Radiotherapy and immunotherapy promote tumoral lipid oxidation and ferroptosis via synergistic repression of SLC7A11

Radiotherapy induces tumor cell ferroptosis

Ferroptosis is a recently discovered form of cell death that differs from apoptosis and results from iron-dependent lipid peroxide accumulation (12,17). It is unknown whether lipid oxidation and ferroptosis are induced by radiotherapy. To test this in vivo, we inoculated ID8 ovarian cancer cells into the peritoneal cavity of C57BL/6 mice and treated the mice with a single dose of 8 Gray (Gy) of ionizing radiation, liproxstatin-1, a ferroptosis antagonist, or both agents concurrently. ID8 tumor bearing mice tolerated radiotherapy without changes in mouse weight (Supplementary Fig. S1A). Radiotherapy, but not liproxstatin-1, reduced tumor growth (Fig. 1A). Unexpectedly, liproxstatin-1 administration concurrent and adjuvant to radiotherapy diminished radiotherapy efficacy (Fig. 1A).

To extend this finding, we inoculated B16F10 melanoma cells into C57BL/6 mice and allowed the tumors to establish. Subsequently, B16F10 bearing mice were treated with a single fraction of 8 Gy. Again, radiotherapy reduced tumor growth (Supplementary Fig. S1B). Dual treatment with liproxstatin-1 and radiotherapy partially and significantly abrogated radiotherapy efficacy. Ferroptosis is accompanied by increased lipid peroxidation, which can be examined using the lipophilic redox sensitive dye C11BODIPY (12). C11BODIPY quantification showed that radiotherapy increased lipid peroxidation in vivo, and concurrent liproxstatin-1 treatment abrogated radiotherapy induced lipid peroxidation in B16F10 tumors (Fig. 1B and Supplementary Fig. S1C).

To orthogonally confirm tumoral ferroptosis occurs following radiotherapy, we generated ferroptosis resistant subclones of B16F10 through serial exposure to RSL-3, a ferroptosis agonist. We showed that RSL-3 resistant B16F10 cells were indeed resistant to RSL-3 and erastin, two ferroptosis agonists (Supplementary Fig. S1D). We then inoculated parental or RSL-3 resistant B16F10 cells into C57BL/6 mice and treated established tumors with radiotherapy. We observed that resistance to ferroptosis significantly reduced the efficiency of radiotherapy (Fig. 1C). Further, RSL-3 resistant tumors failed to manifest a robust increase in lipid peroxidation following radiotherapy as compared to parental tumors (Fig. 1D). We additionally generated ferroptosis resistant ID8 subclones through serial exposure to erastin. We also showed that erastin resistant ID8 cells were resistant to RSL-3 and erastin (Supplementary Fig. S1E). We inoculated parental and erastin resistant ID8 cells into C57BL/6 mice and treated established ID8 tumors with radiotherapy. Again, we observed reduced radiotherapy efficacy in erastin resistant tumors (Fig. 1E). Together, these data suggest that radiotherapy induces tumor ferroptosis and also that ferroptosis is critical for radiotherapy efficacy.

To understand whether radiotherapy directly induces tumor ferroptosis, we performed clonogenic survival assay following radiotherapy in HT1080 human fibrosarcoma cells in vitro. We observed that ferroptosis inhibitors, liproxstatin-1 and ferrostatin-1, the lipophilic antioxidant trolox, and the iron chelator deferoxamine (DFO) increased clonogenic cell survival following radiotherapy (Fig. 1F). To determine whether radiotherapy directly affected lipid peroxidation, we conducted C11BODIPY staining in HT1080 cells in vitro. We observed that radiotherapy resulted in an increase in lipid ROS in HT1080 cells (Fig. 1G). We next examined clonogenic cell survival and lipid peroxidation in B16F10 and ID8 cells. Again, liproxstatin-1, as well as trolox and DFO, enhanced clonogenic survival in B16F10 (Supplementary Fig. S1F) and ID8 (Supplementary Fig. S1G) cells following radiotherapy. Radiotherapy induced an increased lipid peroxidation in B16F10 cells (Supplementary Fig. S1H) and ID8 cells (Supplementary Fig. S1I) in dose dependent manner. These results suggest that radiotherapy directly induces tumor ferroptosis.

Given that radiotherapy can induce tumor ferroptosis (Fig. 1A-1G and Supplementary Fig. S1A-S1I), we wondered whether ferroptosis agonists could enhance radiotherapy efficacy and be exploited as novel radiosensitizers. To explore this possibility, we pretreated HT1080, B16F10, and ID8 cells with low doses of ferroptosis inducers (FINs) including sulfasalazine, RSL-3, erastin, and atorvastatin, and then irradiated these cells. We found that ferroptosis inducers reduced clonogenic cell survival as compared to radiotherapy alone in HT1080 cells in vitro (Fig. 1H). GPX4 utilizes glutathione to reduce oxidized lipid species and to limit ferroptosis. Glutathione levels are regulated by intracellular cystine concentrations. We depleted cystine and cysteine using cyst(e)inase, a recombinant human enzyme (18). Interestingly, cyst(e)inase strongly sensitized HT1080 cells to radiotherapy in vitro (Fig. 1I).

To support our finding in HT1080, we treated B16F10 cells with sulfasalazine, RSL-3, and cyst(e)inase and observed decreased cell survival following radiotherapy as compared to radiotherapy alone (Supplementary Fig. S1J, S1K). Cell death quantification using propidium iodide staining confirmed that sulfasalazine and cyst(e)inase augmented radiation induced cell death (Fig. 1J). Pretreatment with RSL-3, sulfasalazine, cyst(e)inase also increased ID8 cell sensitivity to radiotherapy in vitro (Supplementary Fig. S1L, S1M). To examine the relevance of this finding in vivo, we established B16F10 tumors in C57BL/6 mice and treated mice with cyst(e)inase, radiotherapy, or the combination. While cyst(e)inase had minimal efficacy as monotherapy, it enhanced radiotherapy efficiency (Fig. 1K). Moreover, increased tumor control was accompanied with enhanced lipid oxidation in the combination therapy group (Fig. 1L). Neither single treatment nor the combination treatment caused significant murine weight loss (Supplementary Fig. S1N).

To assess whether FINs can improve radiotherapy efficacy in vivo, we established B16F10 tumors in mice, and treated tumors with radiotherapy, sulfasalazine, or both (dual treatment). We found that dual treatment enhanced tumor control (Supplementary Fig. S1O). Again, this improved tumoral control was accompanied with increased lipid oxidation in tumors treated with both agents (Supplementary Fig. S1P). These data suggest that ferroptosis can be targeted in vivo to enhance radiotherapy efficacy.

It has recently been shown that fatty acid saturation modulates ferroptosis sensitivity. Specifically, long-chain-fatty-acid-CoA ligase 4 (ACSL4) promotes ferroptosis, while ACSL3 limits ferroptosis (19,20). We generated ACSL4 knockout and ACSL3 knockout B16F10 cells (Supplementary Fig. S1Q, S1R). We observed that loss of the ferroptosis effector gene ACSL4 diminished radiotherapy efficacy in vivo (Fig. 1M). In contrast, deletion of the ferroptosis suppressor ACSL3 augmented radiotherapy efficacy in vivo (Fig. 1N).

To demonstrate that diminished cell survival following radiation was due to increased lipid peroxidation and ferroptosis, we quantified lipid peroxidation changes with C11BODIPY following FIN addition. We observed that in B16F10 cells, short exposures to low doses of sulfasalazine, RSL-3, and cyst(e)inase minimally altered lipid ROS. Combinations of FINs with radiation synergistically increased lipid ROS (Fig. 1O). FINs similarly augmented lipid ROS levels in concert with radiation in HT1080 cells (Supplementary Fig. S1S).

To generalize these findings to different radiation doses and fractionation schedules, we treated B16F10 tumors with 5 fractions of 3 Gy as well as a single fraction of 10 Gy or 20 Gy. Quantification of lipid peroxidation changes with C11BODIPY showed that all radiotherapy doses induced increases in C11BODIPY, while higher single fractions of radiotherapy induced more lipid peroxidation (Fig. 1P). To confirm that ferroptosis resistance conferred radiotherapy resistance at ablative doses of radiotherapy, we treated WT and RSL3 resistant B16F10 tumors with a single fraction of 20 Gy (21). We observed that even at higher doses of radiotherapy, ferroptosis resistant tumors remained resistant to radiotherapy in vivo (Supplementary Fig. S1T).

Collectively, these data suggest that radiation induces tumor ferroptosis and this ferroptosis can be pharmacologically augmented by ferroptosis agonists. Thus, targeting ferroptosis may be a novel therapeutic approach for radiation sensitization.

CD8⁺ T cells promote radiotherapy induced ferroptosis via IFN_γ

CD8⁺ T cells are required for efficacy of ablative doses of radiotherapy in vivo (21,22). The cellular and molecular bases for the interaction between T cells and radiotherapy are not fully defined. We first examined the involvement of CD8⁺ T cells in radiation therapy efficacy in B16F10 tumor bearing mouse model at moderate clinically utilized radiation doses. We subcutaneously inoculated B16F10 cells into C57BL/6 and established tumors. Then, mice received a moderate dose of radiation (8 Gy) alone or with anti-CD8 monoclonal antibody (mAb) administration prior to radiotherapy. As expected, radiotherapy reduced tumor volume (Fig. 2A). However, CD8⁺ T cell depletion with anti-CD8 mAb treatment largely abolished radiotherapy-induced tumor reduction (Fig. 2A). This radiation dose induced tumor cell ferroptosis (Supplementary Fig. S1B, S1H). To understand if CD8⁺ T cells promoted ferroptosis in concert with radiotherapy, we treated B16F10 cells with naïve or activated T cell supernatant. We found that exposure to a low dose of T-cell supernatant had minimal effect on B16F10 cell survival. Interestingly, T-cell supernatant synergized with radiotherapy to limit B16F10 clonogenic cell survival (Fig. 2B). Furthermore, T-cell supernatant and radiotherapy each independently enhanced lipid peroxidation in B16F10 cells, their combination further augmented lipid peroxidation (Fig. 2C). Blockade of IFN_γ signaling decreased T-cell supernatant-induced tumor STAT1 phosphorylation (Supplementary Fig. S2A) and diminished tumor lipid ROS production following combination treatment with T-cell supernatant and radiotherapy (Fig. 2C). These results suggest that CD8⁺ T cells may promote tumor ferroptosis and induce radiosensitization via IFN_γ.

To assess a direct link between IFN_γ signaling and radiotherapy efficacy, we treated ID8 cells with IFN_γ and radiotherapy and examined clonogenic cell survival. We observed that IFN_γ and radiotherapy synergistically reduced clonogenic survival (Fig. 2D). We also observed diminished clonogenic cell survival following IFN_γ and radiation treatment in B16F10 cells (Supplementary Fig. S2B). IFN_γ and radiotherapy synergistically increased lipid ROS levels in ID8 cells (Fig. 2E), B16F10 cells (Supplementary Fig. S2C), and HT1080 cells (Supplementary Fig. S2D). To confirm that IFN_γ increased radiation induced cell death, we quantified PI⁺ dying B16F10 cells following treatment with IFN_γ, radiotherapy treatment or the combination. We observed a synergistic increase in cell death following combination treatment of radiotherapy and IFN_γ (Fig. 2F). To implicate ferroptosis in cell death induced by the combination treatment of radiotherapy and IFN_γ, we conducted clonogenic survival in HT1080 cells with IFN_γ, radiotherapy, and liproxstatin-1. We observed that liproxstatin-1 administration blocked the synergistic reduction of cell survival observed upon treatment with IFN_γ and radiotherapy (Fig. 2G and Supplementary Fig. S2E). The data indicates that the combination of IFN_γ and radiotherapy regulates tumor cell ferroptosis in vitro. To examine that IFN_γ alters radiotherapy efficacy in vivo, we inoculated HT1080 cells into NOD-scid IL2γc-deficient (NSG) mice, and treated mice with recombinant human IFN_γ, radiotherapy, or the combination. While low doses of IFN_γ and radiotherapy slightly diminished tumor growth, the combination potently promoted tumor regression (Fig. 2H). To assess whether this is associated with tumor ferroptosis, we performed immunohistochemistry staining in tumor tissues and quantified 4-hydoxynoneal (4-HNE), a lipid peroxidation byproduct (23). Indeed, combination treatment with IFN_γ and radiotherapy significantly increased the tumoral 4-HNE levels (Fig. 2I and Supplementary Fig. S2F). To functionally demonstrate ferroptosis contributed to synergistic tumor regression in vivo, we inoculated HT1080 cells into NSG mice and treated mice with IFN_γ plus radiotherapy, liproxstatin-1, or all three agents. As expected, IFN_γ plus radiotherapy induced tumor regression. Importantly, this effect was abrogated by simultaneous treatment with liproxstatin-1 (Fig. 2J). Together, these data demonstrate that CD8⁺ T cells directly sensitize tumor cells to radiotherapy through IFN_γ by increasing lipid peroxidation and ferroptosis.

Radiotherapy and IFN_γ synergistically suppress system SLC7A11

We next sought to define the molecular mechanism by which IFN_γ sensitizes radiotherapy and promotes tumor ferroptosis. We first performed unbiased RNA sequencing in HT1080 cells following treatment with IFN_γ, radiotherapy, or the combination. Consistent with published results, interferon response genes, including CXCL9 and IRF1, were upregulated strongly by IFN_γ treatment (Fig. 3A) (24). As expected, radiotherapy induced an ATM dependent transcriptional program (25). Dual treatment with radiotherapy and IFN_γ surprisingly downregulated multiple cystine and cysteine transport channels, including a component of the cystine-glutamate transporter system xc⁻, SLC7A11 (Fig. 3A), which is a critical regulator of ferroptosis (26). Other ferroptosis regulators including GPX4, ACSL4, LPCAT3, and lipoxygenases (Supplementary Fig. S3A, S3B) were not altered in the same manner by dual treatment with radiotherapy plus IFN_γ treatment. We next validated this finding. Treatment with IFN_γ and radiotherapy individually reduced SLC7A11 transcripts (Fig. 3B) and protein levels (Fig. 3C) in HT1080 cells. Dual treatment resulted in a dramatic reduction of SLC7A11 in HT1080 cells (Fig. 3B, 3C). We also performed immunohistochemistry staining for SLC7A11 in HT1080 tumor tissues (Fig. 2H) and found IFN_γ and radiotherapy reduced SLC7A11 expression in vivo (Fig. 3D). To determine the functional relevance of SLC7A11 reduction, we examined radiolabeled cystine uptake in HT1080 cells. While both radiation and IFN_γ each reduced cystine uptake, dual treatments resulted in a maximal decrease in cystine uptake in HT1080 cells (Fig. 3E). Cystine is a precursor for glutathione production, which is a required cofactor for GPX4 activity (27). To confirm that diminished cystine import altered cellular antioxidant stores, we quantified glutathione level in HT1080 cells following treatment with radiation, IFN_γ, and the combination. We observed that the combination treatment significantly decreased glutathione level (Fig. 3F). Thus, radiotherapy and IFN_γ synergistically downregulate SLC7A11 expression in tumor cells resulting in diminished cystine transport and depletion of antioxidant stores in tumor cells to induce ferroptosis.

Next, we explored how radiotherapy downregulates SLC7A11 in tumor cells. Cells respond to radiotherapy through defined signaling cascades, including ATM activation (6,25,28). ATM has recently been shown to promote ferroptosis (29). We wondered whether ATM activation is involved in radiation induced ferroptosis by regulating SLC7A11. To test this possibility, we treated HT1080 cells with KU60019 (30), an ATM inhibitor. ATM inhibition abrogated radiotherapy-mediated downregulation of SLC7A11 transcription (Supplementary Fig. S3C) and protein levels (Fig. 3G). Indeed, treatment with KU60019 diminished radiotherapy-induced lipid peroxidation in HT1080 cells (Fig. 3H). Further, siRNA targeting ATM prevented downregulation of SLC7A11 transcription (Supplementary Fig. S3D) and protein (Fig. 3I) following radiotherapy. To confirm this mechanism in vivo, we treated wild type and shATM HT1080 tumors with radiotherapy (Supplementary Fig. S3E). We observed that wild type, but not ATM silenced tumors showed downregulation of SLC7A11 expression (Fig. 3J, Supplementary Fig. S3F). Further, wild type tumors showed increased 4-HNE lipid peroxidation in vivo following radiotherapy as compared to ATM silenced tumors (Supplementary Fig. S3G, S3H). Therefore, radiotherapy transcriptionally represses SLC7A11 expression via ATM to promote tumoral ferroptosis.

We have previously reported that IFN_γ negatively regulates SLC7A11 expression in a JAK-STAT1 dependent manner (16,31). To validate this finding, we silenced STAT1 with siRNAs (si-STAT1) in HT1080 cells and found that IFN_γ downregulated SLC7A11 in STAT1 dependent manner (Supplementary Fig. S3I and Fig. 3K). Interestingly, IFN_γ efficiently and comparably reduced SLC7A11 expression in both ATM-proficient and ATM-deficient HT1080 cells (Supplementary Fig. S3J). Conversely, regardless of STAT1 expression, radiotherapy efficiently reduced SLC7A11 expression in HT1080 cells (Supplementary Fig. S3K). Thus, radiotherapy and IFN_γ synergistically repress tumor SLC7A11 through ATM and STAT1 signaling, respectively. To confirm this mechanism in vivo, we treated wild type and STAT1 knockout HT1080 tumors with IFN_γ (Supplementary Fig. S3L). We observed that wild type but not STAT1 deficient tumors showed SLC7A11 downregulation following IFN_γ treatment (Fig. 3L and Supplementary Fig. S3M). We also observed that wild type tumors showed increased 4-HNE lipid oxidation following IFN_γ treatment as compared to STAT1 knockout tumors (Supplementary Fig. S3N, S3O).

To functionally connect SLC7A11 to tumor ferroptosis induced by radiotherapy, we used small hairpin RNAs against SLC7A11 (shSLC7A11) to knock down SLC7A11 in HT1080 cells (Supplementary Fig. S3P). We found that knockdown of SLC7A11 promoted radiotherapy-induced cell death in HT1080 cells (Fig. 3M) and B16F10 cells (Supplementary Fig. S3Q). This effect was rescued by liproxstatin-1 in HT1080 cells (Fig. 3M) and B16F10 cells (Supplementary Fig. S3Q). To determine the importance of SLC7A11 in radiotherapy in vivo, we used CRISPR-Cas9 to delete SLC7A11 in B16F10 cells. We confirmed deletion by quantifying SLC7A11 mRNA levels, and consistent with loss of system xc⁻ function, knockout cells were unable to export glutamate (Supplementary Fig. S3R, S3S). In line with previous reports (12,26), we found SLC7A11 deficient cells underwent spontaneous cell death, which can be rescue by liproxstatin-1 or 2-mercaptoethanol (BME) (Supplementary Fig. S3T). We inoculated SLC7A11 proficient and deficient B16F10 tumors into C57BL/6 mice, and treated mice with radiotherapy. Radiotherapy efficacy was enhanced in SLC7A11 deficient cells in vivo (Fig. 3N). Examination of lipid peroxidation showed that SLC7A11 deficient tumors had increased baseline levels of lipid ROS (Fig. 3O). Moreover, radiotherapy induced alterations in lipid peroxidation was further increased by loss of SLC7A11 (Fig. 3O). To confirm the importance of SLC7A11 to survival following combination IFN_γ and radiotherapy, we overexpressed SLC7A11 in HT1080 cells (Supplementary Fig. S3U). We observed that overexpression significantly increased clonogenic survival following treatment with radiotherapy and IFN_γ (Fig. 3P). To further confirm the importance of SLC7A11 expression, we performed rescue experiments in which SLC7A11 was expressed in SLC7A11 deficient cells (Supplementary Fig. S3V). Additionally, we observed that overexpression of SLC7A11 reversed the sensitivity of SLC7A11 deficient tumors to radiotherapy in vivo (Fig. 3Q). Collectively, these results suggest that radiotherapy activated ATM and IFN_γ induced STAT1 signaling jointly target SLC7A11 to modulate cystine uptake and promote tumor lipid peroxidation and ferroptosis.

Radiotherapy and immunotherapy synergistically induce tumoral ferroptosis

Immunotherapy augments radiotherapy efficacy in vivo, but the mechanisms through which immune checkpoint blockade enhances tumor cell death in irradiated lesions are not well defined (4,32,33). Checkpoint therapy including anti-PD-L1 and anti-CTLA4 activates CD8⁺ T cells and potentiates IFN_γ signaling pathway to control tumor progression (1). We hypothesized that radiotherapy and immunotherapy would synergize via ferroptosis in vivo to improve the efficacy of both therapies. To test this hypothesis, we established B16F10 tumors in C57BL/6 mice and treated mice with anti-CTLA4 mAb, radiotherapy, or the combination. As previously reported, anti-CTLA4 monotherapy was minimally efficacious in B16F10 tumors (32). However, we found that it potentiated radiotherapy efficacy in irradiated tumors (Fig. 4A). Consistent with our observation (Fig. 1), radiotherapy increased lipid peroxidation. Interestingly, the combination treatment with anti-CTLA4 and radiotherapy further enhanced tumoral lipid peroxidation (Fig. 4B). The combination of anti-CTLA4 and radiotherapy enhanced CD8⁺ T trafficking into tumors (Supplementary Fig. S4A, S4B). Radiotherapy increased tumor infiltrating Ki67⁺CD8⁺ T cells and this was augmented by anti-CTLA-4 (Fig. 4C and Supplementary Fig. S4C). Furthermore, we detected enhanced CD8⁺ T cell function following combined anti-CTLA4 and radiotherapy as marked by increased IFN_γ and granzyme B production (Fig. 4D, 4E and Supplementary Fig. S4D, S4E). Similarly, we observed that combination therapy increased CD4⁺ T cell number and IFN_γ expression (Supplementary Fig. S4F, S4G). To determine whether tumor ferroptosis contributed to the synergy between anti-CTLA4 and radiotherapy in vivo, we inoculated B16F10 tumors in C57BL/6 mice and treated them with the combination of anti-CTLA4 plus radiotherapy, liproxstatin-1, or all three treatments. Interestingly, liproxstatin-1 diminished the combined therapeutic efficacy of anti-CTLA4 and radiotherapy as shown by tumor volume (Fig. 4F), as well as tumor lipid peroxidation (Supplementary Fig. S4H). To extend our findings to an additional model, we inoculated ID8 tumors into the peritoneal cavities of C57BL/6 mice, and treated mice with anti-CTLA4, radiotherapy, or the combination. Anti-CTLA-4 and radiotherapy synergistically enhanced tumor control in vivo (Fig. 4G). Again, we observed increased lipid peroxidation following radiotherapy, and anti-CTLA4 mediated enhancement of lipid peroxidation following dual treatment with radiotherapy and anti-CTLA4 (Fig. 4H). To explore if other forms of immune checkpoint blockade also induced lipid peroxidation, we established B16F10 tumors in mice and treated the tumors with radiotherapy, anti-PD-L1, or both. While anti-PD-L1 or radiotherapy alone controlled tumor growth, the dual treatment was more effective in reducing tumor growth than any single therapy (Fig. 4I). We also detected the highest levels of tumor lipid peroxidation following combination therapy (Fig. 4J).

SLC7A11 deficient cells were more sensitive to radiotherapy in vitro and in vivo because of their increased ferroptosis susceptibility (Fig. 3). We hypothesized that ferroptosis sensitive tumors may benefit more from immune checkpoint blockade. To examine this, we established SLC7A11 proficient or deficient B16F10 tumors in C57BL/6 mice, and treated mice with anti-PD-L1. We detected higher tumoral lipid ROS in SLC7A11 deficient tumor cells than wild type tumor cells regardless of anti-PD-L1 therapy (Supplementary Fig. S4I). Furthermore, SLC7A11 deficient tumors were more sensitive to anti-PD-L1 therapy as shown by increased tumor C11BODIPY fluorescence and reduced tumor volume (Supplementary Fig. S4J) compared to wild type tumors.

To additionally support that ferroptosis is a mechanism of synergy between immunotherapy and radiotherapy, we inoculated parental or RSL-3 ferroptosis resistant cells into mice and treated mice with anti-PD-L1 and radiotherapy. We observed that in contrast to the parental cells, RSL-3 resistant B16F10 cells were largely insensitive to the combined treatment with anti-PD-L1 and radiotherapy (Fig. 4K). Moreover, this insensitivity was accompanied by diminished C11BODIPY in tumor cells (Fig. 4L). To define the mechanisms contributing to resistance of therapy, we established ovalbumin expressing parental or RSL-3 ferroptosis resistant B16F10 tumors and treated the mice with OT-1 transgenic T cells. We observed that OT-1 therapy controlled wild type but not RSL-3 resistant tumor. This suggests that ferroptosis resistant tumors are resistant to T cell effector function (Fig. 4M).

To examine that SLC7A11 modulated the induction of durable immune responses, we established parental or SLC7A11 deficient B16F10 tumors and treated them with radiotherapy and anti-PD-L1 therapy. We observed that SLC7A11 deficient tumors were more sensitive to combination therapy, with complete long-term responses in 56% (14/25) of SLC7A11 deficient tumor inoculated mice (Fig. 4N). To test whether these mice developed immunologic memory, wild type B16F10 tumors were re-inoculated in cured mice. We noted that tumors established in control but not previously cured mice, suggestive of the establishment of T cell memory (Fig 4O). Collectively, these data suggest that immunotherapy synergizes with radiotherapy to induce tumor ferroptosis and establish T-cell immunity.

Reagents

Ferrostatin-1 (347174-05-4), liproxstatin-1 (950455-15-9), trolox (53188-07-1), deferoxamine (138-14-7), RSL-3 (1219810-16-8), erastin (571203-78-6), sulfasalazine (599-79-1), atorvastatin (599-79-1), KU60019 (925701-46-8) were purchased from Cayman. The concentration used is shown in Supplementary Table. S1. C11BODIPY (D3861) was purchased from Invitrogen. Mouse recombinant IFN_γ (485-MI-100), anti-mouse IFN_γ block antibody (MAB485), human recombinant IFN_γ (285-IF-100) were purchased from R&D. Anti-mouse IFNGR1 antibody (16-1193-85) was purchased from Thermo Fisher. The following antibody were used for immunoblots: anti-human SLC7A11 antibody (12691), anti-β-actin (12262), anti-GAPDH (2118), anti-STAT1 (9172), anti-phospho-STAT1 (9167), anti-ATM (2873) and anti-ACSL4 (4047) were bought from Cell Signaling Technology. Anti-GPX4 (ab41787) was from Abcam. Additionally, IgG (BE0087, BE0090), anti-mouse CTLA4 (BE0131), anti-mouse PD-L1 (BE0101) and anti-mouse CD8 (BE0117) were purchased from BioXcell. 4-HNE antibody (ab46545) was purchased from Abcam. Cyst(e)inase was generated as previously described (18).

Cell lines

All the cell lines used here, including B16F10, ID8, HT1080 and HEK-293T cell lines, were bought from ATCC. Cells were regularly tested for Mycoplasma contamination using the Adenosine 5’-triphosphate (ATP) Bioluminescent Assay Kit (Sigma) every two weeks. Cells were thawed at early passage and cultured for up to 16 weeks in total.

Flow cytometry

To assess cell death, cells were treated as indicated, then collected and stained with Propidium Iodide (PI). For in vitro C11BODIPY analysis, cells were treated with test compounds for the indicated times, then harvested and incubated with C11BODIPY (581/591) for 20 min at 37 degrees. For ex vivo C11BODIPY analysis, tumors were filtered and underwent gradient centrifugation, then incubated with 2 μM C11BODIPY and anti-CD45 antibody. C11BODIPY FITC channel MFI data were normalized to untreated controls. To quantify T-cell cytokine production, mononuclear cells were isolated, stained, and analyzed as previously described (40).

Radiotherapy

Radiotherapy was delivered using a Philips RT250 model orthovoltage unit (Kimtron Medical) or a Small Animal Radiation Research Platform (SARRP, XStrahl Medical and Life Sciences) at a dose rate of approximately 2 Gy per minute as previously described (42). Experiments were performed in the Experimental Irradiation Shared Resource (University of Michigan). Focal irradiation was provided to flank tumors via lead shielding or collimation. Whole abdominal radiotherapy was provided to intraperitoneal tumors with shield above the diaphragm and below the pelvis.

Clonogenic survival assay

Clonogenic survival analysis was conducted as previously described. Normalized surviving fraction represented as relative to compound alone as previously described (42).

Animal studies

Six- to eight-week-old NSG and WT C57BL/6 mice were obtained from the Jackson Laboratory. OT-I colony was maintained internally. All mice were maintained under SPF housing with a maximum of five mice per cage. Mice were randomized to experimental groups when tumors reached 50 mm³ (HT1080, B16F10) or on day of first bioluminescent imaging (ID8). Tumor size was quantified and anti-PD-L1 was provided as previously described (16). For interferon experiments, 65 μg of human IFN_γ was given 24 hours prior to radiation (6-8 Gy, single fraction) as well as two days after radiotherapy. The primary endpoint of all experiments was tumor size. All the animal studies were conducted under the approval of the University of Michigan Committee on Use and Care of Animals (PRO00008278).

Activated T cell supernatant harvest

Single cells suspension was prepared from spleens and lymph nodes from OT-I mice, then stimulated with 5 μg/ml OVA peptide (Sigma, S7951) in RPMI1640 medium containing 10% FBS. After 3 days, CD8⁺ T cells were isolated via magnetically labeled antibodies (StemCell Technologies, 19853), then cultured in RPMI1640 medium containing IL-2 (R&D Biosystems) and 2-mercaptoethanol (Gibco, 21985023). After 3 days, harvest supernatant and filtered through 0.2 μm filter prior to subsequent use. For blocking, B16F10 cells were pretreated with 10 μg/ml anti-IFNGR as well as 10 ng/mL anti-IFNγ antibody.

Immunohistochemistry

Tumor tissues were harvested and fixed in 10% formalin and embedded in paraffin. Staining was performed on tumors treated with the indicated conditions using anti-4-HNE or anti-SLC7A11 antibody and counterstained using hematoxylin. The alkaline phosphatase optical density was quantified in imageJ. Results were normalized to untreated controls.

RNA sequencing

Cell lines were treated as indicated, and total RNA were purified using an mRNA kit (Qiagen). Libraries were prepared as previously described, and the edgeR package was used to generate reads per kilobase of transcript per million mapped reads (RPKM) values (43). Differential expression analysis was performed using the limma-R package as previously described. Minmax data normalization was used for heatmap generation. Gene set enrichment analysis (GSEA) was performed to identify significantly enriched gene sets (FDR<0.20).

Real time PCR

Total RNA was extracted using trizol and phenol-chloroform phase separation. Reverse transcription was performed using (Invitrogen, K1691). SYBR Green (Invitrogen, 4368702) was used for quantitative PCR with a StepOne Real Time PCR system (Applied Biosystems). β-actin was used as an internal control. Results are represented as fold change from untreated controls. The sequences of primers for qPCR are as follows:

Human SLC7A11 forward: 5'-TGCTGGGCTGATTTTATCTTCG-3',

Human SLC7A11 reverse: 5'-GAAAGGGCAACCATGAAGAGG-3',

Human β-actin forward: 5'-TGGTATCGTGGAAGGACTC-3',

Human β-actin reverse: 5'-AGTAGAGGCAGGGATGATG-3',

mouse ACSL3 forward: 5'-TCCTGGCTGCGATACACTTG-3'

mouse ACSL3 reverse: 5'-CCAAAGTCAAGGGCTCGGAT-3',

mouse β-actin forward: 5'-TGAGCTGCGTTTTACACCCT-3',

mouse β-actin reverse: 5'-TTTGGGGGATGTTTGCTCCA-3'

Generation of genetically modified cell lines

Transfected HEK-293T cells with a non-targeted retroviral vector, or a retroviral vector encoding shRNA targeting SLC7A11. Viral containing supernatant was harvested, and HT1080 cells and B16F10 cells with transduced and selected prior to subsequent use

Dharmacon shRNA sequence:

Human shSLC7A11: ATAATAAAGAGATAATACG

Mouse shSLC7A11: ATTAGCTGTATAACTCCAGGG

Crispr-Cas9 constructs (Santacruz Biotechnology) were utilized to generate B16F10 subclones deficient in ACSL4 (sc-424503-NIC), and SLC7A11 (sc-424104-NIC). In addition, 2-mercaptoethanol (Gibco, 21985023) was added to support SLC7A11 knockout cells growth. lentiCRISPR v2 (addgene, 52961) based constructs were used for knockout of ACSL3 in B16F10 cells, the designed gRNA sequences were GTAATGATATGCCGCAGACG and GGTGTGTACAATGACACCTT. HEK-293T cells were used to produce lentivirus. B16F10 cells were infected with scramble control or gene targeted lentivirus and selected with 2 μg/ml puromycin prior to subsequent use.

To perform the ectopic SLC7A11 rescue assay, a CMV promoter driven mouse SLC7A11-IRES-GFP construct was generated by Gibson Assembly. This construct was transfected into SLC7A11 deficient B16F10 cells. Clones were selected with G418 and fluorescent activated cell sorting.

RNA interference (siRNA)

HT1080 cells were transfected with control or siRNA targeting human ATM (4392420-S532284, Thermo Fisher) or STAT1 (SASI_Rn02_00204016, Sigma) using lipofectamine RNAiMAX (Invitrogen) prior to experimentation.

Radiolabeled cystine uptake assay

HT1080 cells were treated with either IFN_γ (10 ng/mL), or radiotherapy (10 Gy), or both for 24 hours. Medium was removed and changed to cystine-free medium. L-¹⁴C-Cystine (0.2 μCi/mL) was added and incubated for 15-45 minutes. Cells were washed and lysed. A Beckman liquid scintillation counter was used to quantify radioactive cystine uptake. Results normalized to total protein as determined by Pierce™ BCA Protein Assay Kit(Thermo, 23225).

Glutathione assay

HT1080 cells were treated with either IFN_γ (10 ng/mL), or radiotherapy (16 Gy), or both for 24 hours. Glutathione was quantified using the GSH-Glo assay (Promega). Results normalized to cell viability.

Glutamate assay

The Glutamate Colorimetric Assay Kit (BioVision, K629) was used for detection of extracellular glutamate released into the medium. Parental or SLC7A11 knockout B16F10 cells were seeded overnight in the presence of 2-mercaptoethanol to support knockout cell growth. Results normalized to cell viability.

Cell viability assay

Cells were collected and seeded into 96-well plates. To determine the effect of treatment on cell growth and viability, 10% volume of alamar Blue (Bio-Rad) was added directly into the medium and incubated for 4 hours. Absorbance at wavelengths of 570 nm and 600 nm was measured. Results were normalized to untreated controls and shown as relative cell viability (%).

Statistical analysis

Statistical analyses were performed with an unpaired Student's t-test for experiments with two groups (GraphPad Prism). One-way ANOVA was used to compare multiple experimental groups and two-way ANOVA was used to compare continuous outcomes across multiple experimental groups. For all tests, p<0.05 was considered significant. Sample size was not predetermined. Unless noted, samples were independent biological replicates.