NK Cells Are Required for Dendritic Cell–Based Immunotherapy at the Time of Tumor Challenge

Anthea L. Bouwer, Sarah C. Saunderson, Felicity J. Caldwell, Tanvi T. Damani, Simon J. Pelham, Amy C. Dunn, Ralph W. Jack, Patrizia Stoitzner and Alexander D. McLellan
J Immunol March 1, 2014, 192 (5) 2514-2521; DOI: https://doi.org/10.4049/jimmunol.1202797

Abstract

Increasing evidence suggests that NK cells act to promote effective T cell–based antitumor responses. Using the B16-OVA melanoma model and an optimized Gram-positive bacteria–dendritic cell (DC) vaccination strategy, we determined that in vivo depletion of NK cells at time of tumor challenge abolished the benefit of DC immunotherapy. The contribution of NK cells to DC immunotherapy was dependent on tumor Ag presentation by DC, suggesting that NK cells act as helper cells to prime or reactivate tumor-specific T cells. The absence of NK cells at tumor challenge resulted in greater attenuation of tumor immunity than observed with selective depletion of either CD4 or CD8 T cell subsets. Although successful DC immunotherapy required IFN-γ, perforin expression was dispensable. Closer examination of the role of NK cells as helper cells in enhancing antitumor responses will reveal new strategies for clinical interventions using DC-based immunotherapy.

Introduction

Survival outcomes for patients with advanced metastatic melanoma are poor despite advances in immunotherapy and targeted therapy, with the median survival ≤1 y (1, 2). Current treatment options include surgery in combination with chemotherapy, postoperative radiation therapy, adjuvant therapy with IFN-α-2b, or targeting intracellular pathways such as BRAF (2–4). As melanoma is an immunogenic tumor, vaccines have been investigated in the hope of enhancing antitumor responses through the improvement of Ag presentation to naive and memory T cells (4) or as restimulation protocols following infusion of ex vivo–expanded, tumor Ag–specific T cells (5). The range of vaccines developed have included cell lysates, peptide/protein-based vaccines, cytokine-based vaccines, and whole-tumor cell vaccines with a wide variety of delivery systems, adjuvants, injection frequencies, and routes of administration being investigated (6). To date, vaccines that have been trialed have been derived from melanoma-associated tumor Ags including gp-100, MART-1, TRP-2, and GD3-ganglioside. With the possible exception of available adoptive T cell therapy, vaccines have not yielded a significant improvement in overall survival in patients with resected melanoma above that of standard chemotherapy (2–7).

Dendritic cells (DC) are potent APCs able to activate naive and memory CD4 and CD8 T cells (8). DC-based immunotherapies take advantage of the unique Ag-presenting capabilities of these cells for the treatment of malignancies. The efficiency and quality of the response generated is dependent on the maturation state of the DC, which affects DC migration to the regional lymph node and T cell priming (9).

The lack of clinical effectiveness of current cancer vaccines has led to increased efforts to understand what is required to elicit protective CD4 and CD8 T cell responses, as well as T cell memory cell formation. Most DC immunotherapy strategies aim to activate T cells to directly eliminate cells expressing tumor Ags or via cytokine effects on the tumor or the supporting vasculature (10–14). A further complementary approach is to develop adjuvants that activate both T cells and NK cells. In the case of melanoma, although T cell responses may be useful, the downregulation of MHC class I (MHC-I) on melanoma cells may play a significant role in the pathogenesis and progression of the disease (1).

NK cells detect and destroy malignant and virally infected cells (15–24). Aside from their cytotoxic abilities against low MHC-I tumor cells, interactions between NK cells and DC have been shown to result in the activation of a helper NK cell phenotype, characterized by the production of IFN-γ and associated with enhanced CTL and Th1 responses (15, 16, 25–29). NK cells also possess adaptive features (30), and the NK cell subset responsible for memory-like responses was recently shown to be a liver-derived CD49blo/NK1.1hi subset (31).

IFN-γ has been shown to be a key effector cytokine in antitumor responses, directly affecting tumor cells by decreasing proliferation and metabolic activity, enhancing costimulator molecule and MHC expression, as well as inhibiting angiogenesis by tumor cells through induction of CXC chemokines (13, 14, 16, 32). Adjusting DC-based therapies to augment NK cell function and IFN-γ secretion could therefore complement current CD8 T cell strategies in controlling tumor growth (17–29).

We have investigated the potential of TLR-agonist adjuvants, including intact bacteria, to enhance DC-based immunotherapy through NK cell activation. Streptococcus salivarius was selected for this study as it has previously been shown to induce the highest amount of IL-12 in human PBMC from a panel of 37 taxonomically diverse bacteria tested. S. salivarius induces high-level IFN-γ expression in both human PBMCs (33) and murine NK cells (34). Using a therapeutic tumor model, we found that in vivo depletion of NK cells completely abolished the benefit of the bacteria-stimulated DC immunotherapy. Furthermore, whereas NK cells were critical for the antitumor response, they did not exert an effector function in the absence of tumor Ag vaccination. NK cells isolated from lymph nodes of immunized mice showed enhanced IFN-γ secretion, but displayed similar levels of granzyme B and CD107a (Lamp-1) levels to sham-immunized mice. Moreover, IFN-γ, but not perforin, was shown to be essential for the success of DC immunotherapy, suggesting that soluble factor release, rather than direct cell-mediated killing, was the major mechanism of tumor inhibition by NK cells. Together, these data demonstrate that NK cells act in concert with Ag-driven T cell responses to play an essential role in the antitumor response in DC immunotherapy.

Materials and Methods

Mice

Six- to 8-wk-old BALB/c, C57BL/6 (The Jackson Laboratory, Bar Harbor, ME), perforin-deficient (pfp−/−), and IFN-γ–deficient (ifnγ−/−) mice (obtained from Malaghan Institute, University of Wellington, New Zealand) were bred and housed at the Hercus-Taieri Research Unit under specific pathogen-free conditions at the University of Otago Hercus-Taieri Research Unit as described (35). All experiments were approved by the regional animal ethics committee.

Preparation and stimulation of DC

Bone marrow–derived DC were generated as previously described (36) in GM-CSF–supplemented complete medium (R10) composed of RPMI 1640 (Invitrogen, Auckland, New Zealand) with 10% FBS (PAA Laboratories), 100 U/ml penicillin, 100 μg/ml streptomycin (Invitrogen), and 55 μM 2-ME (Invitrogen). On day 7, DC were pulsed for 24 h with 200 μg/ml OVA protein (OVA; Sigma-Aldrich, Auckland, New Zealand) and overnight with heat-killed (56°C, 1 h) Streptococcus salivarius K12 at a 1:1 ratio or Salmonella typhimurium LPS (1 μg/ml), Staphylococcus aureus lipoteichoic acid (LTA; 1 μg/ml; all from Sigma-Aldrich), Pam3Cys, or CpG-1826 (both 10 μg/ml; Invivogen, San Diego, CA). Cells were then washed three times with PBS and injected s.c. into the regio interscapular, forelimb, or flank, as indicated. S. salivarius strain K12 was maintained on blood agar containing 0.1% CaCO3 (Fort Richard Laboratories, Auckland, New Zealand). Single colonies were grown at 37°C + 5% CO2 in Todd-Hewitt broth overnight, after which the bacterial count was calculated using OD (OD600) with routine CFU calibration. Heat-killed bacteria were washed in 0.1% BSA/PBS/2 mM EDTA (3220 × g, 5 min) and resuspended in R10.

Lymph node analysis, flow cytometry, in vitro cytokine production, and cytotoxicity assays

Forty-eight hours after s.c. immunization, draining lymph nodes (axillary, brachial, and cervical) were harvested. Cells were stained with DX5-PE and TCRβ-FITC (clone H57-597) and analyzed by flow cytometric analysis using an FACScalibur (Figs. 1, 5) or an LSR Fortessa (Figs. 2, 8) (both from BD Biosciences, Auckland, New Zealand) and FlowJo (Tree Star, San Diego, CA) analysis software. Abs to CXCR3 (G025H7-BV421; BioLegend), granzyme B (FGB12-allophycocyanin; Invitrogen), IFN-γ (XMG1.2-allophycocyanin), CD107a (Lamp-1; 1D4B-FITC) CCR7 (4B12-allophycocyanin), H-2K/D (M1/42-PE), or I-A/E (M5/114-PE; all from BD Biosciences) were used to phenotype NK cells or B16-OVA. In some experiments as indicated, lymph node cells at 105/ml in R10 were restimulated at a 1:1 ratio with a YAC-1 or B16-OVA cell line for 4 h prior to flow cytometric analysis. For IFN-γ analysis, monensin (Sigma-Aldrich) was included in coculture experiments at 10 μg/ml. CD107a-FITC was added to the culture medium at 5 μg/ml for the 4-h incubation coculture with YAC-1 or B16-OVA. For flow cytometric analysis, gates were set to include <1% of events in DX-5/TCRβ/isotype-control–labeled cell suspensions. For the in vitro cytokine release assays, splenocytes were cultured at 2 × 106/ml in the presence or absence of S. salivarius K12 in 96-well round-bottom plates (Nunc, Auckland, New Zealand) for 18 h. Soluble IFN-γ was measured by ELISA (BD Biosciences). In vitro cytotoxicity of lymph node or splenic NK cells against the B16-OVA melanoma line was tested by plating 250 target B16-OVA cells in 10 μl R10 in triplicate in Terasaki wells (37). Effector NK cells, isolated from lymph node or spleen using the untouched NK cell Isolation Kit II (Miltenyi Biotec, Pharmaco, Auckland, New Zealand), were overlaid in 5 μl R10 at a 100:1 E:T ratio. After 24 h, NK cells were washed away with PBS and adherent B16-OVA cells enumerated after detection with May-Grünwald Giemsa.

In vivo depletions of lymphocyte subsets and IFN-γ

NK cell depletion was performed using low-dose (20 μl) rabbit anti–asialo GM1 (ASGM) antiserum (Wako) (19) by i.v. injection in 100 μl PBS at day −1 and day 2 relative to tumor inoculation (therapeutic model) or DC immunization (prophylactic model). For CD4 and CD8 T cell depletions YTS191.1 (CD4) and 53-5.8 (CD8β), mAbs were administered i.v. 100 μg/mouse on days −3, −1, and 2 relative to tumor challenge. The efficiency of lymphocyte depletions was determined in pilot experiments in both spleen and lymph nodes by flow cytometry. Mice were depleted of IFN-γ by i.v. administration of 100 μg XMG.D6 (kind gift of Franca Ronchese, Malaghan Institute, Wellington, New Zealand) at days −1, 2, and 4 (relative to tumor inoculation).

Tumor inoculation and monitoring and cytotoxicity assay

B16-OVA, a C57BL/6 mouse-derived melanoma B16-F0 cell line transfected with the full-length OVA gene (38), was cultured in R10 with 50 μg/ml geneticin (Life Technologies, Auckland, New Zealand). For prophylactic tumor experiments, B16-OVA was cultured in serum-free medium to avoid anti-FCS immune responses primed by DC exposed to FCS (39). B16-OVA cells were harvested from logarithmically growing cultures using a cell scraper, filtered through a 70-μm cell strainer, and resuspended in HBSS. In the therapeutic setting, mice were immunized s.c. with 1 × 105 B16-OVA cells in the flank. Three days later, DC (2 × 105) were administered s.c. into the opposite flank. Prophylactic experiments involved mice immunized with OVA/S. salivarius–stimulated DC 30 or 60 d prior to tumor challenge. Tumor growth was determined by measuring the length and width using calipers every 1–2 d. Results are expressed as the mean product of the tumor diameters. Mice were removed from the study when tumors reached 150 mm2. Data are represented as tumor growth curves or as Kaplan-Meier survival plots using Graphpad Prism 5 (GraphPad, San Diego, CA) and analyzed using the Mantel-Cox test. Tumor growth curves represent the mean values of five to eight mice and terminate upon the loss of the first mouse.

Results

Optimized DC immunotherapy

To compare the ability of whole bacteria to standard TLR-agonist adjuvants in tumor immunotherapy, we used a therapeutic setting in which C57BL/6 mice were administered B16-OVA s.c. in the left flank and 3 d later immunized in the opposite flank with OVA-pulsed DC (DC-OVA) stimulated with S. salivarius or other adjuvants. As shown in Fig. 1A, s.c. immunization of mice with S. salivarius or LTA-stimulated DC-OVA caused a significant delay in tumor growth, greater than that observed when unstimulated or LPS-stimulated DC-OVA were administered. The addition of a second DC immunization at day 7 did not markedly further delay tumor growth (Fig. 1B), but confirmed the superiority of S. salivarius in inhibiting tumor growth. In addition, DC matured with S. salivarius were significantly more active than other selected adjuvants at inducing NK cell influx into skin-draining lymph nodes (Fig. 1C). NK cells isolated from the draining lymph nodes or spleen of S. salivarius–DC-immunized mice exhibited no detectable in vitro cytotoxicity against either B16-OVA (Fig. 1D) or B16 parental strain (data not shown). Although CD49b+/TCRβ− lymph node NK cells responding to S. salivarius–DC vaccination displayed similar levels of the CXCR3 or CCR7 chemokine receptors compared with PBS-treated mice (Fig. 2A), NK cells from S. salivarius–DC-immunized mice demonstrated increased potential to secrete IFN-γ in response to YAC-1 recognition (Fig 2B), but not B16-OVA restimulation (data not shown). No increase in granzyme B or CD107a (Lamp-1) expression was noted during coculture with either YAC-1 (Fig. 2B) or B16-OVA (data not shown).

FIGURE 1.
FIGURE 1.

Assessment of the adjuvant properties of TLR ligands or whole bacteria in therapeutic DC-based immunotherapy and lymph node NK cell recruitment. C57BL/6 mice (six per group) were inoculated s.c. with B16-OVA at day 0. (A) Three days later, mice were primed with a single injection of OVA/adjuvant-treated DC in the opposite flank. Survival of the mice in the LTA-DC/OVA and S. salivarius (Ssa)–stimulated DC/OVA groups was significant when compared with the control (p < 0.01). Representative of two experiments performed. (B) Double immunization strategy: mice (eight per group) inoculated s.c. with B16-OVA at day 0. Three and 10 d later, mice were primed with OVA/adjuvant-treated DC. Survival was significantly better in the Ssa-DC/OVA group compared with the tumor-only control, for which p < 0.0001. (C) Whole bacteria or TLR ligand–matured DC (2 × 106) were injected s.c. into the regio interscapular. Two days later, the percentages of CD49b+/TCRαβ− NK cells were measured in the control and DC-draining cervical, brachial, and axilliary lymph nodes (LN). The data are presented as mean ± SEM of three separate experiments in which two mice were analyzed in duplicate. Data were analyzed by one-way ANOVA, applying a Tukey-Kramer posttest to compare all groups in which Ssa-DC were better at recruiting NK cells to LN than both control and CpG-DC (p < 0.05). (D) The cytotoxicity of axilliary and brachial LN or splenic (Spl) NK cells isolated from PBS- or DC-immunized (s.c., forelimb) C57BL/6 mice against B16-OVA. Graph shows percent viable B16-OVA cells remaining after 24 h in NK cell/B16-OVA coculture at 100:1 ratio, in which 100% viability is the mean of triplicate B16-OVA counts in the absence of NK cells. One of two experiments performed.

FIGURE 2.
FIGURE 2.

Phenotype of influxing lymph node NK cells. C57BL/6 mice were immunized s.c. in the forelimb with PBS or bacteria-pulsed DC. Two days later, draining axilliary and brachial lymph nodes were removed and NK cells within the lymph node suspensions either labeled immediately with Abs to CD49b and TCRβ and CXCR3 and CCR7 (A) or granzyme B, CD107a, or IFN-γ (B) following a restimulation with YAC-1 cells (1:1 ratio) for 4 h. Plots show CD49b+/TCRβ− events and gates were set to include <1% of events in DX-5/TCRβ/isotype control–labeled cell suspensions. The percent positive events are represented within each gate of the contour plots. Representative of three mice per group in two independent experiments.

NK cells are required for optimized DC immunotherapy against melanoma

To study the role of NK cells in the observed delay of tumor growth after S. salivarius K12-stimulated DC-OVA immunization, we performed therapeutic tumor experiments in which NK cells were depleted using rabbit ASGM (Fig. 3A) prior to administration of DC therapy. Protection that was previously observed by the administration of DC therapy was completely abrogated by in vivo depletion of NK cells with ASGM (Fig. 3A) or PK136 (data not shown), with tumor outgrowth in NK cell–depleted mice being comparable to that of the control animals. We next determined whether activation of NK cells by mature DC was sufficient for the antitumor response or if an Ag-specific T cell response was also required. The data (Fig. 3B) showed that in vivo stimulation of NK cells with bacteria-matured DC, in the absence of tumor Ag, had little or no effect on the progression of tumor growth, showing that the inhibition of tumor growth was Ag dependent.

FIGURE 3.
FIGURE 3.

NK cell depletion reduces the protection afforded by DC vaccination. (A) C57BL/6 mice were inoculated s.c. with B16-OVA at day 0. Mice were injected i.v. with rabbit ASGM (or control rabbit Ig) to deplete NK cells. On day 3, mice were primed s.c. in the opposite flank with OVA/S. salivarius–stimulated DC. Tumor growth inhibition as well as survival was significantly different in the DC/OVA group with NK cells, compared with untreated or DC/OVA plus NK cell depletion (p < 0.0001). Data are representative of two experiments performed using six mice per group. (B) Mice (five per group) were inoculated s.c. with B16-OVA at day 0. Three days later, mice were primed with (or without) OVA/S. salivarius–stimulated DC. Tumors were measured from day 7 and every second day thereafter. Survival differences between the S. salivarius-DC/OVA group and both the untreated and no OVA groups showed significance (p < 0.05), respectively. Representative of two experiments performed.

Dissection of role of NK cells at the priming and challenge phase of DC immunotherapy

To determine if NK cells were acting at the DC-priming or tumor-challenge phase, we used a prophylactic setting in which mice were depleted of NK cells prior to the administration of DC therapy (Fig. 4A). NK cells were then allowed to regenerate for 30 d before mice were challenged with B16-OVA. A second group of mice were NK cell–depleted immediately prior to tumor challenge. The growth curves (Fig. 4A) suggested that NK cells were involved in both the priming and challenge phases of the immune response. However, while this experiment was being conducted, additional mice were monitored to assess the repopulation of NK cells in ASGM-depleted mice. Surprisingly, it was found that by day 30, only ∼50% of NK cells had returned (Fig. 5A). In addition, NK cell–depleted splenocytes at day 30 post-ASGM produced lower levels of IFN-γ in response to whole bacteria compared with the control mice (Fig. 5A). Because this rapid IFN-γ response to S. salivarius was solely due to NK cell activity and was independent of T cell or NK–T cell activation (34), the observed deficit in NK cell numbers was also manifested in a functional defect in NK cell activity. The slow recovery of NK cell numbers from NK cell–depleted mice at priming could result in impaired NK cell function at the time of challenge. The prophylactic experiment was therefore repeated in which mice were NK cell depleted at either the time of DC immunization or 60 d later at the time of challenge (Fig. 4B) to allow a fuller recovery of NK cell numbers and the IFN-γ response to whole bacteria (Fig. 5B). In the 60-d experiment, the depletion of NK cells around the time of challenge showed a significant loss in efficacy of DC vaccination (Fig. 4B). Although NK depletion at the time of priming inhibited the antitumor effect, this effect was not as profound as the absence of NK cells during the challenge phase (Fig. 4B). The selective depletion of T cell subsets at the time of challenge revealed a potential role for CD4 T cells in the observed protection (Fig. 6A). However, differences in tumor growth rates between control and CD4 T cell subset-depleted groups failed to reach statistical significance, except in the group that was codepleted of both CD4 T cells and NK cells. In contrast, depletion of CD8 T cells had little effect on tumor growth inhibition (Fig. 6B). To determine if NK cell depletion could alter T cell activation in vivo, mice were adoptively transferred with OVA-specific OT-I or OT-II T cells as previously described (36) and then depleted of NK cells using ASGM. Mice were immunized s.c. with S. salivarius–DC 2 d later. However, no defect in lymph node T cell activation was noted, as shown by the similar CFSE dilution profiles between DC immunized control and ASGM mice (Supplemental Fig. 1).

FIGURE 4.
FIGURE 4.

Requirement for NK cells at priming and challenge. Mice were primed s.c. in the flank with OVA/S. salivarius–stimulated DC at day 0. Thirty (A) or 60 d (B) later, mice were inoculated s.c. in the opposite flank with B16-OVA. In one group, mice were depleted of NK cells the day prior to priming and 3 d later. In another group, mice were depleted of NK cells the day prior to tumor challenge and again 3 d later. (A) There was a significant difference in survival between NK cell–replete mice and mice that were depleted at time of priming and challenge as well as between depletion at challenge (p < 0.0005). (B) There were significant survival differences (p < 0.001) between NK cell–replete mice and mice that were depleted of NK cells at the time of challenge only. Data from one experiment performed at each time point using six mice per group.

FIGURE 5.
FIGURE 5.

NK cell regeneration and function following ASGM depletion. Mice were depleted of NK cells with ASGM. Control mice were injected i.v. with the equivalent amount of rabbit Igs. (A) Mice (four per group) were sacrificed, and splenocytes were isolated, labeled with DX5-PE and TCRβ-FITC, and analyzed by flow cytometry. Graph shows the regeneration (means of CD49b+/TCRβ− events of 4 mice ± SD) of NK cells following ASGM administration with a representative flow cytometric plot showing NK cell depletion at day 2 in control (rabbit Ig) or ASGM. Doublets, dead cells, and debris were excluded by forward light scatter/side scatter gating strategies. Representative of two experiments using three to four mice per group. (B) Samples of splenocytes from day 30 or 60 postdepletion groups were also stimulated with S. salivarius (Ssa; multiplicities of infection of 10, 1, and 0.1) overnight and released IFN-γ measured by ELISA. Data represented as mean ± SD from two mice in duplicate. One of two representative experiments.

FIGURE 6.
FIGURE 6.

Investigating the roles of NK and T cells at time of tumor challenge. Mice were administered S. salivarius–stimulated DC/OVA. Sixty days later, mice were depleted of CD4 (A) or CD8 T cells (B) and/or NK cells (A, B) and challenged with B16-OVA. (A) Differences between the +NK+CD4 and −NK−CD4 groups were significant in survival (p < 0.05). (B) Differences between the +NK+CD8 and all groups, except the +NK−CD8 group was significant in survival (p < 0.01). Data representative of two similar experiments using six to seven mice per group.

Role of IFN-γ and perforin in DC immunotherapy

The mechanism of DC immunotherapy was next investigated using neutralizing Ab against IFN-γ as well as ifnγ−/− and pfp−/− mice. The results showed a strict dependence of DC immunotherapy on IFN-γ expression. In contrast, perforin expression was dispensable, and significant antitumor responses were still noted in DC-immunized versus control pfp−/− mice (Fig. 7). In vitro assays showed that direct treatment of B16-OVA with IFN-γ did not retard growth (Fig. 8). However, IFN-γ treatment of our B16-OVA subline stimulated a marked upregulation of MHC-I and MHC class II (Fig. 8), suggesting a role of IFN-γ in enhancing presentation of tumor Ags.

FIGURE 7.
FIGURE 7.

Roles of IFN-γ and perforin in DC immunotherapy. C57BL/6 mice (A) or pfp−/− or ifnγ−/− mice (B) were inoculated s.c. with B16-OVA at day 0 and administered S. salivarius–stimulated DC/OVA at day 3. For (A), anti–IFN-γ Ab was administered at days −1, 2, and 4. DC immunotherapy induced a significant level of protection in control DC groups (p < 0.01), but DC immunotherapy failed to induce a significant survival benefit in anti–IFN-γ–depleted mice. A statistically significant difference in survival was noted between untreated pfp−/− and DC-immunized pfp−/− mice (p < 0.001), but not between untreated and DC-immunized ifnγ−/− mice (B). No significant difference in survival was noted between DC-treated C57BL/6 (A) and DC-treated pfp−/− mice (B), nor between untreated C57BL/6 (A) and untreated or DC-treated ifnγ−/− mice (B). Data from one experiment each using six mice per group.

FIGURE 8.
FIGURE 8.

Effect of IFN-γ on B16-OVA viability and MHC expression. (A) B16-OVA was cultured at 5 × 104 cells/well in a 96-well flat-bottom plate in triplicate in the presence of the indicated concentrations of rIFN-γ (R&D Systems, Auckland, New Zealand) or 250 nM staurosporine (Staur; Sigma-Aldrich) as a positive control for cell death induction in a total of 200 μl of R10 for 24 h. Alamar Blue (Invitrogen; 20 μl/well) was then added for 4 h and dye reduction determined by fluorescence using a Thermo Fisher Varioskan plate reader (Thermo Fisher Scientific, Auckland, New Zealand). Results are plotted as arbitrary units measured at an excitation wavelength (ex) of 540 nm and an emission wavelength (em) of 585 nm. Death induced by staurosporine was significantly different from control cultured B16-OVA (****p < 0.0001) by one-way ANOVA. (B) B16-OVA was cultured for 24 h in six-well plates at 106 cells/well in the presence or absence of 20 ng/ml rIFN-γ. B16-OVA cells were then labeled with Abs to H-2K/D (M1/42-PE) or I-A/E (M5/114-PE) and analyzed by flow cytometry. Shaded area, isotype control; dotted line, media only; solid line, 20 ng/ml IFN-γ. One of three experiments.

Discussion

In this study, we have optimized a DC-based adjuvant strategy for melanoma and have dissected the role of NK cells and IFN-γ in the antitumor response. The DC-based immunotherapy was optimized by providing a potent microbial stimulus of whole bacteria and our results confirm those of others showing that NK cells and IFN-γ are required for optimal DC based therapy (17, 18, 23). Furthermore, we could demonstrate that the major contribution of NK cells to DC-based immunotherapy was at the time of tumor challenge.

NK cell–mediated destruction of tumor cells may lead to the release of tumor Ag into the lymphatic system. Indeed, cell-associated OVA has been shown to be cross-presented much more efficiently than soluble OVA (40), and NK cell destruction of tumor cells could enhance uptake and presentation of dead cell–associated tumor Ag for the induction of T cell responses, similar to localized radiation effects at the tumor site (41). In support of this, Liu et al. (42) showed that tumor infiltration and lysis by NK cells led to the release of tumor Ag and enhanced uptake and presentation of tumor Ag by conventional DC.

Our findings argue against a direct cytotoxic role for NK cells stimulated by DC immunization: NK cells isolated directly ex vivo from lymph nodes draining DC-immunized sites were not cytotoxic against B16 (at least in vitro), nor did restimulated lymph node NK cells show upregulated granzyme B or CD107a expression. Although a convincing role for IFN-γ in DC immunotherapy was shown, perforin was dispensable for the observed antitumor response following DC immunotherapy. In an earlier study (34), we showed that stimulation of splenic NK cells with DC plus Gram-positive bacteria led to rapid NK cell cytokine production, but without enhanced cytotoxic activity against YAC-1 or Ab-sensitized B cell lymphoma target cells. Another potential mechanism for NK cell contribution to anticancer responses is the perforin-dependent lysis of tolerogenic DC in the draining lymph nodes (43). However, a lack of perforin-dependence for NK cell activity in our study suggests that this mechanism is not involved in our particular vaccination setting. Despite the fact that IFN-γ did not suppress the growth of our B16-OVA subline in vitro, it did lead to an upregulation of MHC-I and MHC class II on the B16 tumor, as previously shown by Böhm et al. (32), suggesting that IFN-γ could enhance the presentation of tumor-associated Ags.

It is also unlikely that NK cells are independently involved in tumor lysis or growth inhibition, because administration of SSa-DC in the absence of tumor Ag did not induce an observable antitumor response (Fig. 3B). In contrast, some studies have demonstrated that immunization with syngenic DC alone (in the absence of tumor Ag) can induce NK cell–dependent antitumor responses against melanoma, colon, or lung carcinoma (17, 20, 23). We have previously noted that immunization with syngenic DC in the absence of tumor Ag can induce antimelanoma immunity in the prophylactic setting, but only if FCS Ags are present in both the DC and the melanoma culture medium (37, 39). Unless rigorous attempts are made to exclude FCS from all steps of either DC (37) or melanoma (39) culture, anti-FCS responses could be mistaken for the action of the innate immune response, including NK cell activation.

NK cells are highly effective at limiting the formation of B16 lung metastases, but only exert this antimetastatic effect in the first 6 h following i.v. administration (24). This suggests either that NK cells kill tumor cells lodged in the vasculature of target organs (24) or that melanoma cells acquire membrane expression of MHC-I and become NK cell resistant a few hours following implantation (44, 45). However, in our model, the vascular access of NK cells to s.c.-implanted tumor cells likely limits the role of NK cells in controlling tumor development. In addition, NK cells at distinct anatomic locations inhibit B16 using different mechanisms. For example, liver NK cells, but not lung NK cells, rely on perforin-dependent pathways for destruction of B16 (46).

In our study, cultured B16-OVA demonstrated a basal level of MHC-I expression that was markedly upregulated upon IFN-γ treatment (Fig. 8) (32). Even in the absence of vaccination, MHC-I is upregulated in an IFN-γ–dependent fashion on implanted B16 tumor cells within hours, and this expression is maintained during in vivo growth (44, 47). IFN-γ also leads to NKG2D ligand downregulation on B16 (48). These features may protect B16 from direct NK cell cytolysis. It is also noteworthy that NK cells isolated directly ex vivo are only poorly cytotoxic against B16, unless isolated from hosts systemically activated with polyinosinic-polycytidylic acid or CpG (46, 49).

Microbial products create an inflammatory environment that inhibits tumor growth and vasculogenesis, with key players in this process identified as IP-10, IFN-γ, IL-1, IL-7, and TNF-α (10–12, 50, 51). The streptococcal preparation used in this study was optimized in our laboratory following a screen of a panel of Gram-positive and -negative bacteria. Superior induction of IFN-γ from splenocyte cocultures was achieved with Gram-positive preparations, and the activity was shown to be dependent on NK cells (34). The particular strain of S. salivarius selected to enhance DC activation for tumor immunotherapy has the additional advantage of a lack of mammalian toxin sequences, a demonstrated lack of pathogenicity, and is widely used in humans as an oral probiotic (52), yet is a potent inducer of IL-12 and IFN-γ in human and murine leukocytes (33, 34). Although bacteria induce potent activation of DC enabling optimal T cell stimulation (53, 54), we cannot rule out that host responses to bacterial products associated with S. salivarius–matured DC may also contribute to the potency of the observed immune response. In support of this, Sato and colleagues (55, 56) have shown that the host immune response to OK-PSA, the LTA component of the OK-432 (penicillin-killed Streptococcus pyogenes) vaccine, was dependent on TLR4 expression on both immunizing DC and the host APCs. In our study, both intact Gram-positive bacteria and LTA components were found to be the more effective adjuvants for DC vaccination in a therapeutic setting compared with other TLR agonists. The enhanced adjuvant effect of Gram-positive preparations in our study and that of others (55, 56) may reflect the superior induction of inflammatory cytokines by Gram-positive bacteria, without the induction of IL-10 observed with Gram-negative bacteria (33, 57).

Our data demonstrated that following i.v. depletion by ASGM, NK cells were not fully regenerated by 60 d. It is possible that the rabbit anti-ASGM Abs have a very long t1/2 in vivo, but more likely reflects the slow turnover of NK cells in vivo. The regenerative pathway of NK cells following depletion is still not fully resolved (58); however, Hummel et al. (59) found that the reconstitution of bone marrow NK cells may take as long as 8 wk following a single i.p. injection with ASGM.

In addition to the role of NK cells in DC immunotherapy, our data also suggested a role for CD4 T cells and, to a lesser extent, CD8 T cells in the antitumor response. A requirement for both T cell subsets was recently demonstrated in a prophylactic setting using therapy of DC pulsed with apoptotic B16 melanoma cells (60). In our study, although CD4 or CD8 T cells may be required for optimal antitumor responses, the loss of NK cells resulted in a more profound defect in tumor immunity. Although NK cell depletion did not affect transgenic CD4 or CD8 T cell proliferation (Supplemental Fig. 1), we cannot rule out that NK cells provide a helper function to endogenous T cells following DC immunotherapy. Karimi et al. (18) examined the various contributions of CD4, CD8 T cells, and NK cells to B16 tumor immunity. They found that both CD8 T cells and NK cells were required for tumor protection; however, they used a DC/SIINFEKL (MHC-I–restricted) vaccine that might not uncover a role for CD4 T cells in the observed antitumor response. In another study, the benefit of an adenovirus expressing murine IL-12 plus anti–4-1BB (CD137) therapy combination was lost following depletion of NK cells or CD8 T cells, but not CD4 T cells (61). Interestingly, Prins et al. (20) found that in the absence of CD8 T cells, CD4 T cells may compensate for the loss of CTL in the inhibition of melanoma brain metastases. Together with our data, these studies show that NK cells cooperate with T cells to enhance antitumor response and emphasize the importance of optimizing NK cell activation for tumor immunotherapeutic protocols.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank Roslyn Kemp, Michelle Wilson, John Schofield, Frank Griffin, and Vernon Ward for support, Franca Ronchese for providing mouse strains and IFN-γ–neutralizing Ab, and Geoff Hill and Kelli MacDonald for providing the CD8β Ab. We also thank John Tagg for providing reagents and the S. salivarius K12 strain.

Footnotes

  • This work was supported by a University of Otago Ph.D. scholarship (to A.L.B.), a Lottery Health Ph.D. scholarship (to S.C.S.), a University of Otago Research grant, and the Otago School of Medical Sciences Dean’s Bequest fund.

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    ASGM
    anti–asialo GM1
    DC
    dendritic cell
    DC-OVA
    OVA-pulsed DC
    ifnγ−/−
    IFN-γ–deficient
    LTA
    lipoteichoic acid
    MHC-I
    MHC class I
    pfp−/−
    perforin-deficient.

  • Received October 5, 2012.
  • Accepted December 22, 2013.

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