Cellular and molecular requirements for rejection of B16 melanoma in the setting of regulatory T cell depletion and homeostatic proliferation

Abstract

We have recently demonstrated that adoptive transfer of regulatory T cell (Treg) - depleted polyclonal T cells into lymphopenic mice leads to rejection of B16 melanoma, which generated an opportunity to study host requirements for tumor rejection when it effectively occurred. CD8⁺ T cell priming and tumor rejection required tumor antigen cross-presentation, as evidenced by tumor outgrowth in Kb^−/− bone marrow chimeric or B71/2^−/− mice. CD4⁺ T cells were additionally required for optimal tumor control, thought not through classical CD4 “help”, as the frequency of primed CD8⁺ T cells was similar in the absence of CD4⁺ T cells, and tumor rejection did not depend upon CD40/CD40L interactions or on IL-2 production by CD4⁺ T cells. Rather, CD4⁺ T cells appeared to act at the effector phase of tumor rejection and responded to B16-derived antigens in vitro. At the effector phase, IFN-γ production by transferred T cells, but not host cells, was necessary. IFN-γ acted either on host or tumor cells, and was associated with reduced tumor vascularity. Finally, tumor rejection occurred following transfer of TNF-α, perforin or FasL-deficient T cells. However, perforin/FasL double knockout T cells failed to reject, arguing that that the killing of B16 melanoma cells could occur either via the cytotoxic granule or Fas pathways. Collectively, these results support a model in which host tumor antigen cross-presentation primes adoptively-transferred T cells, which remain functional in the setting of homeostatic proliferation and Treg depletion, and which promote tumor rejection via IFN-γ and lysis via cytotoxic granules and/or FasL.

INTRODUCTION

Recent advances in the efficacy of adoptive T cell therapy have resulted in high rates of objective tumor responses in patients with advanced melanoma. Improvements in the ability to culture and expand tumor infiltrating lymphocytes (TIL) ex vivo, and the addition of chemotherapy or total body irradiation (TBI) conditioning regimens to deplete host T cells and Treg prior to the adoptive transfer of ex vivo activated TIL have resulted in objective tumor response rates exceeding 50% in published reports (1). Currently, several groups have attempted to improve upon these observations through transduction of host T cells with T cell receptors (TCR) specific for known melanoma-derived antigens with some success (2). While a thorough investigation of the antigen-specificity, differentiation state and phenotype of ex vivo expanded T cells ideal for adoptive transfer has been undertaken, little is known regarding either the host requirements or optimal effector functions of adoptively-transferred T cells which are necessary for tumor regression to occur in vivo.

The tumor microenvironment can exploit a number of mechanisms which can inhibit the execution of effective host anti-tumor immune responses (3), which has pointed towards strategies to reverse these immune suppressive pathways and restore immune-mediated tumor control. We have previously reported a murine adoptive T cell therapy model in which tumor-induced T cell anergy was reversed by transfer of polyclonal T cells into lymphopenic hosts, through the process of homeostatic proliferation (HP) (4). Concomitantly, regulatory T cells were depleted using an anti-CD25 mAb-mediated approach prior to adoptive transfer. When Treg-depleted T cells were transferred into lymphopenic hosts (either RAG^−/− or sublethally-irradiated C57BL/6 mice), potent rejection of B16 melanoma ensued, which correlated with robust, persistent IFN-γ production by tumor antigen-specific CD8⁺ T cells (4). This powerful rejection of B16 melanoma provided a rare opportunity to determine the host for requirements for rejection of this tumor which is normally difficult to control in vivo. Although this model does not make use of ex-vivo activated TIL or tumor antigen-specific TCR transgenic T cells, it is an approach which is straightforward and clinically translatable.

While the major role of homeostatic proliferation in this approach appears to be to prevent anergy of anti-tumor T cells, the process of homeostatic proliferation itself has been reported to promote transition of naïve T cells into pseudo-memory phenotype cells (5). Therefore, it was not clear if antigen-specific T cell priming might still be required for productive anti-tumor immunity. Cross-priming via tumor antigen released to host dendritic cells could theoretically still occur and, superimposed upon homeostatic proliferation, preferentially expand tumor antigen-specific effector T cells for tumor control.

In addition to priming of CD8⁺ T cells, which are necessary for tumor eradication, the role of polyclonal CD4⁺ T cells in responding directly to B16 melanoma antigens has not been well elucidated. Previous work has suggested that CD4⁺ T cells contribute to the anti-tumor immune response through several mechanisms, , including providing help during the priming phase of CTL via production of IL-2 (6), licensing of APC through CD40/CD40L interactions (7, 8), production of IFN-γ in the tumor microenvironment (9), and direct tumor cell lysis (10–12). More recently, several groups have demonstrated that TCR transgenic (Tg) CD4⁺ T cells specific for MHC class II-restricted antigens expressed by B16 melanoma can mediate tumor regression (10, 12, 13). While these data are important, they may not translate well into the clinical setting in which polyclonal adoptively-transferred CD4⁺ T cells might also be required to provide classical “help” to antigen-specific CD8⁺ T cells or license APC through CD40L, in addition to directly recognizing and eliminating malignant cells.

Finally, during the effector phase of the anti-tumor immune response, the mechanism of elimination of B16 melanoma tumors in the setting of adoptive T cell transfer was of interest to elucidate. It was conceivable that production of selected cytokines and cytolytic molecules by adoptively-transferred Treg-depleted T cells would be required in order for tumor regression to occur. This could involve actions directly on tumor cells but also indirect effects on stromal cell components. For example, the necessity of IFN-γ production by tumor-specific T cells has previously been shown to be required for tumor rejection in numerous models (14–18). But IFN-γ can act through several, distinct mechanisms, such as inducing MHC upregulation on tumor cells and APCs (14), an anti-proliferative effect on tumor cells in combination with TNF-α (19), activation of innate immune cells, such as macrophages (20), or through induction of molecules which inhibit angiogenesis (15, 21, 22). With these questions in mind, the host requirements for rejection of B16 melanoma by adoptively transferred Treg-depleted T cells in lymphopenic recipient mice were explored.

MATERIALS AND METHODS

Mice and tumor cell lines

C57BL/6 mice were purchased from Jackson Laboratories (Bar Harbor, ME) or Taconic laboratories (Germantown, NY). RAG2^−/−, Kb^−/− and Db^−/− mice on the C57BL/6 background were purchased from Taconic laboratories. RAG1^−/−, B71/2^−/−, CD4^−/−, CD8^−/−, CD40^−/−, CD40L^−/−, IL-2^−/−, IFN-γ^−/−, IFN-γR^−/−, TNF-α^−/−, perforin^−/− and gld/gld mice, all on the C57BL/6 background, were purchased from Jackson laboratories. Perforin^−/−×gld/gld mice were interbred in our animal facility. Mice were housed in micro-isolator cages in a specific pathogen-free barrier facility and treated under NIH guidelines. Animals were maintained and used according to protocols approved by the institutional animal care and use committee of the University of Chicago. RAG1^−/− and RAG2^−/− mice were used as available by our suppliers, and we have previously found that B16.SIY rejection occurs similarly in either strain following adoptive transfer of Treg-depleted T cells (J.K., unpublished observation). Because RAG^−/− mice contain no endogenous mature T cells to confound results, they were used as hosts in experiments investigating the role of CD8⁺ versus conventional CD4⁺ T cells in tumor rejection, and also when T cell-produced effector/lytic molecules (IL-2, IFN-γ, TNF-α, perforin, FasL) were examined. When evaluating host (i.e. non-T cell) factors required for tumor rejection, irradiated, targeted gene knock-out mice on the C57BL/6 background (Kb^−/−, B7.1/B7.2^−/−, CD40^−/−, IFN-γ^−/−, IFN-γR−/−) were utilized, as these knock-out strains are not commercially available on the RAG^−/− background.

B16.F10 is a widely-utilized, poorly-immunogenic and aggressive melanoma cell line. B16.F10 cells were engineered to express GFP fused in frame with the model antigen SIYRYYGL (SIY; B16.SIY) which can be recognized by CD8⁺ T cells in the context of K^b (23). The B16.SIY cell line is useful because it enables monitoring of SIY antigen-specific T cell responses in tumor-bearing hosts using IFN-γ ELISPOT, as well as SIY-pentamer staining and flow cytometry. Expression of the SIY antigen renders B16 cells slightly more immunogenic, and thus B16.SIY tumors grow more slowly (but are not rejected) in syngeneic C57BL/6 hosts compared with the parental B16.F10 cell line (4). The dominant-negative IFN-γR construct (ΔIFN-γR) containing the cDNA for a truncated, non-signaling form of the IFN-γR, has been previously reported (14), and was a kind gift from Dr. Robert Schreiber (Washington University, St. Louis, MO). The ΔIFN-γR cDNA was subcloned into the pMY retroviral construct, and transduced into B16.SIY (SIY.DSRED) cells to generate the B16.SIY.ΔIFN-γR tumor cell line. A control empty-vector pMY construct was similarly transduced into B16.SIY (SIY.DSRED) cells to generate the B16.SIY.EV tumor cell line.

T cell purification and adoptive transfer

Cell suspensions were generated from spleens of indicated donor mice and total CD4⁺ and CD8⁺ T cells were purified by negative selection over magnetic columns using a T cell enrichment kit (Miltenyi Biotechnologies) according to the manufacturer’s protocol. Flow cytometric analysis was periodically performed and routinely showed greater than 95% purity of CD3⁺ T cells. Purified total splenic CD4⁺ and CD8⁺ T cells were additionally depleted of CD25⁺ cells (Treg) by negative selection using a magnetic-bead conjugated anti-CD25 antibody (Miltenyi Biotechnologies). CD25 depletion was also confirmed by flow cytometry to eliminate > 95% of CD25⁺ T cells from total T cell population. For transfer in vivo, purified T cells were washed 3 times with PBS, and then re-suspended at a concentration of 10⁸ cells/ml. For adoptive transfer experiments, a volume of 0.1 ml was injected into the lateral tail vein of mice.

Tumor challenge and measurement

After washing B16.SIY tumor cells 3 times with PBS to remove FCS, they were re-suspended in PBS at a concentration of 10⁷ cells/ml. A volume of 0.1 ml (10⁶ tumor cells) was injected subcutaneously into the lower right abdomen of each mouse. Tumor measurements were performed 2–3 times weekly with calipers by a single individual taking the greatest tumor diameter and its perpendicular to determine an average. Mice with mean tumor diameters greater than 20 mm, or those demonstrating ulceration or showing signs of declining health, were euthanized.

Generation of bone marrow chimeras

Cohorts of C57BL/6 mice were lethally-irradiated with 1000rad TBI. Bone marrow cells were harvested from femurs of C57BL/6, Kb^−/− or Db^−/− mice, and 10⁷ cells were injected into the tail veins of irradiated C57BL/6 mice 1 day following TBI. After 11 weeks, bone marrow chimeric mice received 600rad TBI to induce host lymphodepletion. 24 hours later, mice received adoptive transfer of 10⁷ CD25-depleted T cells isolated from spleens of C57BL/6 mice, followed the next day, by a subcutaneous challenge with 10⁶ B16.SIY cells.

IFN-γ ELISPOT

ELISPOT was conducted with the BD PharMingenTM (San Diego, CA) mouse IFN-γ kit according to provided protocol. Briefly, ELISPOT plates were coated with anti-mouse IFN-γ antibody and stored overnight at 4°C. Plates were then washed and blocked with DMEM supplemented with 10% FCS for 2 hours at room temperature. Splenocytes from tumor-challenged mice were harvested at various indicated time points and plated at 10⁶ cells/well. Stimulation was performed with irradiated B16.SIY tumor cells (10,000rad) at 5×10⁴ cells/well, 80nM SIY peptide, or PMA and ionomycin as a positive control. Plates were stored at 37°C in a 7.5% CO₂ incubator overnight, washed, and coated with detection antibody for 2 hours at room temperature. They were again washed and coated with avidin-peroxidase for 1 hour at room temperature. Plates were then washed and developed by addition of AEC substrate. Developed plates were dried overnight, read using an ImmunoSpot® Series 3 Analyzer, and analyzed with ImmunoSpot® software.

Tetramer staining and FACS analysis

The SIY (SIYRYYGL) and negative control OVA (SIINFEKL) peptide pentamers were purchased from Proimmune (Oxford, UK). For cell staining, the manufacturer’s protocol was followed. Antibodies against the following molecules coupled to the indicated fluorochromes were purchased from BD PharMingen: FITC-anti-CD3, APC-anti-CD8α, PerCP-Cy5.5-anti-CD4, PerCP-Cy5.5-anti-B220, PE-anti-CD25 (PC61), Biotin-anti-K^b, Biotin-anti-D^b. APC-anti-FoxP3 (FJK-16s) was purchased from e-Bioscience (San Diego, CA). FACS analysis was performed on the FACScanto cytometer using BD FACSDiva software (Franklin Lakes, NJ). Data analysis was performed using FlowJo software (Tree Star, Inc Ashland, OR).

Tumor vascular density measurement

B16.SIY tumors were established for 6 days in either irradiated C57BL/6 or IFN-γ^−/− hosts. On day 6, tumor-bearing mice received 600rad TBI. On day 8, 10⁷ CD25-depleted T cells from C57BL/6 or IFN-γ^−/− mice were transferred into tumor-bearing, irradiated C57BL/6 or IFN-γ^−/− hosts, respectively. Approximately 2 weeks later, mice were injected with 100 ug of biotinylated tomato lectin (Vector Laboratories, Burlingame, CA) and euthanized 5 minutes later. Tumors were surgically excised and placed in 4% paraformaldehyde at 4° C for 4 hours, and then in 30% sucrose/PBS overnight at 4° C. Tumor specimens were embedded in 7.5% gelatin/15% sucrose/PBS and flash frozen in liquid nitrogen cooled isopentane at −65° C. 8 micron sections were cut and stored at −80° C until use. For microvessel density analyses, tumor sections were warmed to room temperature and gelatin removed by incubation in PBS at 37° C. Sections were washed twice with room temperature PBS, stained with SA-Alexafluor 594 (Molecular probes) for 20 minutes at room temperature, and then counterstained with Vectashield with DAPI (Vector Laboratories). Stained sections were imaged using an Axiovert200 (Zeiss) microscope. Vessel density was measured using ImageJ (NIH).

Statistical analysis

The student’s t test was utilized to compare differences in tumor size between mice in various treatment arms at the indicated time points, and also to analyze differences in numbers of IFN-γ spot-forming cells and SIY/K^b tetramer-reactive CD8⁺ cells in individual mice assigned to various treatment groups. A p-value of ≤ 0.05 between groups was considered to be statistically significant.

RESULTS

SIY-antigen cross-presentation by host hematopoietic cells is necessary for B16.SIY tumor rejection following adoptive transfer of CD25-depleted T cells

Our previous results have demonstrated that adoptive transfer of polyclonal CD4⁺ and CD8⁺ T cells depleted of CD25⁺ Treg (hereafter referred to as CD25-depleted T cells) into lymphopenic hosts (either RAG2^−/− or sub-lethally irradiated (600rad) C57BL/6 mice) resulted in robust homeostatic T cell expansion which resulted in equivalently-potent rejection of B16.SIY tumors (4), although homeostatic T cell proliferation occurred somewhat more slowly in irradiated C57BL/6 versus RAG2^−/− hosts. Low-dose TBI has been utilized as a lymphodepletion strategy to mimic what might be employed for clinical translation. (4). Following adoptive transfer of CD25-depleted T cells into lymphopenic recipients, tumor rejection occurred in both the prophylactic setting (100% rejection) and also when B16.SIY tumors were established to approximately 5–6 mm in mean tumor diameter prior to transfer of CD25-depleted T cells (Supplementary Figure 1).

Because the process of homeostatic proliferation can drive naïve T cells to acquire a partially activated phenotype (5), it was of interest to determine whether the activation of antigen-specific CD8⁺ T cells in this context would still require tumor antigen cross-presentation by host antigen-presenting cells (APCs). To this end, we generated radiation bone marrow chimeric mice using K^b-deficient donor bone marrow cells, as K^b is the restricting element for the dominant SIY antigen. Eleven weeks later, chimeric mice received 600rad TBI for lymphodepletion and adoptive transfer of CD25-depleted T cells, followed by B16.SIY cell challenge. D^b−/− bone marrow chimeras were included as a control, to ensure that any differences in tumor rejection observed were not due to reduced total expression of class I MHC molecules in K^b−/− chimeric hosts, for example to support homeostatic T cell proliferation. As shown in Figure 1A (top and bottom), K^b and D^b expression were not detected on WBC from K^b−/− → C57BL/6 and D^b−/− → C57BL/6 chimeric mice, respectively. Similar numbers of CD3⁺ T cells were present in spleens of all groups of chimeric mice, suggesting that T cell expansion was supported in all cases (data not shown). As shown in Figure 1B, B16.SIY tumors were rejected, as expected, in lymphopenic C57BL/6 → C57BL/6 and D^b−/− → C57BL/6 chimeras following adoptive T cell transfer. In contrast, in K^b−/− → C57BL/6 chimeras, tumors grew progressively, though slightly more slowly than in chimeric mice that did not receive a T cell transfer, suggesting that some degree of antigen-specific immunity was generated in the K^b−/− → C57BL/6 chimeras. This possibly occurred through recognition of other melanoma antigens on B16 cells, as we and others have previously demonstrated (4, 23). Overall, these results indicate that cross-presentation of antigen on host K^b-expressing hematopoietic cells is necessary for B16.SIY rejection by CD25-depleted T cells in the context of homeostatic proliferation.

Figure 1. SIY antigen cross-presentation by host hematopoietic cells is necessary for B16.SIY rejection.

A. To generate bone marrow chimeras, groups of C57BL/6 mice received 1000rad TBI, followed by infusion of 10⁷ bone marrow cells from C57BL/6, K^b−/− or D^b−/− donors the following day. Approximately 9 weeks following generation of bone marrow chimeric mice, K^b and D^b expression were analyzed on WBC by flow cytometry on peripheral blood samples. Histograms represent K^b or D^b expression on cells in the WBC gate. Solid, dotted and dashed histograms represent K^b (top) or D^b (bottom) expression on WBC from control, Kb^−/− and Db^−/− chimeras, respectively. B. Eleven weeks following generation of chimeric mice (7 per group), each received 600rad TBI, followed by infusion of 10⁷ CD25-depleted T cells purified from spleens of C57BL/6 donor mice. First, total splenic CD4⁺ and CD8⁺ T cells were purified, followed by depletion of CD25⁺ cells. Controls (3 per group) received 600rad TBI, but no T cell infusion. 24 hours later, mice were inoculated subcutaneously with 10⁶ B16.SIY cells, and tumor growth was monitored 2–3 times weekly with calipers. Mean tumor diameter was recorded by taking the average of the bi-perpendicular tumor diameters. *p = < 0.01 for comparison of tumor size on day 37 between K^b−/− and both D^b−/− and B6 chimeric mice following transfer of CD25-depleted T cells. The tumor growth curve is representative of 2 independent experiments each with between 3 (controls) and 7 mice/group.

T cell costimulation through B7-1/B7-2 on host cells is required for tumor rejection by CD25-depleted T cells in the context of homeostatic proliferation

If encounter of transferred T cells with host APCs was necessary for antigen presentation, then it seemed likely that costimulatory signals from host APCs might also be required for T cell priming and tumor rejection. To test this notion, C57BL/6 or B7-1/B7-2^−/− mice were lymphodepleted with 600rad TBI followed by transfer of CD25-depleted T cells from C57BL/6 mice and B16.SIY tumor cell challenge. While B16.SIY tumors were completely rejected in irradiated C57BL/6 mice following a CD25-depleted T cell transfer, they grew progressively in irradiated B7-1/B7-2^−/− hosts (Figure 2A). To assess whether T cell priming paralleled tumor control, SIY/K^b pentamer staining and IFN-γ ELISPOT were performed on spleens harvested approximately 24 days following B16.SIY tumor cell inoculation (Figure 2B and 2C). Consistent with the differences in tumor growth, numbers of SIY-specific CD8⁺ T cells were approximately 2-fold higher in irradiated C57BL/6 versus irradiated B7-1/B7-2^−/− hosts following transfer of CD25-depleted T cells (p=0.05) (Figure 2B upper and lower panels). More strikingly, a 7-fold increase in the number of IFN-γ-producing, SIY-specific CD8⁺ T cells was observed in irradiated C57BL/6 compared with irradiated B7-1/B7-2^−/− mice following T cell transfer (p < 0.001) (Figure 2C). Collectively, these results demonstrate the importance of host B7-1 and/or B7-2 costimulation in promoting the priming and subsequent effector function of tumor antigen-specific CD8⁺ T cell responses in this system.

Figure 2. B7-1/B7-2 costimulation of adoptively-transferred CD25-depleted T cells by host cells is required for B16.SIY rejection.

A. C57BL/6 or B7-1/B7-2^−/− mice received 600rad TBI on day 0. On day 1, 10⁷ CD25-depleted T cells were adoptively-transferred into irradiated mice (a group of irradiated C57BL/6 mice did not receive T cell transfer, serving as controls). 24 hours later, mice were inoculated subcutaneously with 10⁶ B16.SIY cells, and tumor growth was monitored. *p = < 0.01 for comparison of tumor size on day 24 between irradiated B7-1/B7-2^−/− and irradiated C57BL/6 mice following transfer of CD25-depleted T cells. B. 24 days following B16.SIY inoculation, spleens were harvested from tumor-challenged mice and analyzed by flow cytometry for the percentage of CD8⁺SIY⁺ cells among the CD8⁺ T cell population present in individual mice (p = 0.05 for comparison of numbers of SIY-reactive CD8⁺ T cells in irradiated C57BL/6 versus irradiated B7-1/B7-2^−/− mice receiving CD25-depleted T cells (upper panel). Representative SIY and negative control OVA (SIINFEKL peptide) tetramer analysis from individual mice (lower panel). FACS plots demonstrate the percentage of SIY or OVA-reactive CD8⁺ T cells after gating on CD4⁻B220⁻ and CD8⁺ cells. C. IFN-γ ELISPOT. On day 24 following B16.SIY implantation, spleens were harvested from tumor-challenged mice and 10⁶ spleen cells from individual mice in each group were restimulated in vitro overnight with media or SIY peptide (80nM). An IFN-γ ELISPOT assay was performed and results are recorded as spot-forming cells per 10⁶ total cells in individual mice (3 wells per mouse per condition) following SIY restimulation (*p < 0.001 for the comparison of IFN-γ spot-forming cells in irradiated C57BL/6 versus irradiated B7-1/B7-2^−/− mice following adoptive T cell transfer). These results are representative of 2 independent experiments with 5–7 mice/group.

Conventional CD4⁺ T cells are required for complete rejection of B16.SIY tumors in lymphopenic mice following CD25-depleted T cell transfer

CD4⁺ T cells have been reported to contribute to anti-tumor immune responses through several mechanisms. CD4⁺ T cells have been shown to provide “help” for induction of tumor antigen-specific CD8⁺ effector T cells, either through production of IL-2 or via licensing of APCs through CD40/CD40L interactions (6, 7). They also have been shown to promote memory CD8⁺ T cell development (24, 25). However, recent evidence has indicated that CD4⁺ T cells can also act at the effector phase (13), and in some cases directly recognize and kill tumor cells (10, 11). In the human setting, adoptive transfer of tumor antigen-specific CD4⁺ T cell clones can mediate complete tumor rejection in at least a subset of melanoma patients (26). Thus, the role of conventional CD4⁺ T cells in supporting B16.SIY rejection by CD25-depleted T cells was examined. Several T cell populations were prepared prior to adoptive transfer into RAG^−/− hosts: 1) CD25-depleted T cells, consisting of CD8⁺CD25⁻ and CD4⁺25⁻ T cells from spleens of C57BL/6 donor mice, 2) CD8⁺ T cells from the spleens of CD4^−/− donor mice, and 3) CD4⁺25⁻ T cells from the spleens of CD8^−/− donor mice. CD4^−/− and CD8^−/− rather than wildtype C57BL/6 mice were chosen for these experiments because the isolation of CD4⁺ or CD8⁺ T cells from wildtype mice is imperfect, and even a few contaminating T cells can undergo robust homeostatic expansion following adoptive transfer into lymphopenic hosts and potentially confound results. As demonstrated in Figure 3A, RAG1^−/− mice that received only CD8⁺ T cells temporarily controlled B16.SIY outgrowth, but failed to achieve complete tumor rejection, suggesting an important role for conventional CD4⁺ T cells in this model. Interestingly, transfer of CD4⁺25⁻ T cells in the absence of CD8⁺ T cells resulted in a modest delay in the outgrowth of B16.SIY tumors, pointing toward a potential role for conventional CD4⁺ T cells in the direct recognition and control of B16.SIY tumor cells.

Figure 3. Complete rejection of B16.SIY is dependent upon conventional CD4⁺ T cells.

A. T cells were purified from spleens of C57BL-6 mice (CD25-depleted T cell group), CD4^−/− mice (CD8⁺ T cell group) or CD8^−/− mice (CD4⁺25⁻ T cell group). CD25⁺ T cells were subsequently depleted using anti-CD25 microbeads in the CD25-depleted T cell and CD4⁺25⁻ T cell groups. 10⁷ CD25-depleted T cells, CD8⁺ T cells or CD4⁺25⁻ T cells were adoptively-transferred into cohorts of RAG1^−/− recipient mice. 24 hours later, mice were challenged with 10⁶ B16.SIY cells subcutaneously, and tumor growth was monitored. *p < 0.01 for comparison of tumor size at day 21 between RAG1^−/− mice following adoptive transfer of CD25-depleted T cells versus both CD8⁺ and CD4⁺25⁻ T cells. ^#p = 0.02 for comparison of tumor size at day 21 between RAG1^−/− mice following adoptive transfer of CD8⁺ and CD4⁺25⁻ T cells. B. 25 days following B16.SIY inoculation, spleens were harvested from tumor-challenged mice and analyzed by flow cytometry for the percentage of CD8⁺SIY⁺ cells among the CD8⁺ T cell population present in individual mice (p = n.s. for the comparison of SIY-reactive CD8⁺ T cells in RAG1^−/− mice receiving CD25-depleted T cells versus RAG1^−/− mice receiving CD8⁺ T cells. C. SIY and negative control OVA (OT-1) tetramer analysis of spleen cells from individual mice in B. FACS plots demonstrate the percentage of SIY or OVA-reactive CD8⁺ T cells after gating on CD4⁻B220⁻ and CD8⁺ cells. D and E. On day 25 following B16.SIY implantation, spleens from tumor-challenged mice were harvested and 10⁶ spleen cells from individual mice in each group were restimulated in vitro overnight with media, SIY peptide (80nM) or 5×10⁴ irradiated B16.SIY cells (10,000rad). An IFN-γ ELISPOT assay was performed and results are recorded as spot-forming cells per 10⁶ total cells in individual mice (3 wells per mouse per condition) following media, SIY peptide (C.) or irradiated B16.SIY cell (D.) restimulation (*p < 0.001 for the comparison of IFN-γ spot-forming cells in RAG1^−/− mice receiving CD25-T cells versus RAG1^−/− mice receiving CD8⁺ or CD4⁺25⁻ T cells when restimulated with irradiated B16.SIY cells. ⁺p = n.s. for the comparison of IFN-γ spot-forming cells in RAG1^−/− mice receiving CD8⁺ versus CD4⁺25⁻ T cells when restimulated with irradiated B16.SIY cells. ^#p = n.s. for the comparison of IFN-γ spot-forming cells in RAG1^−/− mice receiving CD8⁺ versus CD25-T cells when restimulated with SIY peptide. These results are representative of 2 independent experiments with 4–5 mice/group.

The mechanism by which CD4⁺ T cells might be contributing to tumor control was then investigated. To determine whether the presence of conventional CD4⁺ T cells provided help for the priming of SIY-specific CD8⁺ T cells in B16.SIY tumor-challenged mice, spleen cells were stained with SIY/K^b tetramers (Figure 3B and C) and analyzed by IFN-γ ELISPOT (Figure 3D and E). Surprisingly, similar percentages of SIY⁺CD8⁺ T cells with equivalent IFN-γ production following SIY peptide restimulation were present in spleens of tumor-challenged RAG1^−/− mice that had received either CD8⁺ or CD25-depleted T cells, arguing that the effect of CD4⁺25⁻ T cells on B16.SIY rejection was not primarily through provision of classical “help” to CD8⁺ T cells (Figure 3B and C). Furthermore, a significant number of IFN-γ spot-forming cells were detected in spleens of RAG1^−/− mice that received only CD4⁺25⁻ T cells following restimulation with irradiated B16.SIY cells (Figure 3E), most likely through cross-presentation of MHC class II-restricted antigens derived from dying B16.SIY cells to CD4⁺25⁻ T cells, as B16.SIY cells do not express MHC class II molecules in vitro. This result suggests that CD4⁺ T cells were naturally primed in vivo and could recognize antigens derived from B16.SIY tumors.

IL-2 production and provision of CD40L by CD4⁺25⁻ T cells are dispensable for B16.SIY rejection

Although the presence of conventional CD4⁺ T cells did not appear to significantly increase the priming of SIY-specific CD8⁺ T cells, the results presented above did not formally exclude the possibility that classical mechanisms of CD4⁺ T cell help might contribute to tumor rejection. Therefore, the key defined mechanisms of CD4⁺ T cell help, such as IL-2 production, and provision of CD40L were directly interrogated. First, the role of secreted IL-2 was examined. Wildtype CD8⁺ T cells (3.5 × 10⁶) were combined with CD4⁺25⁻ T cells (6.5 × 10⁶) from either wildtype or IL-2^−/− mice and adoptively-transferred into RAG2^−/− hosts, followed by tumor challenge. Surprisingly, B16.SIY tumors were rejected whether or not the transferred CD4⁺ T cells were capable of IL-2 production (Figure 4A). At the same time, we examined whether IL-2 by transferred T cells was required at all for tumor rejection in this model. Interestingly, when CD25-depleted total T cells from IL-2^−/− mice were utilized for adoptive transfer, no effective tumor control was observed (Figure 4B), and resulted in a marked reduction in SIY-specific T cell responses (Supplementary Figure 2A and B). Together, these data imply that while IL-2 is important for optimal priming of CD8⁺ tumor antigen-specific T cells in this model, it does not need to be provided by CD4⁺ T cells and presumably can be sufficient when produced by the CD8⁺ T cells themselves.

Figure 4. IL-2 production and CD40L provision by conventional CD4⁺ T cells are dispensable for B16.SIY rejection.

A. CD8⁺ and CD4⁺ T cells were isolated separately from spleens of C57BL/6 mice, and CD4⁺ T cells were also isolated from spleens of IL-2^−/− mice. Purified CD4⁺ T cells from C57BL/6 or IL-2^−/− mice were further depleted of CD25⁺ cells, and 3.5 × 10⁶ C57BL/6 CD8⁺ T cells, along with 6.5 × 10⁶ C57BL/6 or IL-2^−/− CD4⁺25⁻ T cells (total of 10⁷ T cells) were adoptively-transferred into groups of 5 RAG2^−/− recipient mice, subsequently challenged with B16.SIY cells, and tumor growth was monitored (p = n.s. for comparison of tumor size on day 26 between RAG2^−/− mice following adoptive transfer of wildtype CD8⁺ and CD4⁺25⁻ T cells versus wildtype CD8⁺ and IL-2^−/− CD4⁺25⁻ T cells). B. CD25-depleted T cells were purified from spleens of C57BL-6 or IL-2^−/− mice, and 10⁷ cells were adoptively-transferred into cohorts of RAG1^−/− recipient mice. 24 hours later, mice were challenged with 10⁶ B16.SIY cells as in A. (*p < 0.01 for comparison of tumor size on day 26 between RAG2^−/− mice following adoptive transfer of wildtype versus IL-2^−/− CD25-depleted T cells). C. Groups of 5 C57BL/6 or CD40^−/− mice received 600rad TBI on day 0. On day 1, 10⁷ CD25-depleted T cells were adoptively-transferred into irradiated mice. 24 hours later, mice were inoculated subcutaneously with B16.SIY cells as in A. (p = n.s. for comparison of tumor size on day 24 between irradiated CD40^−/− and irradiated C57BL/6 mice following adoptive transfer of CD25-depleted T cells). Results of tumor growth curves in A–C are representative of 2 independent experiments with 4–5 mice/group. D. Upper row - representative flow cytometric analysis of CD4⁺ and CD8⁺ (middle panel) and CD4⁺FoxP3⁺ TIL (right panel) from an established B16.SIY tumor in a RAG2^−/− mouse 10 days following adoptive transfer of CD25-depleted T cells. Lower row – bar graphs depicting the mean percentages of CD4⁺ and CD8⁺ TIL (left) and CD4⁺FoxP3⁻ and CD4⁺FoxP3⁺ TIL (right) among 3 tumors analyzed. E. Established tumors were excised and analyzed by flow cytometry for I-A^b expression after gating on viable, GFP⁺ B16.SIY cells. Shadowed histograms represent isotype control staining, the dotted histogram represents I-A^b expression on B16.SIY cells isolated from irradiated IFN-γ^−/− hosts receiving transfer of IFN-γ^−/− CD25-depleted T cells and the solid histogram represents I-A^b expression on B16.SIY cells isolated from irradiated C57BL/6 hosts receiving transfer of C57BL/6 CD25-depleted T cells.

It was also of interest to also determine whether CD4⁺ T cells were providing help through the alternative mechanism of CD40L-mediated activation of host APCs. To explore this possibility, CD25-depleted T cells were transferred into irradiated (600rad TBI) C57BL/6 or CD40^−/− mice followed by B16.SIY tumor challenge. In fact, tumors were similarly rejected in C57BL/6 or CD40^−/− lymphopenic hosts (Figure 4C). As expected, SIY-specific immune responses were similar in the 2 groups, based on SIY/K^b tetramer staining and IFN-γ ELISPOT analysis (data not shown). These data argue against a significant role of the CD40/CD40L pathway in controlling B16.SIY rejection.

Taken together with the observation that CD4⁺ T cell recognition of antigens derived from B16.SIY tumor cells occurred, and that partial tumor control was achieved with transfer of conventional CD4⁺ T cells alone, these data argue that the major role of CD4⁺ conventional T cells in this model is to contribute to direct tumor control in vivo. Consistent with this notion, CD4⁺ T cells were regularly identified within the tumor microenvironment in vivo (Figure 4D). Additionally, only approximately 1% of the tumor infiltrating CD4⁺ T cells were regulatory T cells (Figure 4D), in stark contrast to what we have observed in the B16 tumor microenvironment of unmanipulated, wildtype mice in which between 20–30% of CD4⁺ T cells express FoxP3 (4). Further, we repeatedly observed significant upregulation of MHC class II (I-A^b) molecules on B16.SIY tumor cells when examined directly ex-vivo (Figure 4E), which occurred in a TBI- and IFN-γ dependent manner (Supplementary Figure 2C). Together, these observations provide additional evidence in favor of direct ant-tumor control by conventional CD4⁺ T cells in this model.

Tumor rejection by CD25-depleted T cells plus homeostatic proliferation requires either the cytotoxic granule-pathway or FasL, but not TNF-α

We next turned to investigate the effector mechanisms involved in rejection of B16.SIY tumors by CD25-depleted T cells in the setting of homeostatic proliferation. Two major cytotoxic mechanisms of CTL have been described - that mediated by release of granules containing perforin and granzymes (27), and also engagement of Fas on target cells through surface expression of FasL (28, 29). In Addition, TNF-α produced by T cells is necessary for tumor rejection in some models (18). To address the role of these pathways in rejection of B16.SIY in vivo, T cells from various gene knockout mice were employed.

To investigate whether TNF-α production was necessary for B16.SIY rejection, CD25-depleted T cells were isolated from spleens of C57BL/6 or TNF-α^−/− donor mice and transferred into RAG2^−/− mice. As with wildtype T cells, B16.SIY tumors were completely rejected by TNF-α^−/− CD25-depleted T cells, indicating that TNF-α production by transferred T cells is not necessary for tumor rejection (Figure 5A). Surprisingly, B16.SIY tumors were rejected similarly following transfer of either perforin^−/− or gld/gld CD25-depleted T cells into RAG2^−/− mice, suggesting that expression of neither perforin nor of FasL was required for tumor rejection (Figure 5B and C). However, it was conceivable that either pathway might be sufficient, and that functional compensation could occur. Therefore, perforin^−/−×gld/gld double mutant mice were generated. Interestingly, CD25-depleted T cells isolated from perforin^−/−×gld/gld mice and transferred into RAG2^−/− mice failed to reject B16.SIY tumors (Figure 5D), although partial tumor control was seen. These results argue that adoptively transferred CD25-depleted T cell-mediated killing of tumor cells involves both the perforin and FasL pathways, but that either is sufficient to drive tumor rejection.

Figure 5. Adoptively-transferred CD25-depleted T cells use either the granule pathway or FasL to eliminate B16.SIY cells.

A. 10⁷ CD25-depleted T cells purified from C57BL/6 or TNF-α^−/− mice were adoptviely-transferred into groups of 5 RAG2^−/− recipient mice. 24 hours later, mice were challenged with 10⁶ B16.SIY cells, and tumor growth was monitored. B. 10⁷ CD25-depleted T cells purified from C57BL/6 or perforin^−/− mice were adoptively-transferred into groups of 5 RAG2^−/− recipient mice. 24 hours later, mice were challenged with 10⁶ B16.SIY cells, and tumor growth was monitored. C. 10⁷ CD25-T cells were purified from C57BL/6 or gld/gld mice and were adoptively-transferred into groups of 5 RAG2^−/− recipient mice. 24 hours later, mice were challenged with 10⁶ B16.SIY cells, and tumor growth was monitored. p = n.s. for comparison of tumor size on the day of last tumor measurement between RAG2^−/− mice following adoptive transfer of wildtype versus TNF-α^−/−, perforin^−/− and gld/gld CD25-depleted T cells in A–C) D. 10⁷ CD25-T cells were purified from C57BL/6 or perforin^−/−×gld/gld mice and were adoptively-transferred into groups of 5 RAG2^−/− recipient mice. 24 hours later, mice were challenged with 10⁶ B16.SIY cells, and tumor growth was monitored. *p = < 0.01 for comparison of tumor size on day 27 between RAG2^−/− mice following adoptive transfer of wildtype versus perforin^−/− × gld/gld CD25-depleted T cells.

IFN-γ production by transferred T cells but not by host cells is required for tumor rejection

It was also of interest to clarify whether IFN-γ played a role in promoting B16.SIY rejection in the current model. To determine whether IFN-γ produced by adoptively-transferred T cells was required, CD25-depleted T cells from C57BL/6 or IFN-γ^−/− donor mice were transferred into RAG2^−/− mice challenged with B16.SIY cells. As shown in Figure 6A, tumor rejection was significantly impaired with IFN-γ^−/− T cells, arguing for a critical importance of this cytokine in regulating B16.SIY rejection. The failure of IFN-γ^−/− CD25-depleted T cells to eliminate B16.SIY tumors was not due to decreased numbers of SIY antigen-specific CD8⁺ IFN-γ^−/− T cells in lymphopenic recipients, as significantly higher numbers of SIY-specific CD8⁺ IFN-γ^−/− versus wildtype T cells were routinely identified in spleens of RAG2^−/− hosts (Supplementary Figure 3A and B).

Figure 6. IFN-γ production by donor T cells, but not host cells, is necessary for B16.SIY rejection.

A. CD25-depleted T cells were purified from spleens of C57BL/6 or IFN-γ^−/− mice, and adoptively-transferred into groups of 5 RAG2^−/− recipient mice. 24 hours later, mice were challenged with 10⁶ B16.SIY cells, and tumor growth was monitored. *p < 0.01 for comparison of tumor size on day 20 between RAG2^−/− mice following adoptive transfer of wildtype versus IFN-γ^−/− CD25-depleted T cells. B. CD8⁺ and CD4⁺ cells were isolated separately from spleens of C57BL/6 and IFN-γ^−/− mice. Purified CD4⁺ cells were further depleted of CD25⁺ cells, and permutations of C57BL/6 or IFN-γ^−/− CD8⁺ and CD4⁺25⁻ cells (3.5 × 10⁶ CD8⁺ T cells, and 6.5 × 10⁶ CD4⁺25⁻ cells) were adoptively-transferred into groups of 5 RAG2^−/− recipient mice. Mice were subsequently challenged with B16.SIY cells as in A. *p < 0.01 for comparison of tumor size on day 33 between RAG2^−/− mice following adoptive transfer of wildtype CD25-depleted T cells, wildtype CD8⁺ plus IFN-γ^−/− CD4⁺25⁻ T cells or IFN-γ^−/− CD8⁺ plus wildtype CD4⁺25⁻ T cells versus IFN-γ^−/− CD8⁺ plus IFN-γ^−/− CD4⁺25⁻ T cells. C. Groups of 5 C57BL/6 or IFN-γ^−/− mice received 600rad TBI, followed by adoptive transfer of 10⁷ CD25-depleted T cells from C57BL/6 or IFN-γ^−/− donor mice. Mice were subsequently challenged with B16.SIY cells as in A. *p = n.s. for comparison of tumor size on day 24 between irradiated IFN-γ^−/− and irradiated C57BL/6 mice following adoptive transfer of wildtype CD25-depleted T cells. ^#p < 0.01 for comparison of tumor size on day 24 between irradiated IFN-γ^−/− and C57BL/6 mice following adoptive transfer of IFN-γ^−/− CD25-depleted T cells versus irradiated IFN-γ^−/− and C57BL/6 mice following adoptive transfer of wildtype CD25-depleted T cells. The tumor growth curves in A–C are representative of 2 independent experiments each with 5 mice/group.

Whether IFN-γ production was required by transferred CD8⁺ T cells, conventional CD4⁺ T cells, or both was investigated by transferring permutations of CD8⁺ (3.5 × 10⁶) and CD4⁺25⁻ T cells (6.5 × 10⁶) from C57BL/6 or IFN-γ^−/− donor mice into RAG2^−/− mice (such that each mouse received a total of 10⁷ T cells – 3.5×10⁶ CD8⁺ and 6.5×10⁶ CD4⁺25⁻ T cells). As shown in Figure 6B, IFN-γ production by either CD8⁺ or CD4⁺25⁻ T cells was sufficient to support B16.SIY rejection, while tumors grew progressively following transfer of combined IFN-γ^−/− CD8⁺ and IFN-γ^−/− CD4⁺25⁻ T cells. Therefore, either CD4⁺ or CD8⁺ T cells must produce IFN-γ for tumor rejection to occur. These results provide additional support for the participation of CD4⁺ T cells at the effector phase of the anti-tumor T cell response.

In order to determine whether IFN-γ produced by host cells was also important, C57BL/6 or IFN-γ^−/− mice received 600rad TBI, followed by adoptive transfer of CD25-depleted T cells from C57BL/6 or IFN-γ^−/− donor mice. As before, IFN-γ^−/− CD25-depleted T cells failed to reject B16.SIY tumors in either irradiated C57BL/6 or IFN-γ^−/− mice. However, complete tumor rejection was observed in irradiated IFN-γ^−/− mice receiving C57BL/6 CD25-depleted T cells (Figure 6C). Therefore, production of IFN-γ by host cells (for example by NK cells) was not required for B16.SIY rejection following adoptive transfer of CD25-depleted T cells into lymphopenic hosts.

IFN-γ can act either upon host cells or tumor cells to mediate tumor rejection

IFN-γ has pleiotropic effects on target cells, and may contribute to anti-tumor immunity at multiple levels, mediating potential effects on T cells, tumor cells, and host stromal cells. In order to define the cellular targets for IFN-γ mediated tumor rejection, IFN-γR^−/− mice were utilized. First, to investigate whether host cells or transferred CD25-depleted T cells themselves were responding to IFN-γ to attain tumor rejection, irradiated C57BL/6 or IFN-γR^−/− mice were lymphodepleted prior to transfer of CD25-depleted T cells from either C57BL/6 or IFN-γR^−/− mice and B16.SIY cell challenge (Figure 7A). Surprisingly, all cohorts showed effective tumor control, including irradiated IFN-γR^−/− hosts receiving transfer of IFN-γR^−/− T cells, suggesting that responsiveness to IFN-γ was not absolutely required on transferred T cells or on any host cell.

Figure 7. Rejection of B16.SIY is lost only when IFN-γ signaling is deficient on both host and tumor cells.

A. Groups of 5 C57BL/6 or IFN-γR^−/− mice received 600rad TBI, followed by adoptive transfer of 10⁷ CD25-depleted T cells isolated from C57BL/6 or IFN-γR^−/− mice. 24 hours later, mice were challenged with 10⁶ B16.SIY cells, and tumor growth was monitored (p = n.s. for comparison of tumor size on day 24 between irradiated C57BL/6 or IFN-γR^−/− mice following adoptive transfer of either wildtype of IFN-γR^−/− CD25-depleted T cells). B. Groups of 5 RAG2^−/− mice received adoptive transfer of 10⁷ CD25-depleted T cells from C57BL/6 mice, followed 24 hours later with a subcutaneous challenge with either 10⁶ B16.SIY.EV or B16.SIY.ΔIFN-γR cells, and tumor growth was monitored (p = n.s. for comparison of tumor size on day 45 between RAG2^−/− mice following adoptive transfer of wildtype CD25-depleted T cells and challenge with B16.SIY.EV versus B16.SIY.ΔIFN-γR cells). (Right) B16.SIY.EV and B16.SIY.ΔIFN-γR cells were cultured in the presence or absence of IFN-γ (20 ng/mL) for 48 hours and analyzed for cell surface K^b expression. Shaded histograms represent K^b expression on B16.SIY.EV and B16.SIY.ΔIFN-γR cells cultured in the absence of IFN-γ. Dotted and solid line histograms represent K^b expression on IFN-γ-treated B16.SIY.ΔIFN-γR and B16.SIY.EV cells, respectively. C and D. Groups of 5 C57BL/6 or IFN-γR^−/− mice received 600rad TBI, followed by adoptive transfer of 10⁷ CD25-depleted T cells from C57BL/6 mice. Cohorts of irradiated C57BL/6 and IFN-γR^−/− mice that received no T cell transfer served as controls. 24 hours later, mice were challenged with either 10⁶ B16.SIY.EV or B16.SIY.ΔIFN-γR cells, and tumor growth was monitored. For clarity, tumor growth curves from the control groups are presented in C., and tumor growth curves from experimental groups (those receiving CD25-depleted T cell transfer) are presented in D. *p = 0.18 for comparison of tumor size on day 26 between irradiated C57BL/6 mice challenged with B16.SIY.EV and irradiated IFN-γR^−/− mice challenged with B16.SIY.ΔIFN-γR. Tumor growth curves in A–D are representative of 2 independent experiments, each with 4–5 mice/group.

It was therefore conceivable that IFN-γ produced by adoptively-transferred CD25-depleted T cells might act directly on the implanted tumor cells through upregulation of class I MHC molecules, increased expression of antigen processing machinery or induced production of anti-angiogenic factors, such as the chemokines CXCL9 or CXCL10 (14, 15). We confirmed that IFN-γ induced a substantial increase of K^b expression on B16.SIY melanoma cells, and also induced robust section of CXCL10 in vitro (Supplementary Figures 4A and B).

To determine whether IFN-γ signaling at the level of the tumor cells was required for rejection in this model, B16.SIY cells were transduced with either an empty vector retroviral construct (B16.SIY.EV), or one containing the cDNA for a truncated, non-signaling form of the IFN-γR (dominant-negative IFN-γR; B16.SIY.ΔIFN-γR). B16.SIY.ΔIFN-γR cells failed to upregulate K^b expression following in vitro exposure to IFN-γ (Figure 7B – right panel). CD25-depleted T cells from C57BL/6 donor mice were adoptively transferred into RAG2^−/− mice, followed by challenge with either B16.SIY.EV or B16.SIY.ΔIFN-γR cells. Surprisingly, potent tumor control was still observed against B16.SIY.ΔIFN-γR tumors, although a partial detriment was observed in some experiments (Figure 7B – left panel). We therefore reasoned that IFN-γ signaling might be required on both host and tumor cells, and only when both were insensitive to IFN-γ would tumor rejection fail. To test this notion, C57BL/6 or IFN-γR^−/− mice received 600rad TBI, followed by adoptive transfer of CD25-depleted T cells from C57BL/6 mice. Subsequently, mice were challenged with B16.SIY.EV or B16.SIY.ΔIFN-γR cells. In fact, we observed that tumor rejection often failed when B16.SIY.ΔIFN-γR tumor cells were implanted into IFN-γR^−/− mice (50–75% of B16.SIY.ΔIFN-γR tumors were not rejected following implantation into irradiated IFN-γR^−/− hosts; p = 0.18 for comparison of tumor size at day 28 between irradiated C57BL/6 mice challenged with B16.SIY.EV versus irradiated IFN-γR^−/− mice challenged with B16.SIY.ΔIFN-γR) (Figures 7C and D). Therefore, either host cell or B16.SIY cell responsiveness to IFN-γ was sufficient for tumor rejection to occur in response to CD25-depleted T cells and homeostatic proliferation, suggesting that both host and tumor cells are the targets of IFN-γ produced by transferred CD25-depleted T cells in this system.

Given that tumor rejection required intact IFN-γ signaling in host tissues and that IFN-γ stimulated tumor cells elaborated the chemokine CXCL10 which has known anti-anti-angiogenic properties, we hypothesized that IFN-γ-dependent tumor rejection might involve anti-angiogenic mechanisms. Therefore, the tumor vasculature was analyzed in the presence or absence of IFN-γ. For these studies, the experimental design was altered to analyze pre-established tumors, in order to facilitate analysis of vascular density in tumors in a more reproducible manner.

To this end, C57BL/6 or IFN-γ^−/− mice were challenged with B16.SIY cells. Six days later, mice received 600rad TBI. On day 8, irradiated C57BL/6 mice received transfer of CD25-depleted T cells from C57BL/6 mice, and irradiated IFN-γ^−/− mice received CD25-depleted T cells from IFN-γ^−/− mice. This experimental design was chosen to facilitate an analysis of tumor angiogenesis in a completely IFN-γ-deficient host so as not to require the use of IFN-γR^−/− donor T cells, IFN-γR^−/− irradiated hosts, or B16.SIY.DIFN-γR tumor cells, which would render such an analysis impractical.

As expected, established B16.SIY tumors were controlled in irradiated C57BL/6 following transfer of wildtype CD25-depleted T cells, but grew progressively in irradiated IFN-γ^−/− mice following IFN-γ^−/− CD25-depleted T cell transfer (Figure 8A). Consistent with our in vitro results, CXCL10 levels were significantly higher in B16.SIY tumors (Figure 8B), and K^b expression was upregulated on B16.SIY cells isolated from irradiated C57BL/6 hosts following wildtype CD25-depleted T cell transfer (Supplementary Figure 4C). When these tumors were analyzed for microvessel density at day 22 following tumor challenge, a significantly higher number of blood vessels both by CD34 expression and by lectin staining was observed in B16.SIY tumors isolated from irradiated IFN-γ^−/− mice versus wildtype mice (Figure 8C and D). These observations support the notion that a major effect of IFN-γ produced by T cells in this model may be to inhibit angiogenesis within the B16 tumor microenvironment.

Figure 8. IFN-γ production in the B16 tumor microenvironment is associated with inhibited angiogenesis.

A. B16.SIY cells (10⁶) were inoculated into cohorts of 5 C57BL/6 or IFN-γ^−/− hosts. Mice were irradiated with 600rad TBI on day 6. On day 8, irradiated, tumor-bearing C57BL/6 mice received either no T cells, or CD25-depleted T cells from C57BL/6 donor mice. Similarly, irradiated, tumor-bearing IFN-γ^−/− mice received either no T cells, or CD25-depleted T cells from IFN-γ^−/− donor mice, and tumor size is reported as observed on day 22 (14 days following T cell transfer) (*p = 0.01 for comparison between tumor size in irradiated C57BL/6 mice receiving C57BL/6 CD25-depleted T cells versus irradiated IFN-γ^−/− mice receiving IFN-γ^−/− CD25-depleted T cells). B. CXCL-10 expression from tumors in A. by ELISA controlled for tumor size (*p = 0.02 for comparison of CXCL-10 expression in B16.SIY tumors from irradiated C57BL/6 mice receiving transfer of 10⁷ CD25-depleted T cells from C57BL/6 donors, compared to those in irradiated IFN-γ^−/− mice transferred with 10⁷ CD25-depleted T cells from IFN-γ^−/− donor mice. C. Approximately 22 days after tumor implantation, the mice in A. were injected with biotinylated lectin (100ug) i.v., and sacrificed 5–10 minutes later. Tumors were excised, and analyzed by immunohistochemistry for lectin and CD34⁺ vessels. Representative images from a single tumor in each group are depicted (Top row – B16.SIY tumor from an irradiated C57BL/6 mouse receiving transfer of wildtype CD25-depleted T cells; bottom row – B16.SIY tumor from irradiated IFN-γ^−/− mouse receiving transfer of IFN-γ^−/− CD25-depleted T cells). D. Bar graph representing the average number of functional (lectin positive) vessels/h.p.f (+/− SE) from mice in A. *p = < 0.01 for comparison between numbers of vessels/h.p.f between the 2 groups. Data are representative of 2 independent experiments, each with 4–5 mice/group.

DISCUSSION

A growing body of evidence has suggested that specific immune inhibitory pathways, acting largely at the level of the tumor microenvironment, can blunt the effector function of anti-tumor T cells and thus allow tumor outgrowth. These include metabolic inhibition by indoelamine-2,3-dioxygenase or arginase (30, 31); expression of ligands for inhibitory receptors, such as PD-L1 (32); extrinsic suppression by CD4⁺CD25⁺FoxP3⁺ Tregs (33); or T cell anergy, resulting from TCR ligation in the absence of adequate costimulation (34). Blockade or elimination of such immune-inhibitory pathways has become an attractive strategy for cancer immunotherapy, and numerous approaches have entered clinical trial testing. Understanding the mechanisms by which tumor control occurs when such pathways are uncoupled is therefore vital for elucidating reasons for success versus failure as data unfold during clinical investigation.

The overall goal of this work was to understand the cellular and molecular mechanisms underlying B16 tumor rejection upon elimination of two key immune suppressive pathways, Tregs and T cell anergy, in a clinically-relevant model. At the priming phase, we determined that cross-presentation of antigen and B7 costimulation by host APCs was required. This is despite the ability of homeostatic proliferation to be capable of inducing a “pseudo-memory” T cell phenotype (5), and argues that antigen-specific priming is till required to expand the relevant subset of tumor-reactive T cells from among the adoptively-transferred polyclonal T cell population. Interestingly, we found no evidence that CD4⁺ T cells were involved in providing “help” for the generation of tumor antigen-specific CD8⁺ effector cells in this system. Rather, accumulated evidence supported the participation of CD4⁺ T cells in the effector phase of the anti-tumor T cell response, which is supported by several other recent publications (10–13). It is interesting to consider that depletion of CD25⁺ cells from the polyclonal T cells prior to adoptive transfer may have revealed the activity of a CD4⁺ effector population. It might be of interest in future studies to investigate a role of CD4⁺ T cells in supporting CD8⁺ T cell memory in this model, as has been indicated in models of anti-viral immunity (25).

It is noteworthy that the final event of tumor control by effector T cells in our model could occur similarly with the individual absence of TNF-α, perforin, or FasL. This result is similar to what has been observed in rejection of allogeneic solid organs or in T cell-dependent models of autoimmunity (35, 36), but contrary to published results in which TNF-α and/or perforin have been shown to be absolutely required for T cell-mediated tumor rejection (18). It appears that redundant cytolytic mechanisms are in place to minimize opportunity for immune escape from this process. However, combined elimination of both perforin and FasL in transferred T cells allowed for tumor outgrowth, suggesting that either cytotoxic pathway can be sufficient for tumor control. While other in other model systems a mandatory role for the perforin pathway has been reported (18, 37–39), differences likely have to do with the specific tumor models and T cell populations involved.

IFN-γ production by anti-tumor T cells has been reported to be vital for tumor elimination in many experimental models (14–18). In the current work, our results have also identified IFN-γ as a central mediator of tumor rejection. Through dissection of the contribution of specific cell populations, we have observed that adoptively-transferred T cells and not cells derived from the recipient mice must produce IFN-γ for tumor rejection to occur. Conventional CD4⁺ and CD8⁺ T cells produced IFN-γ following adoptive transfer in response to tumor antigens, and IFN-γ generated by either CD4⁺ or CD8⁺ T cells was sufficient to support tumor rejection. Further, IFN-γR^−/− donor T cells, IFN-γR^−/− recipient mice and dominant negative IFN-γR B16.SIY cells were employed to facilitate a careful dissection into the cellular compartment(s) in which IFN-γ responsiveness was necessary for tumor rejection to ensue. Interestingly, the action of IFN-γ could be either on host cells or on B16.SIY tumor cells, and tumor rejection was lost only when both were insensitive to IFN-γ. While it was not surprising that IFN-γ was absolutely required for tumor elimination in our model, our results clearly define the cell types necessary for both IFN-γ production and responsiveness in order for tumor rejection to ensue.

While IFN-γ has pleiotropic effects and could exert a multitude of influences, the apparent redundant role on either host cells or tumor cells suggested a possible effect on angiogenesis, which is one mechanism that could be shared by these two cellular compartments. Previous studies have suggested that IFN-γ induces expression of CXCL10, which has previously been associated with an anti-angiogenic effect (15). Indeed, we observed CXCL10 production in the tumor microenvironment upon T cell infiltration in our model, and microvessel density was found to be reduced. Alternatively, IFN-γ might direct impact host stromal cells leading to impaired blood vessel development. Our experiments have demonstrated high levels of CXCL10 production by B16 cells in vitro following exposure to IFN-γ. CXCL10 production within B16 tumors in vivo was also observed, but only when T cells capable of IFN-γ production were transferred. It is likely that both host tumor stromal cells and B16 cells themselves were secreting CXCL10 in response to T cell-produced IFN-γ in vivo, even though insensitivity to IFN-γ in either compartment alone did not routinely lead to failure of tumor rejection, possibly as a result of compensation.

However, additional roles for IFN-γ are also conceivable, as class I and II MHC upregulation on B16 cells in vivo was also observed, which presumably led to their increased recognition and elimination by antigen-specific T cells. It is important to keep in mind that IFN-γ also induces upregulation of immune inhibitory molecules, including IDO and PD-L1 (32, 40). Therefore, an optimal amount and timing of IFN-γ production are likely required to ensure maximal anti-tumor effects.

In summary, our results suggest a model in which tumor antigen cross-presentation by host cells to adoptively transferred CD25-depleted polyclonal T cells promotes T cells rejection, which is dependent upon CD4⁺ cells, IFN-γ, and either the perforin or FasL pathways of cytolysis. The elimination of Tregs combined with homeostatic proliferation maintains persistent T cell function in the context of a growing tumor. Combination with inhibition with other immune evasion pathways may also be of interest to investigate. Strategies to reproduce this approach for clinical application in cancer patients are available and will be of great interest to pursue in future studies. Ideally, such clinical/translational investigations should include interrogation of both the priming and effector phases of the anti-tumor T cell response.