Dendritic cells (DCs) encompass a heterogeneous population of cells capable of orchestrating innate and adaptive immune responses. The ability of DCs to act as professional APCs has been the foundation for the development and use of these cells as vaccines in cancer immunotherapy. DCs are also endowed with the nonconventional property of directly killing tumor cells. The current study investigates the regulation of murine DC cytotoxic function by T lymphocytes. We provide evidence that CD4+ Th-1, but not Th-2, Th-17 cells, or regulatory T cells, are capable of inducing DC cytotoxic function. IFN-γ was identified as the major factor responsible for Th-1–induced DC tumoricidal activity. Tumor cell killing mediated by Th-1–activated killer DCs was dependent on inducible NO synthase expression and NO production. Importantly, Th-1–activated killer DCs were capable of presenting the acquired Ags from the killed tumor cells to T lymphocytes in vitro or in vivo. These observations offer new possibilities for the application of killer DCs in cancer immunotherapy.

The cardinal property of dendritic cells (DCs) to function as professional APCs capable of orchestrating adaptive and innate immunity has been the basis for their implementation in vaccination strategies against cancer (13). However, to date, the limited efficacy of DC-based immunization in clinic has failed to spark significant enthusiasm. One possible limitation of this therapeutic approach may stem from the lack of mobilization of the full antitumoral potential of these cells. Indeed, the field has primarily focused on exploiting the APC function of DCs, with limited consideration given to their relatively recently described potential as direct tumor cell killers.

Multiple lines of evidence have indicated that DCs, when appropriately stimulated, can acquire cytotoxic properties against tumor cells, but not normal cells (46). The effector mechanisms underlying DC-mediated tumoricidal function are still being explored and may differ depending on the DC subtype (7). They may include the perforin/granzyme system (8), FasL (913), TNF-family members (TNF-α, TRAIL) (6, 8, 9, 1425), reactive oxygen species, and/or NO (4, 5, 11, 2628). Similarly, multiple modalities for the induction of DC cytotoxic function have been described, including CD40L (18), IFNs (4, 6, 14, 17, 29, 30), LPS, and other TLR agonists (4, 5, 8, 18, 31, 32). We have previously documented in the mouse (5), rat (4), and human (33) that ex vivo generated DCs activated with LPS are capable of killing tumor cells by NO, peroxinitrites, or reactive oxygen species-dependent mechanisms. However, whether DC killing function may be regulated by other immune cells has not been investigated.

In the current study, we show that mouse bone marrow-derived DC tumoricidal activity can be induced by CD4+ Th-1 lymphocytes. The mechanism of induction of killer DC (KDC) cytotoxic activity was not dependent on cell-to-cell contact. Using DCs generated from IFN-γ receptor knockout mice and IFN-γ blocking Abs, we identified IFN-γ as the primary factor responsible for Th-1–mediated induction of DC cytotoxic activity. Killing of tumor cells by Th-1–activated cytotoxic DCs (Th-1 KDCs) required NO production, but not perforin/granzyme or members of the death receptor ligand family. Importantly, Th-1 KDCs efficiently presented Ags derived from the tumor cells they had killed in vitro. Of therapeutic relevance, Th-1 KDCs injected intratumorally migrated to draining lymph nodes, where they were capable of presenting the acquired specific Ags. Thus, IFN-γ–secreting Th-1 lymphocytes promote the nonconventional cytotoxic activity of DCs, which can function as efficient APCs following killing of tumor cells both in vitro and in vivo.

Mice were housed under specific pathogen-free conditions and cared for according to the guidelines of the University of Arizona Institutional Animal Care and Use Committee. Six- to 8-wk-old BALB/c (H2d) and C57BL6 (H2b) mice were obtained from the National Cancer Institute. Inducible NO synthase−/− (iNOS−/−), (B6.129P2-Nos2tm1Lau/J), FasL−/− (B6Smn.C3- Faslgld/J), perforin−/− (CByJ.B6-Prf1tm1Sdz/J), IFNγR−/− (B6.129S7-Ifngr1tm1Agt/J), CD11cDTRGFP [B6.FVB-Tg(Itgax-DTR/EGFP)57Lan/J], OT-I, and OT-II mice were obtained from the Jackson Laboratory (Bar Harbor, ME).

The mouse melanoma cell line B16 was obtained from the American Tissue and Cell Collection. OVA-expressing B16 (B16-OVA) were obtained as reported (5). Mammary carcinoma tumor 4T1 were obtained from the American Tissue and Cell Collection. Cells were cultured at 37°C and 10% CO2 in RPMI 1640 media (Thermo Fisher Scientific, Waltham, MA) containing 10% heat-inactivated FBS (Thermo Fisher Scientific) and supplemented with 2 mM glutamine, 100 U/ml penicillin, 100 μg/ml streptomycin sulfate, 0.025 μg/ml amphotericin B, 0.5 × MEM nonessential amino acids, and 1 mM sodium pyruvate (complete media, CM). Cells were tested routinely and found to be free of Mycoplasma contamination.

NG-methyl-l-arginine (NMMA), LPS, and crystal violet were purchased from Sigma-Aldrich (St. Louis, MO). Murine IL-2, IL-4, IL-6, IL-12, and TGF-β were obtained from Peprotech (Rocky Hill, NJ). Anti–IFN-γ, anti–IL-4, and isotype control Abs were obtained from eBioscience (San Diego, CA).

DCs were generated from mouse bone marrow according to our previously reported procedures (4, 5, 28, 34). Briefly, total bone marrow cells were isolated from mouse femurs and tibias. Red cells were lysed in Pharm Lyse (BD Biosciences, San Jose, CA), and the cell suspension was passed through a 100-μm filter. Cells (5 × 105/ml) were seeded in six-well plates (3 ml/well) in RPMI 1640 medium (Thermo Fisher Scientific) supplemented with 10% heat-inactivated FBS (Thermo Fisher Scientific) and GM-CSF and IL-4 (Peprotech) (10 ng/ml each) and were incubated in 5% CO2 at 37°C. At 3 and 5 d after the beginning of the culture, the medium was replaced. On day 6, CD11c+ cells were selected from the culture, using anti-CD11c microbeads (Miltenyi Biotec, Auburn, CA) and cultured for an additional 2 d with GM-CSF and IL-4. The phenotypical characteristics of the obtained cells post CD11c+ selection after 6 d of culture are depicted in Supplemental Fig. 1. DC cultures did not contain conventional cytotoxic immune cells (Supplemental Fig. 1). T and B lymphocytes and NK cells represented less than 1.5% of the cells.

DC cytotoxic function was assessed as we previously reported (4, 5). Purified CD11c+ DCs were pretreated or not with IFN-γ (5 ng/ml) or with T lymphocyte culture supernatant, from day 6 to day 8 or as indicated. DCs were then washed and plated with B16 melanoma or 4T1 carcinoma cells (tumor cells/DC ratio = 1:5). DCs activated with LPS (1 μg/ml) were used as a positive control. Tumor cell killing was then evaluated as previously described (4, 5, 3537). Briefly, the cells were rinsed with PBS, and the remaining adherent cells were fixed with 95% ethanol and stained with crystal violet dye (100 μl in each well of a 96-well plate) for less than 10 s. The wells were then extensively washed with water. The dye was next eluted with acetic acid (30%). The amount of dye resuspended in the well is proportionate to the number of viable tumor cells. Plates were then read at 570 nm. Data were presented as the percentage of relative absorbance calculated from the following formula: Atest/Acontrol, where Atest is the absorbance of tumor cells cultured with DCs in different conditions and Acontrol is the absorbance of tumor cells cultured alone. DCs are very poorly stained with the dye and minimally contribute to detected absorbance (4, 5, 3537).

Culture supernatants were collected and incubated (50 μl) with an equal volume of Griess reagent. After incubation (15 min) at room temperature, the absorbance was read at 550 nm against 690 nm, following the manufacturer’s instructions (Premege, Madison, WI), and as previously reported (5).

Cells (∼106) were washed in PBS and were first incubated with an Fc receptor blocking Ab (BD Biosciences, Franklin Lakes, NJ) for 10 min and subsequently stained with saturating amounts of the appropriate fluorochrome-conjugated Abs for 30 min. For intracellular staining, cells were fixed and permeabilized according to the manufacturer’s instructions (eBioscience) and stained with the indicated Abs for transcription factor expression detection. Cells were then washed and analyzed using a FACSCalibur (Becton Dickinson Immunocytometry Systems, San Jose, CA). A minimum of 10,000 events were collected for each sample, and data analysis was performed using FlowJo software (Treestar, Ashland, OR) or CellQwest Pro 6.0 (Becton Dickinson). The following Abs were purchased from eBioscience (San Diego, CA): anti-CD4 FITC (GK1.5), anti-Tbet PE (eBio4B10), anti-GATA3 PE (TWAJ), anti-RORγt PE (AFKJS-9), anti-Foxp3 APC (FJK-16S), anti-F4/80 FITC (BM8), anti-CD11c FITC (N418), CD49b FITC (DX5), anti-CD86 FITC (GL1), MHC II FITC (NIMR-4), MHC I PE (SF1-1.1.1), and B220 PE (RA3-6B2); and from BD: CD3 FITC (145-2C11), CD40 FITC (HM40-3), anti–H-2Kb-SIINFEKL (eBio25D1.16) and CD11b PE (M1-70).

To generate Th lymphocyte lineages, CD4+CD25-CD62L+ naive T cells were isolated from mouse spleens using a CD4+CD62L+ T cell isolation kit (Miltenyi Biotec). The purified cells were then incubated for 3 d in 5% CO2 at 37°C in RPMI 1640 medium (Thermo Fisher Scientific) supplemented with 10% heat-inactivated FBS (Thermo Fisher Scientific) with anti-CD3 anti-CD28 coated T cell expander beads (1:1 bead/cell ratio) (Invitrogen, Carlsbad, CA) and with the following specific cytokines and blocking Abs: For Th-1 generation, cells were cultured with IL-2 (20 IU/ml), IL-12 (10 ng/ml), and anti–IL-4 blocking Ab (5 μg/ml). For Th-2 differentiation, cells were incubated with IL-2 (20 IU/ml), IL-4 (30 ng/ml), and anti–IFN-γ blocking Ab (5 μg/ml). Th-17 lymphocytes were generated with TGF-β1 (5 ng/ml), IL-6 (50 ng/ml), anti–IFN-γ and anti–IL-4 blocking Abs (5 μg/ml each). Regulatory T cells (Tregs) were generated by culturing isolated naive T cells in the presence of IL-2 (20 IU/ml) and TGF-β1 (5 ng/ml). All cytokines were purchased from Peprotech and blocking Abs from eBioscience. T lymphocytes were washed and restimulated with anti-CD3 anti-CD28 coated T cell expander beads for 8 h, and their supernatants were collected to activate DCs as described above.

Real-time RT-PCR was used to evaluate expression of different cytokines, transcription factors, or chemokine receptors in different T cell lineages and in DCs treated with LPS or IFN-γ. Total RNA was isolated using TRIzol reagent (Invitrogen), and its integrity was confirmed by denaturing agarose gel electrophoresis and calculated densitometric 28S/18S ratio. Then 250 ng total RNA was reverse transcribed using the iScript cDNA Synthesis Kit (Bio-Rad, Hercules, CA). Subsequently, 20 μl of the PCR reactions was set up in 96-well plates containing 10 μl 2× IQ Supermix (Bio-Rad), 1 μl TaqMan primer/probe set (ABI, Foster City, CA), 2 μl of the cDNA synthesis reaction (10% of RT reaction), and 7 μl nuclease-free water. Reactions were run and analyzed on a Bio-Rad iCycler iQ real-time PCR detection system. Cycling parameters were determined, and resulting data were analyzed by using the comparative Ct method as means of relative quantification, normalized to an endogenous reference (TATA box binding protein) and relative to a calibrator (normalized Ct value obtained from control mice) and expressed as 2−ΔΔCt (Applied Biosystems User Bulletin #2: Rev B “Relative Quantification of Gene Expression”).

DCs were isolated, cultured, and activated with LPS, IFN-γ, or Th-1 supernatant, as described above. Cells were collected and centrifuged at 350 g for 5 min at 4°C. Cells were then rinsed with cold PBS and were lysed with radioimmunoprecipitation assay buffer plus a protease and phosphatase inhibitor mixture (Thermo Scientific, Rockford, IL). The cell lysate was briefly sonicated and centrifuged at 8000 g for 8 min at 4°C. Protein concentration was determined using the BCA Protein Assay kit (Thermo Scientific). Next, 30 μg total cell lysate was boiled for 5 min and resolved in 10% Tris-Glycine Gel (Invitrogen) by electrophoresis at 125 V for 105 min. Proteins were then transferred to a polyvinylidene difluoride membrane (Millipore, Billerica, MA). The membrane was blocked with 5% milk in TBST containing 0.1% Tween 20 for 30 min before overnight incubation at 4°C with the indicated primary Abs. The blot was rinsed with TBST and incubated for 2 h at room temperature with secondary Ab. Reactive bands were visualized by exposure to film, using SuperSignal Chemiluminescent Substrate (Thermo Scientific). Abs against iNOS were purchased from Cell Signaling Technology (Beverly, MA). Abs against Actin were purchased from Sigma-Aldrich. The secondary Abs, peroxidase conjugated goat anti-rabbit and peroxidase conjugated goat anti-mouse, were purchased from Jackson ImmunoResearch (West Grove, PA).

The concentrations of IFN-γ, IL-10, and IL-17 in T lymphocyte culture supernatants were determined using ELISA kits according to the manufacturer’s procedures (eBioscience).

DCs were activated or not with the supernatant of Th-1 lymphocytes, washed, and cocultured for 48 h with B16 or B16-OVA tumor cells. Positive controls consisted of LPS-activated KDCs (5). At the end of the culture, DCs were selected using anti–CD11c-microbeads and were cocultured with B3Z cells (DC/B3Z ratio = 1:10). B3Z is a mouse CD8+ T-cell hybridoma that contains an Escherichia coli lacZ reporter gene driven by NF of activated T cell elements from the IL-2 promoter. The specific recognition of the SIINFEKL peptide of OVA (OVA257–264) in the context of MHC class I by the TCR of B3Z results in the expression of the enzyme β-galactosidase. The activity of this enzyme is detected by evaluating the subsequent conversion of a chemoluminescent substrate measured by luminometry (Novagen, Madison, WI), as previously documented (5).

CD11c-DTR mice (H2b) [B6.FVB-Tg(Itgax-DTR/EGFP)57Lan/J] were injected with 1 × 106 B16 or B16-OVA tumor cells s.c. in both flanks. When tumors were palpable, diphtheria toxin (DT; 5 ng/g body weight) was administered (i.p. injection) on 2 d consecutively (Sigma-Aldrich). This resulted in a significant depletion of endogenous DCs, as verified by flow cytometry (Supplemental Fig. 4A). On the day of the second DT injection, 20 × 106 day 8 DCs, previously activated with LPS, IFN-γ, or Th-1 supernatant, were injected intratumorally. At 36 h later, mice were euthanized, and CD11c+ cells were reisolated from the draining lymph nodes using CD11c microbeads (Miltenyi Biotec). Approximately 2 × 105 to 2.5 × 105 CD11c+ cells can be recovered. The ability of these reisolated DCs to induce B3Z activation was assessed, as explained in the previous section. In other experiments, the capability of reisolated CD11c+ DCs to induce MHC-I–restricted proliferation of OVA-specific OT-I or MHC-II–restricted proliferation of OVA-specific OT-II lymphocytes was determined using BrdU incorporation assays (Millipore), as previously reported (38).

Unless specified otherwise, all experiments were reproduced three times and performed in triplicate. A two-sided Student t test with paired samples was used to determine significant differences (p < 0.05) between groups. For real-time PCR experiments, statistical significance was determined by ANOVA, followed by the Fisher protected least significant difference post hoc test with StatView software package v.4.53 (SAS Institute, Cary, NC). Data are expressed as mean ± SD of mean.

The cytotoxic function of ex vivo generated DCs may be triggered by different approaches. We and others have previously reported on the induction of the tumoricidal activity of these cells by the TLR-4 ligand LPS (4, 5, 28, 39). However, the possible modulation of DC cytotoxic potential by T lymphocytes had not been elucidated. To assess whether DC killing function may be regulated by helper T cells, day 6 CD11c+ bone marrow-derived DCs were incubated for 48 h with anti-CD3– and anti-CD28–activated CD4+ T lymphocytes. DCs were then separated from T cells, using CD11c microbeads, and their ability to kill tumor cells was determined as previously described (4, 5). The results depicted in Fig. 1A indicate that CD4+ T cells triggered the ability of DCs to kill 4T1 mammary carcinoma cells. Induction of DC killing function was dependent on the DC/T cell ratio, with a significant cytotoxic activity obtained at a 1:5 ratio (Fig. 1A). Identical results were obtained using B16 melanoma as targets (Fig. 1B). Similar to our previous reports (5), DC-mediated tumor cell killing depended on the effector DC/target tumor cell ratio and was prominent at 5:1 E/T ratios (data not shown). The induction of DC cytotoxic activity did not require direct cell to cell contact and therefore involved soluble factor or factors produced by activated CD4+ T lymphocytes (Fig. 1C, 1D).

FIGURE 1.

Activated CD4+ T cells induce DC tumoricidal activity. A and B, Day 6 CD11c+ DCs were cultured for 48 h with activated CD4+ T lymphocytes at the indicated ratios. CD11c+ DCs were then separated from T cells, washed, and subsequently incubated for an additional 48 h with 4T1 (A) or B16 (B) tumor cells. Tumor cell survival was determined using a crystal violet assay. LPS-activated (1 μg/ml) DCs ([DC] LPS) were used as positive controls and untreated DCs (Untreated DCs) as negative controls. The data represent the mean ± SD from triplicate wells. Significant difference when compared with tumor cells cultured with control untreated DCs, *p < 0.001. C and D, 4T1 (C) or B16 (D) tumor cells were cultured for 48 h with untreated DCs (Untreated DCs), with LPS-activated (1 μg/ml) KDCs ([DC] LPS), or with DCs that had first been cocultured for 48 h and separated from activated CD4+ T cells by a microporous membrane ([DC] CD4 (TW)), as described in 1Materials and Methods. Tumor cell viability was determined as described above. Mean ± SD of triplicate cultures, *p < 0.0001.

FIGURE 1.

Activated CD4+ T cells induce DC tumoricidal activity. A and B, Day 6 CD11c+ DCs were cultured for 48 h with activated CD4+ T lymphocytes at the indicated ratios. CD11c+ DCs were then separated from T cells, washed, and subsequently incubated for an additional 48 h with 4T1 (A) or B16 (B) tumor cells. Tumor cell survival was determined using a crystal violet assay. LPS-activated (1 μg/ml) DCs ([DC] LPS) were used as positive controls and untreated DCs (Untreated DCs) as negative controls. The data represent the mean ± SD from triplicate wells. Significant difference when compared with tumor cells cultured with control untreated DCs, *p < 0.001. C and D, 4T1 (C) or B16 (D) tumor cells were cultured for 48 h with untreated DCs (Untreated DCs), with LPS-activated (1 μg/ml) KDCs ([DC] LPS), or with DCs that had first been cocultured for 48 h and separated from activated CD4+ T cells by a microporous membrane ([DC] CD4 (TW)), as described in 1Materials and Methods. Tumor cell viability was determined as described above. Mean ± SD of triplicate cultures, *p < 0.0001.

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CD4+ T lymphocytes represent a heterogeneous cell population composed of functionally and phenotypically distinct subsets primarily characterized as Th-1, Th-2, IL-17–producing Th-17, and immunosuppressive Foxp3+ Tregs (40). We further evaluated the respective contribution of these CD4+ T cell lineages to the induction of DC tumoricidal function. Naive T cells were cultured for 3 d in different conditions to generate Th-1, Th-2, Th-17, and Treg lymphocytes, as described in 1Materials and Methods. Cell lineage identity was confirmed by flow cytometry analysis of specific transcription factor expression (Tbet for Th-1, GATA3 for Th-2, RORγt for Th-17, and Foxp3 for Treg) (Fig.2A) and by ELISA and real-time PCR for detection of specific cytokine production (IFN-γ, IL-10, IL-17) (Fig. 2B, 2C).

FIGURE 2.

Generation and characterization of Th lymphocyte lineages. Splenic CD4+CD25CD62L+ T cells were isolated by magnetic cell sorting and cultured for 3 d in different polarization conditions, as described in 1Materials and Methods, to generate Th-1, Th-2, Th-17, and Treg cells. A, Phenotypical analysis of the obtained T lymphocytes. Cells were stained with the indicated Ab and were analyzed by flow cytometry. B, On day 3, cells were washed and restimulated with anti-CD3/CD28 Ab coated beads for 8 h, and the culture supernatants were collected and analyzed by ELISA. C, Real-time PCR analysis of the cells obtained in B for the expression of the indicated cytokine mRNAs. Significant difference when compared with naive T cell groups; *p < 0.05, **p < 0.0001.

FIGURE 2.

Generation and characterization of Th lymphocyte lineages. Splenic CD4+CD25CD62L+ T cells were isolated by magnetic cell sorting and cultured for 3 d in different polarization conditions, as described in 1Materials and Methods, to generate Th-1, Th-2, Th-17, and Treg cells. A, Phenotypical analysis of the obtained T lymphocytes. Cells were stained with the indicated Ab and were analyzed by flow cytometry. B, On day 3, cells were washed and restimulated with anti-CD3/CD28 Ab coated beads for 8 h, and the culture supernatants were collected and analyzed by ELISA. C, Real-time PCR analysis of the cells obtained in B for the expression of the indicated cytokine mRNAs. Significant difference when compared with naive T cell groups; *p < 0.05, **p < 0.0001.

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Day 6 CD11c+ DCs generated from BALB/c mice were exposed for 48 h to the supernatant of Th-1, Th-2, Th-17, or Treg cultures, and their ability to kill 4T1 tumor cells was evaluated. Our results indicate that only Th-1–derived factor or factors were capable of inducing DC cytotoxic function (Fig. 3A). Similar results were obtained when B16 melanoma cells were used as targets (Fig. 3B), indicating that Th-1–activated KDCs (Th-1 KDCs) are capable of killing both syngeneic and allogeneic tumor cells. Similar results were obtained with DCs and T cells generated from C57BL/6 mice (Supplemental Fig. 2). No new cancer cell colony was observed when detached cells from tumor cell and Th-1 KDC cocultures were incubated for a week in complete medium. Phenotypically, these Th-1 KDCs demonstrated increased expression of MHC II, CD40, and CD86, compared with untreated DCs (Fig. 3C).

FIGURE 3.

Th-1, but not Th-2, Th-17, or Tregs, is capable of inducing DC tumoricidal activity. Day 6 CD11c+ DCs were cultured for 48 h with the supernatants from Th-1, Th-2, Th-17, and Treg cultures. DCs were then washed extensively and incubated for 48 h with 4T1 (A) or B16 (B) tumor cells. KDCs activated with LPS (1 μg/ml) ([DC] LPS) were used as positive controls. Tumor cell survival was then determined. Mean ± SD from triplicate cultures, *p < 0.001. C, Phenotype of DCs obtained after 48 h of culture with the culture supernatant from Th-1, Th-2, Th-17, and Treg cultures. Cells were stained with the indicated Ab and analyzed by flow cytometry.

FIGURE 3.

Th-1, but not Th-2, Th-17, or Tregs, is capable of inducing DC tumoricidal activity. Day 6 CD11c+ DCs were cultured for 48 h with the supernatants from Th-1, Th-2, Th-17, and Treg cultures. DCs were then washed extensively and incubated for 48 h with 4T1 (A) or B16 (B) tumor cells. KDCs activated with LPS (1 μg/ml) ([DC] LPS) were used as positive controls. Tumor cell survival was then determined. Mean ± SD from triplicate cultures, *p < 0.001. C, Phenotype of DCs obtained after 48 h of culture with the culture supernatant from Th-1, Th-2, Th-17, and Treg cultures. Cells were stained with the indicated Ab and analyzed by flow cytometry.

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We next sought to determine the mechanism or mechanisms underlying Th-1–mediated induction of DC killing activity. Because the data depicted in Fig. 1C and 1D indicated that induction of DC killing activity required soluble factors produced by activated CD4+ T lymphocytes, we investigated the role of the prominent Th-1 cytokine, IFN-γ (Ref. 41; Fig. 2B), in the triggering of DC cytotoxic function. CD11c+ DCs were exposed to Th-1 supernatant for 48 h in the presence or absence of anti–IFN-γ blocking Ab. DCs were then washed and cocultured for another 48 h with tumor cells. As illustrated in Fig. 4A and 4B, IFN-γ neutralization prevented Th-1 lymphocyte-mediated induction of DC cytotoxic activity. Consistent with this result, recombinant IFN-γ triggered DC killing activity (Fig. 4A, 4B). We further confirmed the central role of IFN-γ by demonstrating that Th-1 cell supernatant failed to trigger tumor-killing activity of DCs generated from IFN-γ receptor knockout mice (Fig. 4C).

FIGURE 4.

The induction of DC killing activity by Th-1 lymphocytes depends on IFN-γ. Day 6 DCs were treated with Th-1 supernatant ([DC] Th-1) with or without anti–IFN-γ blocking Ab (+ IFN-γ blocking Ab) or isotype control Ab (+ Isotype control) or were treated with recombinant IFN-γ (5 ng/ml) ([DC] IFN-γ) for 48 h. DCs were then washed extensively and plated at a 5:1 DC/tumor cell ratio with 4T1 (A) or B16 (B) tumor cells. C, 4T1 tumor cells were cultured with DCs generated from wild-type (WT) or from IFN-γ receptor−/− mice (IFN-γ R−/−) and treated with Th-1 culture supernatant ([DC] Th-1) or IFN-γ ([DC] IFN-γ). AC, Tumor cell viability was then evaluated. Mean ± SD from triplicate cultures. Untreated DCs (Untreated DC) and DCs treated with LPS (1 μg/ml) ([DC] LPS) were used as negative and positive controls, respectively, *p < 0.001.

FIGURE 4.

The induction of DC killing activity by Th-1 lymphocytes depends on IFN-γ. Day 6 DCs were treated with Th-1 supernatant ([DC] Th-1) with or without anti–IFN-γ blocking Ab (+ IFN-γ blocking Ab) or isotype control Ab (+ Isotype control) or were treated with recombinant IFN-γ (5 ng/ml) ([DC] IFN-γ) for 48 h. DCs were then washed extensively and plated at a 5:1 DC/tumor cell ratio with 4T1 (A) or B16 (B) tumor cells. C, 4T1 tumor cells were cultured with DCs generated from wild-type (WT) or from IFN-γ receptor−/− mice (IFN-γ R−/−) and treated with Th-1 culture supernatant ([DC] Th-1) or IFN-γ ([DC] IFN-γ). AC, Tumor cell viability was then evaluated. Mean ± SD from triplicate cultures. Untreated DCs (Untreated DC) and DCs treated with LPS (1 μg/ml) ([DC] LPS) were used as negative and positive controls, respectively, *p < 0.001.

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We next investigated the mechanisms underlying Th-1 KDC cytotoxic function. Th-1 KDC killing activity was dependent on a direct cell-cell contact, as separation of target and effector cells by a microporous membrane prevented tumor cell death (Fig. 5A). Death receptor ligands (TRAIL or Fas-L) and perforin and granzyme have been described as effector molecules responsible for DC cytotoxic activity (6, 8, 16, 1821, 4245). We therefore evaluated whether Th-1 KDC tumoricidal function was mediated by any of these molecules. TRAIL was not detected in Th-1 KDCs by real-time PCR (Fig. 5B). Fas-L, perforin, and granzyme were also not expressed at significant levels (data not shown). Using knockout mice, we further confirmed that Fas-L and TRAIL did not participate in Th-1 KDC killing activity (Fig. 5C; data not shown). Of note, DCs from Fas-L−/− mice were slightly more effective at killing tumor cells than were DCs from wild-type mice. Similarly, Th-1–activated KDCs from perforin knockout mice were not impaired in their capability to induce tumor cell death (Fig. 5D).

FIGURE 5.

Th-1–induced KDC tumoricidal activity requires a direct cell-cell contact but does not depend on death receptor ligands. A, 4T1 or B16 tumor cells were cultured, separated or not by a Transwell insert (TW), with untreated DCs (Untreated DC), or DCs treated with IFN-γ ([DC] IFN-γ) or Th-1 supernatant–treated DCs ([DC] Th-1). Significant difference compared with the corresponding group without the TW, *p < 0.0001. B, Expression of TRAIL detected by RT-PCR analysis of RNA isolated from day 8 DCs treated for 24 h with LPS (1 μg/ml), IFN-γ (5 ng/ml), or Th-1 supernatant. DCs generated from FasL−/− (C) or perforin−/− (D) mice were treated with IFN-γ or Th-1 supernatant and cultured for 48 h with 4T1 tumor cells. Significant difference compared with wild-type DCs, *p < 0.01. A, C, and D, Tumor cell killing was determined after 48 h. Mean ± SD from triplicate cultures. Untreated DCs (Untreated DC) and DCs treated with LPS (1 μg/ml) ([DC] LPS) were used as negative and positive controls, respectively.

FIGURE 5.

Th-1–induced KDC tumoricidal activity requires a direct cell-cell contact but does not depend on death receptor ligands. A, 4T1 or B16 tumor cells were cultured, separated or not by a Transwell insert (TW), with untreated DCs (Untreated DC), or DCs treated with IFN-γ ([DC] IFN-γ) or Th-1 supernatant–treated DCs ([DC] Th-1). Significant difference compared with the corresponding group without the TW, *p < 0.0001. B, Expression of TRAIL detected by RT-PCR analysis of RNA isolated from day 8 DCs treated for 24 h with LPS (1 μg/ml), IFN-γ (5 ng/ml), or Th-1 supernatant. DCs generated from FasL−/− (C) or perforin−/− (D) mice were treated with IFN-γ or Th-1 supernatant and cultured for 48 h with 4T1 tumor cells. Significant difference compared with wild-type DCs, *p < 0.01. A, C, and D, Tumor cell killing was determined after 48 h. Mean ± SD from triplicate cultures. Untreated DCs (Untreated DC) and DCs treated with LPS (1 μg/ml) ([DC] LPS) were used as negative and positive controls, respectively.

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We and others have previously reported that NO or peroxynitrites are essential short-half-life effector molecules that contribute to the tumoricidal activity of rat, mouse, and human DCs and that require proximity with the target cells (4, 5, 26, 28, 46). We therefore assessed whether these cytotoxic products may also be involved in Th-1 KDC-mediated killing of tumor cells. The concentration of nitrites (the main metabolites of NO) was significantly increased in the cultures of DCs preincubated with Th-1 supernatant (Fig. 6A). In line with this result, iNOS expression was significantly increased in DCs treated with Th-1 supernatant (Fig. 6B). Moreover, NMMA, an inhibitor or iNOS, abrogated Th-1 KDC-mediated tumor killing activity (Fig. 6C). Further confirming these data, the cytotoxic potential of Th-1 supernatant-activated DCs generated from iNOS−/− mice was significantly impaired (Fig. 6D). These findings were reproduced using the B16 model in C57BL/6 mice (data not shown). Taken together, these data demonstrate that NO is essential for Th-1 KDC-mediated tumor cell killing.

FIGURE 6.

Th-1 KDC cytotoxic activity depends on NO. A, Detection of nitrites in the supernatants of DCs treated for 48 h with the supernatants of Th-1, Th-2, Th-17, and Treg cell cultures, or with LPS or IFN-γ. B, iNOS expression was determined by Western blot (left panel) and RT-PCR (right panel) in day 8 DCs treated for 24 h with LPS (1 μg/ml) ([DC] LPS), IFN-γ (5 ng/ml) ([DC] IFN-γ), or Th-1 supernatant ([DC] Th-1). Significant compared with untreated DCs, *p < 0.05. C, DCs activated with LPS, IFN-γ, or Th-1 culture supernatant were incubated for 48 h with 4T1 tumor cells, with or without the iNOS inhibitor NMMA (1 mM). D, DCs generated from iNOS−/− or wild-type mice were treated with IFN-γ or Th-1 supernatant and cultured for 48 h with 4T1 tumor cells. C and D, Tumor cell killing was determined after 48 h. Mean ± SD from triplicate cultures. A, C, D, *p < 0.001. Untreated DCs and DCs treated with LPS were used as negative and positive controls, respectively.

FIGURE 6.

Th-1 KDC cytotoxic activity depends on NO. A, Detection of nitrites in the supernatants of DCs treated for 48 h with the supernatants of Th-1, Th-2, Th-17, and Treg cell cultures, or with LPS or IFN-γ. B, iNOS expression was determined by Western blot (left panel) and RT-PCR (right panel) in day 8 DCs treated for 24 h with LPS (1 μg/ml) ([DC] LPS), IFN-γ (5 ng/ml) ([DC] IFN-γ), or Th-1 supernatant ([DC] Th-1). Significant compared with untreated DCs, *p < 0.05. C, DCs activated with LPS, IFN-γ, or Th-1 culture supernatant were incubated for 48 h with 4T1 tumor cells, with or without the iNOS inhibitor NMMA (1 mM). D, DCs generated from iNOS−/− or wild-type mice were treated with IFN-γ or Th-1 supernatant and cultured for 48 h with 4T1 tumor cells. C and D, Tumor cell killing was determined after 48 h. Mean ± SD from triplicate cultures. A, C, D, *p < 0.001. Untreated DCs and DCs treated with LPS were used as negative and positive controls, respectively.

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To investigate the ability of Th-1 KDCs to process and present Ags from the tumor cells they kill, Th-1–activated DCs were cocultured for 48 h with B16 melanoma cells expressing the model Ag OVA (B16-OVA). DCs were then reisolated using CD11c microbeads and either stained with anti-CD11c and an Ab recognizing MHC class I–SIINFEKL complexes (Kb/SIINFEKL) or cultured for 24 h with B3Z (a CD8+ T cell line expressing a TCR that specifically recognizes the SIINFEKL peptide of OVA in the context of MHC class I) (5). We did not detect CD11c H-2Kb/SIINFEKL+ cells (contaminating tumor cells) following isolation, but MHC class I–SIINFEKL complexes were detected at the surface of isolated CD11c+ DCs (Supplemental Fig. 3). In addition, purified Th-1 DCs were capable of activating B3Z cells (Fig. 7A). These results therefore indicate that Th-1 KDCs are able to acquire, process, and present Ags from the OVA-expressing B16 cells they kill.

FIGURE 7.

Th-1 KDCs are capable of presenting tumor Ags from the tumor cells they have killed to tumor-specific T cells. CD11c+ DCs were treated with LPS (1 μg/ml) ([DC] LPS), IFN-γ (5 ng/ml) ([DC] IFN-γ), or Th-1 supernatant ([DC] Th-1) for 48 h. A, DCs were then washed and cultured for 24 h with B16 or B16-OVA melanoma cells. DCs were selected from the culture using CD11c microbeads and incubated for an additional 24 h with B3Z cells (DC/B3Z ratio = 1:10), as outlined in 1Materials and Methods. The activity of β-galactosidase was measured by evaluating the conversion of its substrate into a chemoluminescent product (RLU, relative luminescence unit). B, CD11c-GFP-DTR mice were injected with B16-OVA melanoma cells. When tumors become palpable, DT was administered to deplete host CD11c+ cells. CD11c+ DCs generated in vitro and treated with LPS or IFN-γ or Th-1 supernatant were then injected intratumorally (20 × 106 cells/tumor). After 36 h, draining lymph nodes were harvested, and CD11c+ cells were isolated using CD11c microbeads. The obtained DCs were then cultured with the OVA-specific T cell line B3Z. Specific recognition of tumor-derived OVA peptide was measured as outlined above. Significant difference when compared with untreated DC group, *p < 0.01.

FIGURE 7.

Th-1 KDCs are capable of presenting tumor Ags from the tumor cells they have killed to tumor-specific T cells. CD11c+ DCs were treated with LPS (1 μg/ml) ([DC] LPS), IFN-γ (5 ng/ml) ([DC] IFN-γ), or Th-1 supernatant ([DC] Th-1) for 48 h. A, DCs were then washed and cultured for 24 h with B16 or B16-OVA melanoma cells. DCs were selected from the culture using CD11c microbeads and incubated for an additional 24 h with B3Z cells (DC/B3Z ratio = 1:10), as outlined in 1Materials and Methods. The activity of β-galactosidase was measured by evaluating the conversion of its substrate into a chemoluminescent product (RLU, relative luminescence unit). B, CD11c-GFP-DTR mice were injected with B16-OVA melanoma cells. When tumors become palpable, DT was administered to deplete host CD11c+ cells. CD11c+ DCs generated in vitro and treated with LPS or IFN-γ or Th-1 supernatant were then injected intratumorally (20 × 106 cells/tumor). After 36 h, draining lymph nodes were harvested, and CD11c+ cells were isolated using CD11c microbeads. The obtained DCs were then cultured with the OVA-specific T cell line B3Z. Specific recognition of tumor-derived OVA peptide was measured as outlined above. Significant difference when compared with untreated DC group, *p < 0.01.

Close modal

We next assessed the antitumoral potential of Th-1 KDCs in vivo. B16-OVA tumors were established in CD11c-GFP-DTR mice. Animals were then treated with DT to eliminate host CD11c+ DCs when tumors became palpable. This approach prevented possible interference by endogenous DCs. Th-1–activated killer DCs were then injected into the tumor beds, and CD11c+ cells were isolated from the tumor draining lymph nodes after 36 h. These purified CD11c+ DCs were GFP negative and therefore entirely of donor origin (Supplemental Fig. 4A). Their ability to activate the OVA-specific B3Z hybridoma was then evaluated. Our results indicate that only DCs activated with LPS or IFN-γ, and Th-1 KDCs (e.g., killer DCs), but not untreated nonkiller DCs, were capable of inducing B3Z activation (Fig. 7B). Th-1 KDCs were also capable of inducing the proliferation of lymphocytes isolated from OT-II or OT-I mice (Supplemental Fig. 4B; data not shown). These results therefore demonstrate that Th-1–activated killer DCs are capable of migrating from the tumor site to the lymph nodes, and that functionally these killer DCs retained their ability to process and present in a MHC class I- and class II-restricted manner the in vivo acquired Ags, as demonstrated by ex vivo assays.

DCs play an essential role in the initiation and regulation of tumor-specific immune responses, as they are endowed with the unique potential to efficiently activate anticancer effector cells such as Th and cytotoxic T cells (1). This capacity has been extensively exploited, leading to the development of DC-based cancer immunotherapies. However, the initial evidence that protective antitumor immunity can be successfully generated by vaccination with tumor Ag-loaded DCs has been undermined by the limited clinical responses observed in cancer patients. Therefore, the prospect of exploiting the nonconventional direct tumor-killing function of DCs represents an important step forward for the advancement of these cells in cancer therapies. Not only are KDCs indeed endowed with the capacity of directly killing tumor cells (and thereby participating in the effector mechanisms of immune responses), but they can also generate their own source of released tumor Ags immediately available for uptake, processing, and presentation to specific T lymphocytes (5, 28).

Several agents have been reported to trigger DC killing potential, including cytokines, such as IFNs, TNF-α, IL-12, IL-18, IL-2, and IL-15 (14, 17, 19, 24, 30, 47), or TLR ligands (8, 18, 31), such as the TLR4 ligand LPS, as we and others have previously reported (4, 5). However, the optimal mode of activation of DC cytotoxic activity remains to be determined. DC-Th lymphocyte cross-talk critically contributes to the regulation of conventional DC function (48), but the putative effects exerted by T cells on the cytotoxic activity of DCs had not, to our knowledge, been previously delineated. In the current report, we provide evidence that only activated proinflammatory Th-1 lymphocytes can drive the activation of bone marrow-derived DCs into potent tumor killers.

The mechanism of induction of DC killing activity by these Tbet+ Th-1 cells does not depend on direct cell contact but requires the Th-1–related cytokine IFN-γ. The possibility that IFN-γ may induce DC cytotoxic activity has been controversial. This observation has been documented primarily in the case of human DCs and only when high and less physiologically achievable concentrations of this cytokine were used (4, 6, 14, 17, 29, 30). In addition, only recombinant IFN-γ was tested. We provide novel evidence that IFN-γ produced by Th-1 at physiological levels can activate mouse bone marrow-derived DC tumoricidal potential. In our study, the concentration of IFN-γ detected in Th-1 supernatants capable of inducing DC killing activity was significantly lower than that tested in previous reports (4, 6, 14, 17, 29, 30). In addition, in most of these studies cytotoxic effects were detected only when significantly higher effector KDC/target tumor cell ratios were used (4, 6, 14, 17, 29, 30).

The mode of tumor cell killing by DCs may involve the death receptor family and their ligands (6, 8, 1013, 15, 1720, 25, 31), perforin and granzyme (8), or, as we previously reported, NO or peroxynitrites (4, 11, 23, 26, 27). NO has been shown to sensitize tumor cells to Fas ligand-mediated killing by immature or spontaneously matured DCs (9). A NO donor was used at low (and noncytotoxic concentrations) to modulate antiapoptotic pathways (9). In the same study, the NO donor alone was, however, capable of inducing tumor cell death when used at higher concentrations (9). We demonstrated that iNOS expression is significantly increased in Th-1 KDCs and identified NO as the primary cytotoxic effector molecule implicated in Th-1 KDC-mediated tumor cell elimination, as DCs from iNOS−/− mice were not capable of killing cancer cells. The levels of NO produced by activated DCs were sufficient to trigger the death of tumor cells, and ligands of the TNF superfamily were not involved in this process. The short half-life of these cytotoxic molecules is also consistent with a cell to cell contact-dependent mechanism of tumor cell killing.

Typically, only immature DCs are efficient at taking up Ags (49). The observation that activated mature Th-1 KDCs, following killing of cancer cells, are still capable of capturing tumor Ags is therefore of importance. This property allows them to subsequently process and present the derived antigenic peptides to T cells and therefore to initiate adaptive immune responses. Of therapeutic relevance, Th-1 KDCs injected into the tumor beds that migrated to the draining lymph nodes were capable of activating T lymphocytes, indicating that they acquired Ags and have the ability of presenting them in vivo. This capacity of Th-1 KDCs to efficiently present Ags is contingent upon induction of their killing function, as nonkiller DCs were significantly less potent APCs. Further supporting this observation, DCs from iNOS−/− mice were less potent at activating specific T cells following culture with tumor cells, compared with their wild-type counterparts (data not shown).

The potential for DCs to act not only as APCs but also as tumor cell killers has revitalized their attractiveness as immunotherapeutic agents against cancer. Our results further advocate for the implementation of KDCs in immunotherapy strategies and highlight new possibilities related to the mode of induction of their killing and Ag presenting function, and underline the need for rethinking DC-based vaccine approaches.

We thank Anthony Pilutti for assistance.

This work was supported by National Institutes of Health Grant R01 CA104926 (to E.K. and N.L.), Cancer Biology Training Grant T32CA009213 (to S.C.), Arizona Cancer Center Support Grant CA023074 (to E.K. and N.L.), Tee Up for Tots, and PANDA Funds (to E.K. and N.L.).

The online version of this article contains supplemental material.

Abbreviations used in this article:

DC

dendritic cell

DT

diphtheria toxin

iNOS

inducible NO synthase

KDC

killer DC

NMMA

NG-methyl-L-arginine

Th-1 KDC

Th-1–activated KDC

Treg

regulatory T cell.

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The authors have no financial conflicts of interest.