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Conception and design: Y. Guo, C.A. Ahonen, E. Dmitrovsky, M.J. Turk, R.J. Noelle
Development of methodology: Y. Guo, K. Pino-Lagos, C.A. Ahonen, J.L. Napoli, R. Blomhoff, R.A. Chandraratna, E. Dmitrovsky, R.J. Noelle
Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): Y. Guo, J. Wang, J.L. Napoli, R. Blomhoff, S. Sockanathan, R.J. Noelle
Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): Y. Guo, J.L. Napoli, R. Blomhoff, E. Dmitrovsky, R.J. Noelle
Writing, review, and/or revision of the manuscript: Y. Guo, K. Pino-Lagos, E. Dmitrovsky, M.J. Turk, R.J. Noelle
Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): Y. Guo, K.A. Bennett
Study supervision: R.J. Noelle
While vitamin A has been implicated in host resistance to infectious disease, little is known about the role of vitamin A and its active metabolite, retinoic acid (RA) in host defenses against cancer. Here, we show that local RA production within the tumor microenvironment (TME) is increased up to 5-fold as compared with naïve surrounding tissue, with a commensurate increase in RA signaling to regionally infiltrating tumor-reactive T cells. Conditional disruption of RA signaling in CD8+ T cells using a dominant negative retinoic acid receptor α (dnRARα) established that RA signaling is required for tumor-specific CD8+ T-cell expansion/accumulation and protective antitumor immunity. In vivo analysis of antigen-specific CD8+ T-cell responses revealed that early T-cell expansion was RA-independent; however, late T-cell expansion and clonal accumulation was suppressed strongly in the absence of RA signaling. Our findings indicate that RA function is essential for the survival of tumor-reactive CD8+ T cells within the TME.
The morphogenic role of the active form of vitamin A, retinoic acid (RA), in controlling spatial and temporal developmental patterning has underscored the powerful and essential function of this mediator during embryogenesis (1, 2). Similarly, within the immune system, RA has been shown to exert profound effects as a differentiation factor in inducing gut homing of leukocytes (3-6), the differentiation and stability of adaptive regulatory T cells (7-9), the differentiation of CD4+ T cells toward T-helper 1(TH1)/TH17 cells (10, 11), IgA class switching in B cells (6), and the differentiation of myeloid cells (12, 13). It has been proposed that regionalized production of RA is critical for its role as an immune differentiation factor (11). Genetic approaches of ablating RA signaling in T cells have established how RA influences CD4+ T-cell response in vivo (10, 11), but little is known about its role in governing CD8+ T-cell responsiveness. Given the well-recognized role of RA in supporting T-cell responses, and the need for robust T-cell responses in the development of protective antitumor immunity (14), the role of RA in the host resistance against cancer was addressed with a focus on CD8+ T-cell expansion and function. The studies presented herein show that RA is abundantly produced within the tumor microenvironment (TME) and accumulates to levels much higher than in surrounding tissue. Selective disruption of RA signaling in CD8+ T cells incapacitates their ability to undergo effective clonal expansion in vivo and as such, interferes with the development of protective antitumor immunity.
C57BL/6 (CD45.1) and C57BL/6 (CD45.2) were purchased from National Cancer Institute (Bethesda, MD). CD4Cre and OTI transgenic mice were from Jackson Laboratory. The DR5-Luciferase (4) and dominant negative retinoic acid receptor α (dnRARα) mice (15) are as previously described. All animals were maintained in a pathogen-free facility at Geisel School of Medicine at Dartmouth (Hanover, NH).
B16.Ovalbumin (B16.OVA) melanoma cell line was generated (16) by overexpressing chicken OVA-RFP in B16-F10 (obtained from Mary Jo Turk in 2005). B16.OVA was transduced with plasmid containing DR5-Luciferase to generate B16.OVA-DR5-Luciferase cell line. Both cell lines were periodically authenticated by morphologic inspection and tested negative for Mycoplasma contamination by PCR tests in 2008 to 2012, and last time tested in February 2012. For tumor growth curve measurement, 0.5 × 105 B16.OVA cells were injected into mice intradermally and measured 3 times a week. For whole body imaging (WBI), OVA-tetramer staining and IFN-γ enzyme-linked immunosorbent spot (ELISPOT) assay, 1.5 × 105 B16.OVA cells were injected. To deplete CD4+ T cells, mice received 250 μg αCD4 (clone GK1.5, BioXcell). For pan-RAR antagonist treatment, recipient mice were treated intraperitoneally (i.p.) 3 times per week with control vehicle [dimethyl sulfoxide (DMSO)] or 25 μg/mouse Pan-RAR antagonist (NRX 194310, NuRX Pharmaceuticals).
The following FITC-, PE-, PerCP-, APC-Cy7-, Pacific Blue- or APC-conjugated antibodies were used: α-CD45.1 (A20), α-CD45.2 (104), α-CD8 (53–6.7), α-CD11c (N418), α-CD62L (Mel-14), α-CD11b (M1/70), α-CD44 (IM7), α-MHCII (M5), α-IFN-γ (XMG1.2), α-CD69 (H1.2F3), α-BrdUrd (PRB-1), and α-α4β7 (DATK32). All antibodies were purchased from BioLegend except α-BrdUrd and anti-α4β7 (BD Biosciences). MitoTracker Green was purchased from Invitrogen. 7-AAD (BD Biosciences) or LIVE/DEAD Fixable Near-IR Dead Cell Stain Kit (Invitrogen) was used to exclude dead cells in fluorescence-activated cell sorting (FACS) analysis.
WBI and luciferase activity of purified cells (5 × 105 cells per well) was conducted as previously described (11). For in vitro cultured B16.OVA-DR5-Luciferase tumor cells, cells were cultured for 24 hours with RA and Pan-RAR antagonist (2.5 μg/mL), plated at 1 × 106 cells per well, administrated d-luciferin at 150 μg/mL, and imaged. Analysis and images were obtained using the Living Image Software (version 2.6.1).
Tumor, tumor-draining lymph node (TDLN), and spleen were taken from day 6 B16.OVA-bearing or naïve mice. Liquid chromatography/tandem mass spectrometry (LC/MS-MS) was used as described previously to measure all-trans retinoic acid (ATRA) concentration (17).
RALDH activity in individual cells was estimated using ALDEFLUOR staining kits (StemCell Technologies), according to the manufacturer's protocol as previously described (18). For immunophenotyping of ALDHbri cells, the cells were subsequently stained with PE-, PerCP-, APC-, PE-Cy7-, or APC-Cy7–conjugated monoclonal antibody (mAb) in ice-cold ALDEFLUOR assay buffer. Cells were analyzed using FACS Calibur or MACS Quant (Miltenyi Biotech).
C57BL/6 (CD45.1) or DR5-Luciferase (CD45.2) mice were lethally irradiated and received 5 × 106 bone marrow cells harvested from DR5-Luciferase and C57BL/6 (CD45.1), respectively. Reconstitution was confirmed 8 weeks later by staining with α-CD45.1 and α-CD45.2. All BMCs exceeded 96% reconstitution efficiency.
dnRARα and dnRARαCD4Cre mice were immunized with 500 μg Ovalbumin (Sigma–Aldrich), 50 μg αCD40 (BioXcell), and 50 μg polyI:C (InvivoGen) by intraperitoneal injection.
MHCI-I tetramer and IFN-γ staining was conducted as previously described (19). Four-color FACS data were collected on a BD FACSCalibur flow cytometer and analyzed using FlowJo software. Analysis typically pregated on CD8+MHCII− cells.
ELISPOT was done according to the procedures described previously (20). Briefly, CD8+ effector T cells were harvested from spleen, TDLN, or tumor-infiltrating lymphocytes (TIL) of day 12 B16.OVA-bearing mice, and plated at 1:1 ratio with irradiated T-cell–depleted C57BL/6 splenocytes pulsed with 10 μg/mL SIINFEKL peptide (or without peptide as control). Plates were incubated for 20 hours at 37°C and then developed with aminoethylcarbazole chromogen.
Enrichment was done at different time points as described previously (21) and stained as described in the text.
Data graphs were made using GraphPad Prism software and expressed as the mean ± SEM or mean ± SD. Differences for graphs with one grouping variables were analyzed by Student t test (2 groups). For studies of B16.OVA tumor growth and OVA tetramer kinetics, 2-way ANOVA was used to assess significance.In all analysis,*, P<0.05;**, P <0.01; ***, P < 0.001, and ns denotes P > 0.05.
We previously reported that inflammatory mediators induced a spatially and temporally restricted induction of RA synthesis and signaling in vivo (11). To determine if tumor growth in vivo induced restricted and heightened RA signaling and synthesis, a tumor that reports RA signaling was injected in vivo. During the growth of B16.OVA-DR5-Luciferase, in which the RA response element, DR5 repeat, is coupled to a luciferase reporter, an increase of RA signaling over the tumor growth course (Fig. 1A) was observed. The reporting signal was RA-dependent, as it was inhibited by the administration of pan-RAR antagonist (NRX 194310; data not shown). The same RA reporting signal was also observed using B16-DR5-Luciferase cells, indicating that expression of OVA was not critical for induction of RA (data not shown). To confirm that enhanced RA synthesis at the tumor site was induced during tumor growth, DR5-Luciferase transgenic reporter mice were used, which express DR5-luciferase in all tissues (4). B16.OVA tumor cells were injected into DR5-Luciferase mice and RA signaling could be readily detected primarily at the growing tumor site by day 6 (Fig. 1B). The induced RA signaling was also observed in other tumor models, such as EL4 thymoma, MB49 urothelial cacrcinoma, and B16 melanoma (data not shown). To distinguish RA signaling in hematopoietic cells and/or nonhematopoietic compartment in the TME, bone marrow chimeras (BMC) were generated (designated as donor bone marrow → lethally irradiated recipient): DR5-Luciferase→C57BL/6 and C57BL/6 → DR5-Luciferase. Local RA responses in the TME were observed in both recipients as nonreconstituted mice, indicating RA signaling in both hematopoietic and nonhematopoietic cells (Fig. 1C).
To directly measure RA synthesis within the TME, ATRA concentrations from TDLN, spleen, and tumor site were quantified by LC/MS-MS. Data showed that ATRA was significantly increased (~5-fold) in tumor tissue as compared with naïve skin (Fig. 1D). The same trend was also true when ATRA quantity in TDLN was compared with that in naïve lymph node (LN; ~2-fold increase). Conversely, no significant difference was observed in the ATRA levels between spleen from naïve mice and B16.OVA-bearing mice. Taken together, these results showed that RA synthesis and signaling is elicited locally in response to tumor growth in vivo.
A series of studies were designed to determine the relative contribution of the host and tumor cells to elevated levels of RA in the TME. First, real-time (RT-PCR) analysis for the expression of Aldh1a1–3 (encoding RALDH1–3, critical enzymes for RA synthesis) in B16.OVA cells compared with the expression in granulocyte macrophage colony-stimulating factor (GM-CSF) or RA-stimulated dendritic cells (DC; ref. 18) showed (Fig. 2A) that B16.OVA cells expressed no detectable Aldh1a1 or Aldh1a2, and expressed very low Aldh1a3. Second, luciferase activity measured in B16.OVA-DR5-Luciferase cells in vitro was only detectable if exogenous RA (as little as 1nM RA) was added, indicating that the cells produce little if any RA that can drive reporter activity (Fig. 2B). This reporting signal was RA-dependent, as evidenced by its inhibition by Pan-RAR antagonist. Third, RA synthesis in vivo by tumor cells was assessed by ALDEFLOUR (ALDH) staining. As can be seen (Fig. 2C), ex vivo analysis of the CD45−RFP+ cells (tumor) did not report as ALDHbri [0.053% −DEAB (diethylaminobenzaldehyde) vs. 0.047% +DEAB], whereas the CD45+RFP− cells in the tumor site were ALDHbri (0.11 −DEAB vs. 0.013% +DEAB). Taken together, tumor cells contribute little, if at all, directly to elevated tissue levels of ATRA.
Attention was turned to identifying host cells producing elevated levels of ATRA within the TME. RA is constitutively produced in naïve skin (Fig. 2D, top) as reported previously (22). However, on day 5 posttumor inoculation, there was a significant increase in ALDHbri% within the TME as compared with the naïve skin (Fig. 2E, 2.83% ±0.91% vs. 0.36% ± 0.09%, P < 0.05). The frequency of ALDHbri cells increased as the tumor grows, showed by higher ALDHbri% of TILs on day 14 and 22 than day 5 of tumor growth (data not shown). Further analysis showed that the ALDHbri cells comprises various DCs and macrophage subsets, including CD11c+CD11bhi, CD11c+ CD11b− DCs, and CD11c−CD11b+ macrophages (Fig. 2F). Analysis indicated that approximately 50% of ALDHbri cells express high levels of MHCII (Fig. 2F), indicating they may be mature antigen-presenting cells (APC) within the TME. Taken together, the data suggest that the tumor-infiltrating DCs and macrophages contribute to RA-enriched TME.
Further studies were designed to determine the lineage of cells being signaled by RA within the TME and TDLN. To identify the hematopoietic cells reporting RA signaling, specific cell lineages were purified from TIL, TDLN (inguinal LN), and spleen of the DR5-Luciferase mice, and whole-cell luciferase assays were conducted. There were no significant differences between the basal level of luciferase activity in cells from naïve DR5-Luciferase (Luc+) and control mice (Luc−; data not shown). B16.OVA growth induced about approximately 6-fold increase in luciferase activity in CD8+ T cells from the TME in DR5-Luciferase mice (Fig. 3A; TIL Luc+ vs. Luc−). However, when the luciferase activity was measured in the same number of cells, there was only approximately 2.5- and 1.2-fold increase in CD4+ T cells and CD11c+ DCs from TME (TIL Luc+ vs. Luc−), respectively. The same trend was observed in TDLN and spleen, except that there was also a significant increase in RA signaling in CD11c+ DCs from the spleen (tumor vs. naïve). Thus, CD8+ T cells in the TME showed a marked response to elevated concentrations of ATRA in vivo.
To measure tumor-specific CD8+ T-cell response to heightened RA concentration in the TME, naive OVA-specific CD8+ T cells from DR5-Luciferase × OTI (OTILuc) mice were transferred into B16 and B16.OVA-bearing mice, respectively. CD8+ T cells in OTILuc mice recognize OVA antigen and elicit luciferase activity when RA signaling occurs. As shown in Fig. 3B and C, in B16.OVA-bearing mice, OTILuc T cells showed robust RA response signal at the tumor site as compared with nondetectable RA reporting signals in B16-bearing mice. This further confirmed that CD8+ T cells are responsive to RA produced at the tumor site in a tumor-specific manner.
High ATRA tissue levels and RA signaling induced at the tumor site suggests that RA may be important in regulating tumor-specific immunity. To address the functional importance of RA signaling to CD8+ T-cell function, a dominant negative RARα403 (dnRARα; ref. 15) was overexpressed in T cells by interbreeding the dnRARα with CD4Cre mice. Studies showed that dnRARαCD4Cre mice expressed Cre-dependent dnRARα in both CD4+ and CD8+ T cells (Supplementary Fig. S1A). In addition, it was confirmed that CD8+ T cells in dnRARαCD4Cre were unresponsive to RA in vitro, as shown by the inability to upregulate α4β7 in response to RA (Supplementary Fig. S1B) and that the dnRARαCD4cre did not disrupt signaling through peroxisome proliferator-activated receptor γ and vitamin D receptor (23; data not shown).
With specificity of the dnRARα for RA signaling confirmed, the role of RA signaling on tumor-specific CD8+ T-cell function in vivo was evaluated. The endogenous OVA-specific CD8+ T-cell responsein B16.OVA-bearing dnRARαCD4Cre and dnRARα mice was measured. As previously published in B16 melanoma model (24), CD4+ T-cell depletion induces greater endogenous CD8+ T-cell priming against tumor-associated antigen in B16.OVA model. Therefore, to measure more robust CD8+ T-cell response in the absence of CD4+ T cells and irrefutably assign any functional impact of RA signaling disruption to CD8+ T cells, B16.OVA were injected into CD4+ T cell-depleted dnRARα and dnRARαCD4Cre mice. Ablation of RA signaling in the dnRARαCD4Cre mice resulted in a marked decrease of CD44hiOVA-tetramer+ CD8+ T cells as compared with the equivalent cells in the control mice over the tumor growth course (Fig. 4A and B). Furthermore, there was a significant decrease in IFN-γ–producing CD8+ T cells from tumor tissue (TIL 4.3 ± 0.67 vs. 269.0 ± 12.9, P < 0.001), TDLN (1.0 ± 0.57 vs.171 ± 15.1, P < 0.001), and spleen (0.66 ± 0.33 vs. 173.37 ± 21.37, P < 0.01) in dnRARαCD4Cre as compared with dnRARα mice (Fig. 4C), respectively. The fewer functional OVA-specific CD8+ T cells in dnRARαCD4Cre mice suggests that RA signaling is required for tumor-specific CD8+ T-cell accumulation and function in the TME.
Faster B16.OVA growth was observed in dnRARαCD4Cre mice in comparison with dnRARα mice (Fig. 4D), suggesting reduced immune surveillance. The enhanced tumor growth was independent of deficient CD4+ T cell help because the same effect was observed when CD4+ T cells were depleted in both groups over the entire tumor growth course (Fig. 4E). Chemical inhibition of RA signaling recapitulated the phenotype observed in the dnRARαCD4Cre mice (Fig. 4F). In vitro culture experiments established that the pan-RAR antagonist used in this study did not inhibit B16.OVA growth over a wide dose range (data not shown). In conclusion, these studies establish that RA signaling to CD8+ T cells in vivo is critical for optimal suppression of tumor growth.
To gain greater insights into the underlying mechanism of RA controlling CD8+ T-cell clonal expansion, a more robust CD8+ T-cell response induced by CD40 agonist, together with a TLR agonist and soluble antigen system was exploited(19). As observed in the B16.OVA tumor model, dnRARαCD4Cre mice showed a significant decrease in the frequency of OVA-specific CD8+ T cells as compared with control dnRARα mice (14.9% vs. 0.6%; Fig. 5A and B). CD44hiOVA-tetramer+ CD8+ T cells were quantified in the blood, spleen, mesenteric LN (MLN), and peripheral LN (PLN) on peak response time of day 6 to ensure that the reduced frequency in the blood was not due to a trafficking defect but rather was representative of the overall CD8+ T-cell response (Fig. 5C). When restimulated ex vivo by SIINFEKL peptide or α-CD3, the frequency of IFN-γ–producing CD8+ T cells in dnRARαCD4Cre was significantly lower than in dnRARα mice (Fig. 5D and E). Identical results were obtained in mice that were depleted of CD4+ T cells (Supplementary Fig. S2), establishing that intrinsic RA signaling deficiency in CD8+, but not CD4+ T cells, accounted for the defective CD8+ T-cell accumulation.
Tetramer enrichment allowed for the quantification of total OVA-tetramer+ CD8+ T cells at very early time-points following immunization (21). On day 1 and 4 postimmunization, CD69 expression as well as increases in forward scatter, respectively, of OVA-tetramer+ CD8+ T cells from dnRARαCD4Cre and dnRARα mice were indistinguishable (Fig.6A and B), indicating that the expression of dnRARα imparts no effect on early T-cell activation. During the initial expansion between day 0 and 4, OVA-tetramer+ CD8+ T cells in dnRARα CD4Cre mice accumu-lated to the same extent as their counterparts in dnRARα mice, showed by the same approximately 4-fold increase over naïve control (Fig. 6C). However, OVA-specific CD8+ T cells failed to accumulate further between day 4 and 6 in dnRARαCD4Cre mice, whereas the counterparts in dnRARα mice expanded logarithmically (~50-fold).
Analysis of proliferation of OVA-specific CD8+ T cells by bromodeoxyuridine (BrdUrd) incorporation between day 4 and 6 revealed a slight but not significant decrease in OVA-tetramer+ cells incorporating BrdUrd between dnRARα and dnRARαCD4Cre (both between 40% and 50%; Fig.6D and E). To determine if there was a survival deficiency of the proliferating CD8+ T cells in dnRARαCD4Cre mice, enriched OVA-specific CD8+ T cells were stained with MitoTracker on day 5 postimmunization. Interestingly, while approximately 20% of the cells were MitoTracker-negative (undergoing apoptosis) in control dnRARα mice, approximately 50% of OVA-tetramer+ CD8+ T cells were MitoTracker-negative in dnRARαCD4Cre mice (Fig. 6D). Therefore, the abortive accumulation of OVA-specific CD8+ T cells in dnRARαCD4Cre mice between day 4 and 6 postimmunization may be due to an equal number of cells undergoing proliferation and apoptosis simultaneously (Fig. 6E). In conclusion, although early CD8+ T-cell expansion is independent of RA signaling in vivo, late CD8+ T-cell clonal accumulation requires RA signaling to maintain better survival during proliferation and effector development.
Accumulating evidence suggests that heightened tissue levels of RA develop at sites of inflammation (11). In the present study, we reported for the first time that the tumor induces a temporally and spatially restricted production and heightened accumulation of RA within the TME. Host tissue DCs and macrophages instead of tumor cells, were major contributors to high-level ATRA within the TME. The marked CD8+ T-cell responses to regionally produced RA suggested that the host may exploit RA to facilitate the development of CD8+ T-cell–mediated protective antitumor immunity. The functional importance of RA signaling in CD8+ T cells was confirmed by studies in mice in which the RA signaling in T cells was genetically impaired. RA signaling-deficient CD8+ T cells failed to expand/accumulate and produce IFN-γ, thus leading to enhanced B16.OVA tumor growth. These studies support the notion that the expansion, accumulation, and differentiation of tumor-specific CD8+ T cells are dependent on intrinsic RA signaling. An in-depth in vivo analysis of OVA-specific CD8+ T cells revealed RA signaling disruption did not impact on upregulation of early activation markers, cell enlargement, or early in vivo expansion induced by OVA immunization. The RA signaling-deficient CD8+ T cells proceed through early rounds of division but fail to ultimately accumulate due to dominant apoptosis in the late clonal expansion. As such, RA signaling seems to render these proliferating cells resistant to apoptosis at the effector phase. Because previous studies have suggested that RA signaling may influence IL-2 expression in activated T cells (25, 26), dnRARαCD4cre mice were treated with IL2/aIL2 complex to restore IL-2 levels. However, this did not overcome the deficiency in OVA-specific CD8+ T-cell expansion in immunization model (data not shown). Additional investigative studies on the molecular basis for how intrinsic RA signaling controls CD8+ T-cell survival at the effector phase are underway.
The authors thank Dr. Pierre Chambon for insightful discussions on data and manuscript preparation, Drs. Edward Usherwood, Victor C. de Vries, and Jose R. Conejo-Garcia for critically reading the manuscript, and Dr. Joshua J. Obar for technical help.
This work was supported by the NIH (AT005382, AI048667, CA062275, and CA120777), the Medical Research Council Centre for Transplantation, the Biomedical Research Council, The Wellcome Trust, and Samuel Waxman Cancer Research Foundation.
Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).
Disclosure of Potential Conflicts of Interest
R. Blomhoff and has an ownership interest (including patents) in Cgene. E. Dmitrovsky has ownership interest (including patents) in a pending patent. R. A. Chandraratna is employed (other than primary affiliation; e.g., consulting) in Io Pharmaceuticals. He is a consultant/advisory board member, holds title of SAB Chair, and has ownership interest (including patents) in Io Pharmaceuticals. No potential conflicts of interest were disclosed by the other authors.