MHC class I-independent activation of virtual memory CD8 T cells induced by chemotherapeutic agent-treated cancer cells

Keywords: DNA-damaging agents, B-cell lymphomas, virtual memory CD8 T cells, cancer immunotherapy, MHC class I

Subject terms: Tumour immunology, Immune evasion

Mice and in vivo treatment

Mice (6–8 weeks) were injected subcutaneously in both flanks with 1 × 10⁶ A20 lymphoma cells (WT BALB/c), MC38 colon cancer cells (WT C57BL/6 (B6)), or DHL16 lymphoma cells (humanized BRGS). When tumors reached a certain size, tumor-bearing recipient mice were randomized into groups, and treated as indicated. Humanized BRGS mice were provided by the Humanized Mouse Core of the Translational Research Networking and Preclinical Models (TRNPM) facility at the University of Colorado Anschutz Medical Campus (AMC). Briefly, humanized mice were generated via injection of CD34⁺ HSCs isolated from cord blood (CB) into sublethally irradiated newborn BRGS pups (1–3 days old), as described previously.^40–43 CD34-CB-BRGS humanized mice (hu-CB-BRGS) were provided to us, and human chimerism was >25% in the blood at both 10 and 15 weeks post engraftment for humanized mice to be used in subsequent experiments. All mice were maintained under specific pathogen-free conditions in the vivarium facility of the University of Colorado AMC. Animal work was approved by the Institutional Animal Care and Use Committee of the University of Colorado Anschutz Medical Campus (Aurora, Colorado, USA).

Tumor dissociation and flow cytometry

Tumors were harvested from tumor-bearing mice. Tumor weight was measured before dissociation, and tumors were processed into single-cell suspensions. The antibodies used for flow cytometry are listed in Supplementary Table 1. Dead cells were excluded by the Live/Dead Fixable Green Dead Cell Stain Kit (Invitrogen). BD Fix/Permeabilization buffer was used for intracellular staining of IFN-γ and granzyme B (GZMB) in TILs. Before IFNγ staining, equal numbers of tumors were cultured in vitro for 6 h in the presence of 50 ng/mL phorbol 12-myristate 13-acetate (PMA) (Sigma Aldrich), 1 μg/mL ionomycin (Sigma Aldrich), and 5 μg/mL BFA (BioLegend). For Eomes staining, a True-nuclear Transcription Factor Buffer Set was used (BioLegend). Data were acquired on a BD Fortessa or a BD FACSCalibur, and analyzed with FlowJo software V10 (Oregon, USA).

Cell culture

A20 lymphoma cells and MC38 cells were obtained from the cell line vendor ATCC in 2017. The SU-DHL-16 cell line was a gift from Dr. Wing C. (John) Chan (City of Hope Medical Center) in 2016 and cultured, as described previously.³ Cell line authentication and mycoplasma testing were performed by the Molecular Biology Service Center at the Barbara Davis Center (University of Colorado, Anschutz Medical Campus) in 2019. The cells were tested and authenticated with PCR assays as described (http://www.barbaradaviscenter.org/). The number of passages between thawing and use in the described experiments generally ranged from two to five.

Tumor cells were cultured at 0.5 × 10⁶/ml and treated with Ara-C or doxorubicin (DOX) at the indicated concentrations for 16 h. Tumor cells were thoroughly washed with PBS at least three times before coculture with CD8 T cells. Total CD8 T cells were isolated from the spleen and/or lymph nodes of WT naive BALB/c or B6 mice by a negative selection kit (Stem Cell Technologies, Canada), and cultured with either untreated or treated tumor cells in the presence of human IL-2 (33 IU/ml). Four days after coculture, CD8 T cells were analyzed by flow cytometry. For cell sorting, cells were stained with the desired antibodies and sorted with a BD Aria Fusion Sorter. Human CD8 T cells were isolated from human blood or hu-CB-BRGS mice by a positive selection kit (Stem Cell Technologies, Canada), cultured with untreated or treated tumor cells in the presence of human IL-2 (100 IU/ml), and collected 4 days after coculture for flow cytometry analysis.

Chemotherapy-treated tumor cells activate CD8 T cells that acquire VM phenotypes

To test our hypothesis that DNA-damaged cancer cells may directly activate CD8 T cells, we set up a new experimental system by coculturing chemotherapy-treated or untreated tumor cells with syngeneic CD8 T cells. We first tested A20 cells, a B-cell lymphoma line derived from the BALB/c mice,⁴⁴ that were untreated or treated with DOX or Ara-C. Chemotherapeutic drugs were thoroughly washed off, and equal numbers of untreated or treated A20 cells were cultured with total CD8 T cells from WT naive BALB/c mice. DOX- or Ara-C-treated A20 cells promoted CD8 T-cell proliferation (Supplementary Fig. 1a) and activated CD8 T cells by enhancing granzyme B (GZMB) (Fig. 1a) or IFN-γ (Supplementary Fig. 1B) production. Notably, CD8 T-cell activation was only induced by live cancer cells, but not dead cells (Supplementary Fig. 2A); thus, we titrated the concentration of chemotherapeutic drugs used to treat cancer cells (Supplementary Fig. 2B–E). Untreated A20 lymphomas failed to activate CD8 T cells (Fig. 1a; Supplementary Fig. 1A, B). Since lymphomas grow aggressively, it is possible that untreated lymphomas cannot activate CD8 T cells due to competition for growth factors or nutrients in culture media. To test this, we titrated the input number of untreated lymphoma cells while keeping the same number of CD8 T cells. We observed no activation of CD8 T cells regardless of the input number of untreated A20 lymphoma cells (data not shown). We conclude that only DNA-damaged A20 lymphoma cells activate CD8 T cells.

Consistently, DOX-treated B16F10 melanoma cells also activated CD8 T cells by enhancing GZMB production (Fig. 1b). Furthermore, DOX-treated MC38 cells, a colon cancer cell line, activated CD8 T cells in a similar fashion (Fig. 1c), thereby implicating the generality of our findings to other types of cancers. We characterized the phenotypes of activated CD8 T cells and found that CD8⁺GZMB⁺ T cells were also CD44⁺CD122⁺NKG2D⁺; furthermore, the majority of them were CD49d⁻ (Fig. 1d; Supplementary Fig. 3A), consistent with VM CD8 T cell phenotypes. All CD8 T cells activated by Ara-C-treated A20 tumor cells expressed Eomes, and upregulated the coinhibitory receptors CTLA-4, Tim-3, and PD-1 (Fig. 1e). Moreover, these activated CD8 T cells also expressed phosphor-AKT (Supplementary Fig. 3B). Taken together, these findings lead us to conclude that chemotherapeutic drug-treated cancer cells can activate CD8 T cells that acquire VM CD8 T-cell phenotypes.

Chemotherapeutic treatment enhanced total CD8 TILs and preferentially increased VM CD8 TILs in different tumor models

To determine whether CD8 T cells contribute to chemotherapeutic effects, A20 B-cell lymphoma cells were transplanted into syngeneic WT BALB/c mice, which developed secondary tumors approximately 2 weeks after tumor inoculation. When the tumor size reached ∼1000 mm³, we randomized recipient mice into two groups and treated them with vehicle control or Ara-C. A single dose of Ara-C treatment eliminated A20 lymphomas (Fig. 2a); however, depleting CD8 T cells completely abrogated the durable effects of Ara-C treatment since all tumors relapsed (Fig. 2b). We analyzed TILs 5 days after Ara-C treatment and found that the number of CD8 TILs was significantly increased by Ara-C treatment (Fig. 2c). Furthermore, Ara-C treatment induced more production of IFN-γ from CD8 TILs than no treatment in tumor-bearing recipients (Fig. 2d, e). Further characterization of CD8 TILs showed that Ara-C treatment significantly increased the percentage and number of VM CD8 TILs (CD8⁺CD44⁺CD122⁺CD49d⁻) (Fig. 2f–h).

To generalize our findings, we next tested the effects of another chemotherapeutic drug, DOX, on CD8 TILs. MC38 colon cancer cells were transplanted into syngeneic WT B6 mice, and when the tumor size reached ~200 mm³, we treated tumor-bearing mice with vehicle control or DOX. Similar to what we observed in Ara-C-treated tumors, the number of total and VM CD8 TILs was significantly increased in DOX-treated MC38 tumors compared with controls (Fig. 2i–k), and the percentage of VM CD8 T cells was preferentially increased in total CD8 TILs in DOX-treated MC38 tumors (Fig. 2l). To examine the functionality of CD8 T cells, we transplanted MC38 cells into syngeneic CD8^−/− mice. When the tumor size reached ~200 mm³, tumor-bearing mice were randomized into four groups: vehicle control, DOX-treated, CD8 T cell-transferred, or combination treatment (DOX plus CD8 T cells). The combination treatment significantly inhibited tumor growth compared with other treatments (Supplementary Fig. 4A, B). Overall, our data show that CD8 T cells, especially VM CD8 T cells, play a critical role in mediating chemotherapeutic effects.

Mechanisms of CD8 T-cell activation induced by chemotherapy-treated tumors

To dissect the mechanism of CD8 T-cell activation, we physically separated the Ara-C-treated A20 cells and CD8 T cells using transwells and found that GZMB upregulation in CD8 T cells was abrogated compared with that in the untreated negative or Ara-C-treated positive control (Fig. 3a). We conclude that CD8 T-cell activation requires cell–cell contact between Ara-C-treated lymphoma cells and CD8 T cells. Next, we asked whether this mode of CD8 T-cell activation requires MHC class I. We previously employed the CRISPR/Cas9 technique to generate A20 β2M-KO lymphoma cells.³ Using WT and β2M-KO lymphomas, we showed that Ara-C-treated WT and β2M-KO A20 lymphomas activated CD8 T cells to a similar extent (Fig. 3b). Our data demonstrate that Ara-C-treated lymphoma cells can activate CD8 T cells independent of tumor-derived MHC class I. Furthermore, we employed an MHC class I-blocking antibody⁴⁵ in our coculture assay and found that blocking MHC class I had no effects on the activation of CD8 T cells mediated by Ara-C-treated B2M-KO A20 lymphoma cells (Supplementary Fig. 5A), further confirming an MHC class I-independent response.

We previously showed that Ara-C treatment increased cyclin A2 protein expression.⁴⁶ Thus, we used cyclin A2 expression as an indicator of Ara-C treatment efficacy. The translation inhibitor cycloheximide (2 or 20 μg/ml) efficiently inhibited the upregulation of cyclin A2 induced by Ara-C treatment (Supplementary Fig. 5B). However, A20 lymphoma cells cotreated with cycloheximide plus Ara-C still activated CD8 T cells similarly to Ara-C-monotherapy-treated A20 lymphoma cells (Supplementary Fig. 5C). These data suggest that Ara-C-treated lymphomas do not activate CD8 T cells through newly synthesized proteins; instead, the putative activating factors might be pre-existing and translocate to the cell membrane, where they activate CD8 T cells. To identify such putative activating factor(s), we applied various blocking antibodies to the coculture system. However, blocking CD44, PD-1, or LAG-3 had no effects on the activation of CD8 T cells (Supplementary Fig. 5D, F). In addition, CD8 T-cell activation was not affected by an anti-CD8-blocking antibody (Supplementary Fig. 5G), consistent with our data showing that MHC class I is not required for such a mode of CD8 T-cell activation. While Ara-C-treated A20 cells upregulated Rae-I (data not shown), a ligand for NKG2D, blocking NKG2D had no effects on the CD8 T cell activation induced by Ara-C-treated A20 tumors (Supplementary Fig. 5E).

Given that these activated CD8 T cells also expressed phosphor-AKT (Supplementary Fig. 3B), we next tested whether the PI3K pathway is involved in the activation of CD8 T cells induced by chemotherapy-treated tumor cells. Various kinase-specific inhibitors were employed in our coculture assays. Inhibitors of Src, Syk, or PIK3CD (p110δ), a catalytic subunit of PI3K, partially blocked the CD8 T-cell activation induced by Ara-C-treated A20 lymphomas, whereas an inhibitor of PIK3CA (p110α) appeared to completely block CD8 T-cell activation (Fig. 3c). Consistently, the Src, Syk, or PIK3CD inhibitors partially reduced the phosphor-AKT level, while the PIK3CA inhibitor almost abrogated it (Fig. 3d). Moreover, these inhibitors at the indicated concentrations had no effect on tumor cell growth, and pretreating tumor cells with such inhibitors did not affect CD8 T-cell activation, suggesting a direct inhibitory role of these inhibitors on CD8 T cells (data not shown). Overall, these data reveal a critical role of the Src, Syk, and PI3K pathways in the CD8 T-cell activation induced by chemotherapy-treated tumor cells.

To further investigate the mechanisms of CD8 T-cell activation, we employed additional inhibitors and found that an inhibitor of Toll-like receptor 4 (TLR4), TAK242, partially inhibited the activation of CD8 T cells mediated by chemotherapy-treated cancer cells (Fig. 3e). These data imply that the TLR signaling pathway may be involved in CD8 T-cell activation; moreover, these data suggest that additional molecules or signaling pathways may also contribute to such a mode of CD8 T-cell activation.

VM CD8 T cells preferentially responded to chemotherapy-treated tumors, and CD8 T cells activated by chemotherapy-treated tumors exhibited potent cytotoxicity independent of MHC class I

Since the phenotypes of activated CD8 T cells resemble those of VM CD8 T cells, we sorted different subsets of CD8 T cells from WT naive BALB/c mice and confirmed their purity, and then tested which subset preferentially responded to chemotherapy-treated tumors (Supplementary Fig. 6A, B). Upon coculture with Ara-C-treated B2M-KO A20 tumor cells, the CD8⁺CD44⁺C122⁺CD49d⁻ subset (VM CD8 T cells) produced the highest level of GZMB (Supplementary Fig. 6A), demonstrating that VM CD8 T cells preferentially responded to chemotherapy-treated tumor cells independent of tumor-derived MHC class I. In contrast, the CD44⁻CD122⁻CD49d⁻ subset did not produce a high level of GZMB (Supplementary Fig. 6A), suggesting that naive CD8 T cells did not acquire VM CD8 T-cell phenotypes during coculture and that GZMB production in the unsorted total CD8 T cells was likely attributed to the expansion of pre-existing VM CD8 T cells.

To test whether activated CD8 T cells exhibit cytotoxicity to untreated tumor cells, we set up a complex sequential coculture assay (Fig. 4a). We first cocultured total CD8 T cells with either untreated or Ara-C-treated B2M-KO A20 cells for 4 days. Then, we isolated these CD8 T cells and added them into another coculture with untreated B2M-KO A20 cells. Two days later, flow cytometry analysis was performed to examine tumor cell death (Fig. 4a). Untreated B2M-KO A20 tumor cells that were cultured alone (group I) showed little cell death (Fig. 4b, c). CD8 T cells that were not activated by Ara-C-treated B2M-KO A20 cells also led to a minimal level of tumor cell death in groups II and III (Fig. 4b, c). In contrast, due to coculture with CD8 T cells previously activated by Ara-C-treated B2M-KO A20 cells, untreated B2M-KO A20 cells exhibited the highest level of cell death in group IV, demonstrating a potent effector function of these activated CD8 T cells (Fig. 4b, c).

Our results showed that upon coculture with chemotherapy-treated tumor cells, CD8 T cells produced GZMB (Fig. 1a–c), which can mediate the apoptosis of target cells. To further investigate how tumor cells were killed, we first examined GZMB expression in tumor cells and found that tumors did not express any GZMB in the absence or presence of Ara-C treatment (Supplementary Fig. 7). In contrast, we detected GZMB in Ara-C-treated tumor cells when they were cocultured with CD8 T cells (Supplementary Fig. 7). Furthermore, in our sequential coculture assay, a GZMB inhibitor (Z-AAD-CMK) significantly reduced tumor cell death upon coculture with activated CD8 T cells compared with the vehicle control (Fig. 4d, Group IV); however, NKG2D blockade had no effects on tumor cell death (Fig. 4d, Group IV). We also included cleaved caspase 3 as an apoptotic marker in our sequential coculture assay. Upon coculturing with activated CD8 T cells, tumor cells expressed much more cleaved caspase 3 than other control groups (Fig. 4d, bottom panel, Group I–III vs. Group IV). Taken together, our data show that CD8 T cells activated by chemotherapy-treated tumors exhibit strong cytotoxicity against untreated tumor cells independent of tumor-derived MHC class I, but dependent on GZMB.

VM CD8 T cells led to a better therapeutic effect on MC38 B2M-KO tumors

We showed that DOX-treated MC38 tumor cells activated CD8 T cells (Fig. 1c); however, it remains unknown whether MHC class I affects CD8 T-cell activation. We employed the CRISPR/Cas9 approach as described previously³ to generate MHC class I-deficient MC38 cell lines (MC38 B2M^−/−) (Fig. 5a). The deletion of the B2M gene was confirmed by PCR and Sanger sequencing, as shown with an example of the MC38 B2M^−/− colon cancer line (Fig. 5b) that harbored a deletion on each allele of the B2M gene (Supplementary Fig. 8A, B). Western blot results further confirmed the absence of B2M protein in the MC38 B2M^−/− cells (Fig. 5c). CD8 T cells were activated to the same extent by chemotherapy-treated WT or B2M^−/− MC38 tumor cells in vitro (Fig. 5d, e), demonstrating a dispensable role of tumor-derived MHC class I in the CD8 T-cell activation induced by chemotherapy-treated tumor cells.

To compare the therapeutic effects conferred by total vs. VM CD8 T cells, we sorted the total or VM CD8 T cells from syngeneic WT B6 mice and transferred them to CD8^−/− mice-bearing MC38 B2M^−/− tumors that were either untreated or treated with DOX (see schematics in Fig. 5f). Treatment with either T cells or DOX alone had little effect on tumor growth (Fig. 5f, g). In contrast, adoptive transfer of total CD8 T cells plus DOX treatment significantly inhibited the growth of MC38 B2M^−/− tumors (Fig. 5g). Furthermore, VM CD8 T cells showed a better therapeutic effect than total CD8 T cells when combined with DOX treatment (Fig. 5g). In addition, treatment with VM CD8 T cells plus DOX led to a significantly longer survival of recipient mice compared with treatment with total CD8 T cells plus DOX, indicating that VM CD8 T cells may be more potent at inhibiting tumor growth and prolonging recipient survival than total CD8 T cells (Supplementary Fig. 9). Of note, treatment with total CD8 T cells plus DOX also led to a significantly longer survival of recipient mice compared with treatment with the vehicle control (Supplementary Fig. 9), implying that other subsets of CD8 T cells may also contribute to the therapeutic effects.

Chemotherapy-treated human lymphomas activated human VM CD8 T cells

Previous studies reported that human VM CD8 T cells express KIR/NKG2A and Eomes.^47,48 To examine the effects of chemotherapeutic treatment on human VM CD8 T cells, we transplanted SU-DHL-16 (DHL-16) lymphoma cells into BRGS mice that had been humanized with cord blood stem cells (hu-CB-BRGS). Once the tumor size reached ~500 mm³, tumor-bearing mice were treated with Ara-C, and TILs were analyzed with flow cytometry. Ara-C treatment increased the percentage of total CD8 T cells in human CD45⁺ TILs (Fig. 6a, b) and the number of total human CD8 TILs (Fig. 6c). Ara-C treatment preferentially increased the percentage of human VM CD8 TILs in the human CD8 TIL population (Fig. 6d) and drastically enhanced the number of human VM CD8 TILs (Fig. 6e).

We next tested whether chemotherapy-treated DHL-16 lymphoma cells directly increase and activate human CD8 T cells isolated from hu-CB-BRGS mice. To prevent the mismatch effect of MHC class I between DHL-16 lymphoma cells and cocultured human CD8 T cells, we employed the CRISPR/Cas9 approach to delete the B2M gene in the DHL-16 lymphomas, which led to the loss of HLA-A, B, and C expression in DHL-16 cells (Fig. 6f). Compared with the groups of T cell only or untreated DHL-16 B2M^−/− cells, the group of DOX-treated DHL-16 B2M^−/− cells had a significantly increased percentage of VM CD8 T cells (Fig. 6g) and enhanced production of GZMB in CD8 T cells (Fig. 6h). Last, when we employed human CD8 T cells isolated from the peripheral blood of healthy donors, similar findings were observed (Supplementary Fig. 10A, B). Taken together, our data demonstrate that chemotherapy-treated human tumors also activated human CD8 T cells, especially VM CD8 T cells, independent of tumor-derived MHC class I.

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