qRT-PCR. Although Cre⁻SOCS1^fl/fl BMDCs had robustly increased expression of socs1 transcript following IFN-γ treatment, no socs1 transcript was detectable in Cre⁺SOCS1^fl/fl BMDCs after cytokine priming (Fig. 1B). To confirm functional loss of SOCS1 activity, STAT1 phosphorylation was measured in BMDCs following prolonged stimulation with LPS, IFN-γ, or both. BMDCs lacking SOCS1 activity exhibited increased IFN-γ–mediated STAT1 phosphorylation compared with control littermates (Fig. 1C). The increased response to IFN-γ had phenotypic consequences, because although untreated Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl BMDCs exhibited similar expression of costimulatory molecules, BMDCs lacking SOCS1 activity exhibited a greater upregulation of CD86 following IFN-γ treatment (Fig. 1D). Naive Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates exhibited similar proportions and absolute numbers of CD11c⁺MHCII⁺, CD11c⁺MHCII⁺CD8α⁺, and CD11c⁺MHCII⁺CD11b⁺CD8α⁻ splenic DCs (Supplemental Fig. 1A, 1B). However, SOCS1-deficient DCs showed higher basal expression levels of MHCII in CD11c⁺ cells (Fig. 1E) and higher basal levels of CD86 in CD11c⁺MHCII⁺CD8α⁺ and CD11c⁺MHCII⁺CD11b⁺CD8α⁻ DCs (Supplemental Fig. 1Ci, 1Cii). Furthermore, XCR1 expression levels were significantly higher in CD11c⁺MHCII⁺CD8α⁺ DCs when SOCS1 was ablated (Supplemental Fig. 1Ci, 1Cii). These data demonstrate that mice lacking SOCS1 are highly responsive to IFN-γ treatment and exhibit increased basal activation in vivo.

To determine whether vaccine-driven Ag-specific responses are improved in mice with DC-specific loss of SOCS1 expression, we used an L. monocytogenes–based vaccine. L. monocytogenes vaccines directly infect DCs in the spleen following systemic administration, and these DCs are required to initiate CD8⁺ T cell responses to bacterial Ags (4). Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates were vaccinated with ΔactA-QV L. monocytogenes expressing four well-characterized T cell epitopes, including the SIINFEKL epitope of OVA. Mice with wt SOCS1 expression exhibited a strong Ag-specific T cell response to vaccination, with IFN-γ⁺ SIINFEKL-specific CD8 T cells readily detectable in the spleen 7 d following vaccination; however, unexpectedly, mice with DCs lacking SOCS1 demonstrated a significantly lower CD8 T cell response to Ag (Fig. 2A). Similarly, CD8 T cell responses to the other epitopes in the vaccine, B8R and A42R, were significantly reduced in Cre⁺SOCS1^fl/fl mice (Fig. 2B). To determine whether the T cell response to other infectious agents was similarly compromised in these mice, mice were vaccinated with a recombinant vaccine strain of vaccinia virus expressing OVA. Again, mice with wt SOCS1 expression exhibited a strong response to OVA, whereas mice with DCs lacking SOCS1 demonstrated a significantly lower CD8⁺ T cell response to OVA (Fig. 2A), indicating that this failure is not unique to L. monocytogenes. To determine whether this response was specific to infectious agents, mice were vaccinated with anti–DEC-205–OVA and anti-CD40, which delivers OVA to splenic DCs and drives efficient cross-presentation to T cells to generate robust OVA-specific CD8⁺ T cell expansion (23). Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates exhibited equivalent responses to vaccination with anti–DEC-205–OVA and anti-CD40 (Fig. 2A), suggesting that DC function is intact when Ag is delivered via this route. Furthermore, the success of this vaccination strategy was not due to the activity of anti-CD40, because delivery of anti-CD40 to mice vaccinated with ΔactA-QV L. monocytogenes was not able to restore T cell responses in Cre⁺SOCS1^fl/fl mice (Supplemental Fig. 2). Interestingly, the CD4⁺ T cell response to Listeria-associated Ag was not diminished in Cre⁺SOCS1^fl/fl mice (Fig. 2C), suggesting that the failure in response was associated with intracellular Ags from the infectious agents that are cross-presented by DCs to CD8⁺ T cells. To exclude the possibility that CD8⁺ T cell function was broadly diminished in Cre⁺SOCS1^fl/fl mice, Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates were adoptively transferred with a low dose of TCR-transgenic CD8⁺ OT1 T cells, which are specific for the SIINFEKL epitope of OVA presented on H2K^b and have wt SOCS1 expression. Vaccination of these mice with ΔactA-QV L. monocytogenes resulted in significantly lower OT1 CD8⁺ T cell responses in mice whose DCs lack SOCS1 expression (Fig. 2D). To assess the function of T cells from Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl mice independently of DC function, T cells from Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates were adoptively transferred to Rag1^−/− mice with wt SOCS1 expression, and these mice were vaccinated with ΔactA-QV L. monocytogenes. Although the proportion of CD8⁺ T cells from Cre⁺SOCS1^fl/fl mice responding to Ags was slightly lower than CD8⁺ T cells from Cre⁻SOCS1^fl/fl mice, there was no difference in the number of responding T cells after vaccination (Fig. 2Ei, 2Eii). Again, the CD4⁺ T cell response to LLO was equivalent between strains (Fig. 2Eiii). These data demonstrate that, when DCs lack SOCS1 expression, Ag-specific CD8⁺ T cell responses to infectious agents are significantly decreased.

L. monocytogenes generates innate and adaptive immune responses that each contribute to clearance of the bacterium. To determine whether the innate response was altered in mice whose DCs lack SOCS1, we examined cytokine expression in the serum 24 h following infection with wt L. monocytogenes. In mice whose DCs lack SOCS1, there was a significantly lower expression of IFN-γ, CCL2, and IL-6 but a significantly higher level of TNF-α (Fig. 3A). Consistent with prior reports (24), infection with ΔactA L. monocytogenes did not result in IL-10 secretion in control mice or in mice lacking SOCS1 in DCs (data not shown). Because NK cells can be an important source of IFN-γ in early innate responses to infectious agents, including L. monocytogenes (8), and NK cells expressed low levels of Cre (Fig. 1A), we investigated the role of NK cells in Cre⁺SOCS1^fl/fl mice. NK cells were decreased in number and proportion in Cre⁺SOCS1^fl/fl mice (Fig. 3B), mostly as a result of decreased numbers and proportions of mature CD11b⁺CD27⁺ and CD11b⁺CD27⁻ NK cells (Fig. 3C). Following infection with L. monocytogenes, IFN-γ production by NK cells was significantly decreased in Cre⁺SOCS1^fl/fl mice (Fig. 3D), indicating a failure of NK cell activation in response to infection. However, NK cells isolated from Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates were equally able to produce IFN-γ in response to stimulation in vitro (Supplemental Fig. 3), suggesting that the defect was not intrinsic to the NK cell but rather their response to infection. To determine whether NK cells were required for T cell Ag-specific responses to infection, wt mice were depleted of NK cells prior to vaccination with ΔactA-QV L. monocytogenes using a depleting NK1.1 Ab (Fig. 3Ei, Supplemental Fig. 3). These mice demonstrated loss of NK cells and significantly decreased IFN-γ in the serum following infection, but Ag-specific T cell responses were unchanged (Fig. 3Eii). These data demonstrate that in vivo NK cell function is changed in Cre⁺SOCS1^fl/fl mice but that early NK cell production of IFN-γ is not required for CD8⁺ T cell responses and so does not explain the poor response in mice whose DCs lack SOCS1.

Infection of CD8α⁺ DCs is critical to generate Ag-specific responses following infection with L. monocytogenes. To determine whether this infection was deficient in Cre⁺SOCS1^fl/fl mice, we analyzed infection of a range of cell populations at early time points following in vivo infection with L. monocytogenes wt strain. Similar amounts of bacteria could be detected at this early time point in the spleen of Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates (Fig. 4Ai), and, as anticipated, the majority of L. monocytogenes was present in CD11c⁺CD8α⁺ DCs, with small amounts of the bacterium in CD11c⁺CD8α⁻ DCs and CD11b⁺ monocytes (Fig. 4Aii). The number of bacteria in DCs was not different between Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates (Fig. 4Aii), indicating that the bacteria are similarly infecting CD11c⁺CD8α⁺ DCs, despite a failure to generate T cell responses. To determine whether DCs from these animals have a defect in functional Ag presentation, BMDCs from Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates were pulsed with OVA, SIINFEKL peptide, or anti–DEC-205–OVA and then tested for their ability to stimulate proliferation of OT1 CD8⁺ T cells in vitro. In each case, OT1 CD8⁺ T cell proliferation was equivalent in each group (Fig. 4B). To assess Ag presentation in vivo, Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates were challenged with ΔactA-QV L. monocytogenes (which also contains the SIINFEKL epitope) 4, 3, 2, or 1 d prior to adoptive transfer of a high dose of OT1 CD8⁺ T cells and were assessed for OT1 CD8⁺ T cell proliferation 3 d later. OT1 CD8⁺ T cell proliferation was significantly decreased in Cre⁺SOCS1^fl/fl mice at each time point following infection (Fig. 4C), indicating a significantly decreased capacity to functionally present Ag in vivo. To determine whether this functional difference in the cross-presenting capacity of splenic DCs was true ex vivo, we sorted CD11c⁺ splenic DCs from Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates, pulsed them with OVA, SIINFEKL peptide, or anti–DEC-205–OVA, and tested their ability to stimulate proliferation of OT1 CD8⁺ T cells in vitro. Cre⁺SOCS1^fl/fl DCs showed a slightly increased capacity to stimulate OT1 proliferation following cross-presentation of OVA and were equally able to stimulate T cells with pulsed peptide and anti–DEC-205–OVA (Fig. 4D). These data demonstrate that DCs lacking SOCS1 are functional in classic in vitro assays of cross-presentation but dysfunctional in stimulating adaptive immunity following infection in vivo.

To determine whether the failure in mice lacking SOCS1 in DCs is limited to priming of naive T cell responses, Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates were vaccinated with anti–DEC-205–OVA plus anti-CD40 to generate an equivalent and functional in vivo response and challenged 21 d later with ΔactA-QV L. monocytogenes. Mice lacking SOCS1 in DCs exhibited a significantly lower CD8⁺ T cell response following rechallenge (Fig. 5A), indicating that CD8⁺ T cell memory expansion is also impaired following L. monocytogenes rechallenge. To determine whether a failure in CD8 T cell responses impacted bacterial clearance, naive mice or mice that had been vaccinated with ΔactA-QV L. monocytogenes 21 d prior were challenged with 1 LD₅₀ (1 × 10⁵ CFU) wt L. monocytogenes, and bacterial load in the spleen was determined 3 d later. Cre⁻SOCS1^fl/fl mice that were vaccinated demonstrated significantly improved bacterial clearance compared with naive mice (Fig. 5B). In contrast, Cre⁺SOCS1^fl/fl littermates were not protected by vaccination (Fig. 5B). However, surprisingly, naive mice lacking SOCS1 in DCs appear to exhibit improved innate control of the infection at 3 d postinfection compared with mice with normal SOCS1 expression (Fig. 5B). Daily analysis of bacterial load in infected mice demonstrated that wt L. monocytogenes was able to infect and replicate in Cre⁺SOCS1^fl/fl mice, and although bacterial counts are initially lower, the mice are similarly susceptible to progressive infection as control mice (Supplemental Fig. 4A). Similarly, ΔactA-QV L. monocytogenes showed initially lower bacterial counts in the spleen and livers of Cre⁺SOCS1^fl/fl mice but similar clearance of the attenuated ΔactA-QV L. monocytogenes strain by day 5 postinfection (Supplemental Fig. 4B) Thus, the differing T cell response to infection cannot be adequately explained by differential clearance or persistence of bacteria in mice. To examine cell populations that may be participating in the innate clearance of L. monocytogenes, Cre⁻SOCS1^fl/fl and Cre⁺SOCS1^fl/fl littermates were analyzed for their monocyte and neutrophil populations. Naive Cre⁺SOCS1^fl/fl mice have higher numbers of CD11b⁺Ly6C^hi monocytes in the spleen than Cre⁻SOCS1^fl/fl mice and equivalent numbers of Ly6G^hiLy6C^hi neutrophils (Fig. 5Ci, 5Cii). Monocytes have been shown to be recruited into the spleen of infected mice and differentiate into TipDCs, which participate in innate bacterial clearance (25). As we demonstrated, wt L. monocytogenes–infected Cre⁺SOCS1^fl/fl mice display elevated TNF-α compared with control littermates (Fig. 3A), suggestive of increased TipDC activity. To determine whether there was increased TipDC activity, iNOS expression was measured in splenic cells following infection. We observed increases in TipDCs over time following infection with wt L. monocytogenes, and the number of these cells was significantly elevated in Cre⁺SOCS1^fl/fl mice compared with control littermates (Fig. 5D). These data suggest that mice lacking SOCS1 in DCs exhibit a dual phenotype. These mice generate an increased innate response associated with increased induction of iNOS in TipDCs but an impaired T cell response due to poor expansion of CD8⁺ T cells.

We demonstrate that SOCS1 deficiency in CD11c⁺ cells results in poor activation of CD8 T cell responses to Ags in bacterial and viral vaccines. This deficiency is caused by poor expansion of CD8 T cells by the critical CD8α⁺ DC population. Prior publications have described improved Ag presentation in DCs with SOCS1 deficiency; however, these experiments were performed with ex vivo–derived BMDCs and not in an intact animal (18, 26). We demonstrate for the first time, to our knowledge, that restricted cell-specific loss of SOCS1 in DCs in vivo redirects the immune response following L. monocytogenes infection away from an adaptive response and toward an innate response. We found that SOCS1 deficiency causes an increase in TNF-α secretion in serum, as well as an increase in iNOS⁺CD11b^intLy6C^highCD11c^intMHCII⁺ TipDCs in spleens, during the infection that could improve innate, rather than adaptive, control of infection.

L. monocytogenes LLO–mediated entry into the cytoplasm of DCs is required for efficient cross-presentation to CD8 T cells (27). L. monocytogenes produces cyclic dinucleotides, which are critical for bacterial function (28) but also activate the cytoplasmic sensor stimulator of IFN genes (STING), resulting in type I IFN production (29). Archer et al. (30) demonstrated that overactivation of STING can result in excess IFN production that limits CD8 T cell responses to L. monocytogenes. Similarly, overactivation of STING using exogenous administration of cyclic dinucleotides has resulted in type I IFN–mediated suppression of CD8 T cell responses (31). However, despite decreased adaptive immune responses, L. monocytogenes that overactivates STING does not have increased virulence and, in fact, can exhibit decreased virulence in vivo (32). This would be consistent with our data demonstrating an increased innate control of infection concomitant with decreased adaptive responses. TNF-α and iNOS are essential for defense against infection with L. monocytogenes (25, 33, 34). In addition, IFN-γ secretion from NK cells has been described as crucial for activation of monocytes to differentiate into TipDCs (8). Although the levels of NK cell–derived IFN-γ are low in mice lacking SOCS1 in DCs during the first 24 h postinfection, it may be sufficient to induce iNOS production due to the higher sensitivity to IFN-γ (17, 18). Thus, the balance of innate and adaptive responses to L. monocytogenes infection can be varied and still control the infection; however, strong protective T cell–mediated immunity requires carefully controlled inflammation at challenge (35, 36).

The CD8a⁺ subpopulation of DCs is critical to generate CD8 T cell responses via cross-presentation of viral- and bacterial-associated Ags (4, 5, 37). This same DC population is required for cross-presentation of cell-associated Ags (38); thus, these cells direct adaptive immune responses to cancer (39). Importantly, overexuberant inflammatory responses have similarly been shown to diminish immune control of cancer through “rebound immune suppression” (40). Thus, as with bacterial and viral vaccination approaches, strategies that aim to generate CD8 T cell–mediated immunity to tumors may similarly need to avoid overactivation of cross-presenting DCs to optimize the adaptive immune response.

The authors have no financial conflicts of interest.