Low-dose carboplatin modifies the tumor microenvironment to augment car t cell efficacy in human prostate cancer models

ABSTRACT Chimeric antigen receptor (CAR) T cells have transformed the treatment landscape for hematological malignancies. However, CAR T cells are less efficient against solid tumors,

largely due to poor infiltration resulting from the immunosuppressive nature of the tumor microenvironment (TME). Here, we assessed the efficacy of Lewis Y antigen (LeY)-specific CAR T cells

in patient-derived xenograft (PDX) models of prostate cancer. In vitro, LeY CAR T cells directly killed organoids derived from androgen receptor (AR)-positive or AR-null PDXs. In vivo,

although LeY CAR T cells alone did not reduce tumor growth, a single prior dose of carboplatin reduced tumor burden. Carboplatin had a pro-inflammatory effect on the TME that facilitated

early and durable CAR T cell infiltration, including an altered cancer-associated fibroblast phenotype, enhanced extracellular matrix degradation and re-oriented M1 macrophage

differentiation. In a PDX less sensitive to carboplatin, CAR T cell infiltration was dampened; however, a reduction in tumor burden was still observed with increased T cell activation. These

findings indicate that carboplatin improves the efficacy of CAR T cell treatment, with the extent of the response dependent on changes induced within the TME. SIMILAR CONTENT BEING VIEWED

BY OTHERS PSMA-TARGETING TGFΒ-INSENSITIVE ARMORED CAR T CELLS IN METASTATIC CASTRATION-RESISTANT PROSTATE CANCER: A PHASE 1 TRIAL Article 21 March 2022 THE POTENTIAL OF CAR T CELL THERAPY

FOR PROSTATE CANCER Article 08 July 2021 PSCA-CAR T CELL THERAPY IN METASTATIC CASTRATION-RESISTANT PROSTATE CANCER: A PHASE 1 TRIAL Article Open access 12 June 2024 INTRODUCTION

Immune-based therapies have transformed the treatment of many cancers, but not prostate cancer1. Chimeric-antigen receptor (CAR) T cell therapy, in which a patient’s own T cells are

genetically modified to target tumor-associated antigens, has greatly improved clinical outcomes in advanced acute lymphocytic leukemia and B cell lymphoma2,3. However, success in treating

solid tumors has been very limited due to the immunosuppressive tumor microenvironment (TME), which prevents T cell infiltration and activation at the tumor site4. Herein we address the

challenge of how to modify the TME to enhance CAR T cell recruitment and effector function using combination approaches. Target antigens for CAR T cells in prostate cancer include

prostate-specific membrane antigen (PSMA), prostate stem cell antigen (PSCA), KLK2, and six-membrane epithelial antigen of the prostate-1 (STEAP-1), mainly as monotherapies and with limited

success thus far5,6,7,8,9. In this study, we examine the efficacy of CAR T cells directed against the Lewis Y (LeY) glycolipid antigen. LeY is an onco-fetal glycolipid antigen with minimal

expression in normal tissues following birth; however, >50% of human carcinomas reactivate LeY biosynthesis at very high levels10. Studies in the 1990s demonstrated that LeY is

prominently expressed in many high-grade prostate cancers and their metastases11,12. LeY CAR T cells demonstrated high safety and tolerability in a Phase 1a clinical trial in patients with

LeY-positive acute myeloid leukemia13. In addition to ensuring antigen specificity, it is important to consider the immunosuppressive TME, which can interfere with T cell differentiation,

proliferation, and exhaustion in solid tumors14,15. To overcome this issue, novel approaches including the engineering of CARs or therapeutic combinations have been tested in preclinical

cancer models7,16. Promising effects have been observed with checkpoint blockade in various cancer types and are being tested in clinical trials (i.e., ACTRN12622001542785 by our team).

Checkpoint inhibitors effectively block immune checkpoints on T cells and their ligands on tumor cells (i.e. PD-1/L1) and checkpoint blockade allows CAR T cells to penetrate the TME, exert

their actions, and control tumor growth16. Alternatively, accumulating evidence suggests that chemotherapies can increase the immunogenicity of tumor cells and dampened the immunosuppressive

TME. Chemotherapies, including docetaxel, oxaliplatin, and carboplatin in prostate, lung, and ovarian cancer, have been shown to increase tumor antigen presentation, increase immune

checkpoint expression and promote immune cell infiltration and activation5,7,17,18,19,20,21,22. This shift from an immunologically cold to warm TME may prime the tumor to immunotherapies,

including CAR T cells. However, studies on pre- and post-treated ovarian cancer biopsies show different patterns of immune response to chemotherapy, suggesting that TME modulation may be

tumor-specific19. Therefore, whilst there is a strong rationale for modifying the TME with chemotherapy, further work needs to optimize their roles in enhancing immunotherapy and identifying

which tumors will respond to this treatment strategy. In this study, we tested the combination of LeY CAR T cells with either carboplatin, docetaxel, or the checkpoint inhibitor nivolumab

on tumor growth using patient-derived xenografts (PDXs) of prostate cancer. Our data show that carboplatin can synergize with CAR T therapy to facilitate early and persistent CAR T cell

infiltration and activation within select prostate tumors to maximize therapeutic response. RESULTS Using immunohistochemistry, we examined LeY expression in 800 prostate cancer specimens

from 739 patients. As cell surface expression of the target antigen is critical for CAR T cells to form an immune synapse, we quantified both membrane- and cytoplasmic LeY staining in this

unselected patient cohort. 12.6% of tumors had \(\ge\)10% membrane LeY-positive cells (the cut-off for admission into our current Phase I clinical trial in LeY-positive lung cancer); more

specifically, 12.3% (87/709) and 20% (6/30) of patients with localized and metastatic disease respectively demonstrated this level of membrane staining (Fig. 1a). However, in 49 PDXs from

the Monash Urology Research Alliance (MURAL) collection, which are typically derived from more aggressive cases of prostate cancer23, 57% (28/49) had \(\ge\)10% membrane LeY-positive cells,

with far higher expression in many specimens, based on percent cell positivity and staining intensity (Fig. 1b, c). LeY-positive specimens included AR-null tumors as well as AR-positive

tumors (Fig. 1c), and its expression was independent of AR or PSMA (Fig. 1c). This suggested that anti-LeY CAR T cells may provide another treatment option for patients who would not benefit

from AR- or PSMA-directed therapies, or who had progressed following these therapies. To show the on-target action of LeY CAR T cells in vitro, we used organoids derived from PDXs, which

maintain LeY expression (Fig. 2a and Supplementary Fig. 1a, b). We selected six PDXs with different phenotypes and genomic features (Fig. 1c and Supplementary Fig. 1c)23, including three

androgen receptor (AR)-positive adenocarcinomas and three AR-null tumors that express neuroendocrine markers, with a range of LeY expression (Supplementary Fig. 1a, b). Co-culture with LeY

CAR T cells, but not control empty vector T (ev-T) cells, caused morphological destruction of organoids with pronounced propidium iodine uptake, indicating secondary necrosis (Fig. 2b–d).

Importantly, tumor cell death correlated with LeY expression, with strongest PI uptake in organoids with high (\(\ge\) 90%) LeY expression (Fig. 2d, Supplementary Fig. 1a, b). In contrast,

PDX-435.1A-Cx organoids, with 10% LeY expression, showed a weaker response, and PDX-201.1A-Cx organoids, with 5% LeY expression, had no response to LeY CAR T cells (Fig. 2d and Supplementary

Fig. 1a, b). This is consistent with data from cell lines, with lysis of the LeY-positive prostate cancer cells PC-3, DU-145, and 22Rv1, but not the LeY-negative melanoma cell line

MDA-MB435, when co-cultured with LeY CAR T cells (Supplementary Fig. 1d, e). The mechanism of cell death in organoids involved the perforin/granzyme-dependent granule exocytosis pathway as

granzyme B and/or perforin inhibitors markedly reduced cell death in PDX-287R and PDX-472M organoids (Fig. 2e). Since LeY can be targeted on primary prostate tumors in vitro, we next

administered LeY CAR T cells to mice bearing PDX-287R grafts derived from a patient with aggressive adenocarcinoma, as previously described23. PDX-287R has 70% LeY expression by

immunohistochemistry and 95% by flow cytometry (Fig. 1c and Supplementary Fig. 2a). As expected, CAR T cells administered alone did not inhibit tumor growth (Fig. 3a). As immune checkpoint

blockade is often combined with CAR T cells in the setting of solid cancers24, we combined CAR T therapy with nivolumab, an anti-PD-1 antibody that blocks its inhibitory interaction with its

ligand, PD-L1. Despite the tumors expressing PD-L1 (Supplementary Fig. 2b, c), nivolumab had no effect on tumor growth alone or in combination with LeY CAR T cells (Fig. 3b). We next

trialed CAR T cells in combination with a single dose of the taxane-based chemotherapy docetaxel, or the platinum-based chemotherapy carboplatin, given a week prior to CAR T cell infusion.

Docetaxel given alone, or with LeY CAR T cells, had no effect on tumor growth (Fig. 3c). Carboplatin treatment alone or with ev-T cells induced an initial decrease in tumor burden that was

sustained for up to 3 weeks before tumors grew back (Fig. 3d, e and Supplementary Fig. 2d). In contrast, carboplatin with CAR T cells caused tumors to regress to <1% of their starting

volume, with no regrowth for 6 weeks post-treatment (Fig. 3d, e). Following LeY CAR T cell plus carboplatin treatment only scattered tumor cells could be detected in 5 of the 6 residual

grafts (Supplementary Fig. 2e), with phospho-histone H3 (pHH3) staining indicating greatly decreased proliferation compared to tumor cells in other treatment groups (Fig. 3f). Residual tumor

cells lacked or had minimal LeY expression (Supplementary Fig. 2f). However, treatment with carboplatin alone did not alter LeY expression in tumors in vivo, as well as in organoids

following short-term carboplatin treatment in vitro, suggesting that the lack of LeY expression in residual tumors was not due to carboplatin treatment (Supplementary Fig. 2g–i). These data

showed that pre-treatment with carboplatin could negate the poor therapeutic response of CAR T cells alone in PDX-287R. To determine whether this combination therapy is effective in other

PDX lines, we treated PDX-224R-Cx, a primary neuroendocrine cancer responsive to LeY CAR T cells in vitro (Fig. 2d). Consistent with PDX-287R, LeY CAR T cells alone did not alter tumor

growth (Fig. 3g). Carboplatin treatment alone caused an initial regression in tumor burden that was sustained for a week before grafts grew back (Fig. 3g). Treatment with carboplatin plus

LeY CAR T cells significantly decreased tumor burden compared to no T cell control, carboplatin, LeY CAR T cells alone and carboplatin with ev-T cells at the end of the treatment period

(Fig. 3g). However, in contrast to PDX-287R, tumors continued to grow across the experimental period (Fig. 3g), suggesting that the response to carboplatin-CAR T cell therapy may depend on

the effect of carboplatin in individual tumors. To investigate this, we first analyzed T cell recruitment and activation, which is a major barrier to CAR T cell therapy in solid tumors. We

assessed the localization of recruited LeY CAR T cells in PDX-287R grafts by staining for the human T cell marker CD3. There was significant enrichment of LeY CAR T cells at the tumor site 6

weeks after treatment with carboplatin, compared to all other treatment groups and controls (Fig. 4a, b), demonstrating that the carboplatin-induced long-term CAR T retention at the graft

sites. To determine if CAR T cells homed directly to the tumor, or if infiltration was secondary to tumor shrinkage, we assessed LeY CAR T cell infiltration 48 h after infusion. Remarkably,

we found high numbers of LeY CAR T cells in grafts 48 h after infusion following pre-treatment with carboplatin compared to grafts treated with CAR T cells alone (Fig. 4c, d), with

concurrent accumulation of mouse F4/80+ macrophages (Fig. 4e, f). Using flow cytometry, we showed that the presence of CAR T cells in the tumor was accompanied by a decrease in CAR T cells

in the spleen and peripheral blood, when compared to CAR T cells administered alone (Fig. 4g). Elevated expression of T cell activation markers CD25 and CD137, and the production of

pro-inflammatory Th1 cytokines IL-2, TNF-α, and IFN-γ, at the tumor site were also consistent with heightened T cell activation when administered after carboplatin (Fig. 4h, i and

Supplementary Fig. 3a–d). In contrast to PDX-287R, similar analysis on PDX-224R-Cx grafts 48 h post-CAR T cell infusion demonstrated that carboplatin failed to increase the infiltration of

CAR T cells into PDX-224R-Cx tumors (Fig. 4j). However, the limited CAR T cells that were present in the tumors were activated following carboplatin treatment (Fig. 4k, l and Supplementary

Fig. 3e–h). Therefore, while TME-modulation with carboplatin increased the activation of LeY CAR T cells in PDX-224R-Cx, the response was restricted by a lack of CAR T cell infiltration into

the tumor. We next interrogated changes in the TME using flow cytometry seven days after carboplatin treatment, concurrent to the time of CAR T cell infusion. Carboplatin-treated PDX-287R

grafts had a significant decrease in the proportion of EpCAM+ tumor epithelial cells 1 week following treatment compared to vehicle-treated grafts (Fig. 5a, b). This was accompanied with a

significant increase in hFAS-positive tumor cells (Fig. 5c and Supplementary Fig. 4a), a death receptor associated with extrinsic apoptosis. In contrast, despite PDX-224R-Cx initially

responding to carboplatin treatment in vivo (Fig. 3g), by 1 week after treatment there was no change in the proportion of tumor cells or hFAS expression (Fig. 5d–f and Supplementary Fig. 

4b). To investigate the response of tumor cells to carboplatin treatment in more detail, RNA-sequencing was performed on FACs-isolated tumor cells from PDX-287R at 1 week post-treatment.

Carboplatin resulted in the differential expression of 2017 genes in tumor cells, with 982 upregulated and 1036 downregulated genes (Fig. 5g and Supplementary Data 1). Consistent with the

increase in FAS expression observed by flow cytometry, gene set enrichment analysis showed an enrichment for apoptotic pathways, with significant increased expression of the pro-apoptotic

genes _FAS, BAX, BBC3, IFI6_, and _JUN_ (Fig. 5h and Supplementary Fig. 4c). Gene set enrichment analysis also revealed enrichment of interferon response, JAK/STAT signaling, p53, and

chemokine signaling pathways (Supplementary Fig. 4c). Previously, carboplatin has been shown to activate the cGAS/STING pathway, resulting in an increase in chemokine expression by lung

tumor cells22. Here, we also found an increase in STING signaling, with an enrichment for the cytosolic DNA sensing pathway and increased expression of _STING1, STAT1_, and _STAT2_ (Fig. 5i

and Supplementary Fig. 4c). There was also a significant increase in the expression of genes involved in T cell chemotaxis, _CXCL10, CXCL11, CCL20_ (Fig. 5j and Supplementary Data 1). This

increase in chemokine expression may contribute to early recruitment of CAR T cells into the tumor. Collectively, carboplatin-induced cell death in tumor cells likely initiated a

pro-inflammatory phenotype in prostate cancer PDXs. At 1 week post-treatment, flow cytometry analysis also showed that carboplatin caused an influx of CD45+ immune cells into PDX-287R grafts

(Fig. 5a and Supplementary Fig. 4d). The majority of immune cells in both control and treated grafts were F4/80+ cells of the myeloid monocytic lineage, and RNA seq of isolated immune cells

demonstrated no change in macrophage phenotype between control and treated grafts (Supplementary Fig. 4e–h and Supplementary Data 2). In contrast, in PDX-224R-Cx grafts there was no change

in CD45+ immune cell infiltration following carboplatin treatment (Fig. 5d and Supplementary Fig. 4i). PDX-287R grafts treated with docetaxel also showed no change in tumor cell proportion

and FAS expression at 1 week post-treatment, and a reduction in immune cell infiltration was observed (Supplementary Fig. 4j–n). Therefore, a response to combination therapy with CAR T cells

may depend on initial sensitivity to the chemotherapeutic agent, with tumor cell death at the time of CAR T cell infusion required to promote T cell recruitment into the tumor. To further

understand the long-term changes following carboplatin treatment in PDX-287R that may facilitate CAR T cell retention and the sustained response to LeY CAR T cells, we used scRNA-seq to

transcriptionally define immune and non-immune cell populations and anti-tumor innate immune signaling within the TME 3 weeks post-carboplatin treatment, when the response to carboplatin was

maximal. Consistent with the previous experiment, a single dose of carboplatin initially induced a decrease in tumor burden before tumors grew back (Supplementary Fig. 5a). Indeed, by 3

weeks post-carboplatin treatment 97% (28/29) of grafts from across the two experiments had regressed from starting volume (Supplementary Fig. 5b). Given the tumor mass was considerably

reduced by this timepoint, we expected a substantial reduction in tumor cells. This was demonstrated by scRNA-seq, where epithelial cells comprised only 2.1% of the cellular content of the

graft, compared to 86.1% in vehicle control, and confirmed histologically (Fig. 6a, Supplementary Fig. 5c, and Supplementary Data 3). Unlike the 1 week time point, there was no significant

difference in the expression of pro-apoptotic genes; however, _BAX_ and _BAD_ were still upregulated suggesting ongoing apoptosis (Supplementary Fig. 5d and Supplementary Data 4). As the

epithelial cell content was reduced in carboplatin-treated PDX tumors, there was a proportionate increase in mouse stroma (97.9%) compared to vehicle control (13.9%; Fig. 6a, Supplementary

Fig. 5c, and Supplementary Data 3). This was also accompanied by changes in the relative populations of stromal cell types, with an increase in macrophages and monocytes, and fewer

fibroblasts, endothelial and perivascular cells (Fig. 6b and Supplementary Data 3). UMAP (Uniform Manifold Approximation and Projection) density-based clustering analysis separated

PDX-287R-associated mouse stromal cells, showing distinct clusters of stromal cell types with differential gene expression patterns between cells from carboplatin-treated and vehicle grafts

(Fig. 6c). The gene expression profile of each cell type was cross-referenced to known cell population markers25,26,27,28,29,30 to identify the cell types represented by each cluster in our

analysis (Supplementary Fig. 5e–g). In contrast to the epithelial cells, carboplatin treatment led to increased expression of pro-survival factors (_Bcl2, Mcl1, Ier3, Ier5, Hmox1_) in

stromal cell populations, thus preserving their viability and potentially providing a means of stabilizing and prolonging the anti-tumor response (Supplementary Fig. 5d, Supplementary Data 

5–7). In contrast to 1 week post-treatment, gene set enrichment analyses demonstrated that by 3 weeks following carboplatin treatment macrophages had undergone polarization to the

pro-inflammatory M1, but not M2, phenotype, with upregulated of _Cd68_, _Cd86_, and _MHCII (H2-Aa, H2-Eb1_, and _H2-Ab1)_ (Fig. 6d, e and Supplementary Data 5). Consistent with their M1

differentiation, the macrophages upregulated expression of the chemotactic factors _Ccl5_ and _Cxcl10_, responsible for recruiting T lymphocytes (in this instance CAR T cells) into the tumor

stroma (Fig. 6f). The transcriptional profile of cancer-associated fibroblasts (CAFs) was also substantially altered 3 weeks after a single dose of carboplatin. The proportion of CAFs in

carboplatin-treated grafts was markedly reduced (Fig. 6b, c), and there was a shift from a myofibroblast (myCAF) to a pro-inflammatory phenotype (iCAF) (Fig. 6g and Supplementary Data 6).

This involved the downregulation of genes encoding the immunosuppressive factors _Tgfß-1_ and _Vegfa_, as well as the _Loxl_ gene family, responsible for stabilizing the extracellular matrix

(ECM; Fig. 6h and Supplementary Data 6). Following carboplatin treatment, CAFs increased their expression of numerous MMP genes (_Mmp2, Mmp3, Mmp13, Mmp14, Mmp27_); proteases involved in

the degradation of the tumor-supporting matrix (Fig. 6h and Supplementary Data 6). Carboplatin-treated CAFs also upregulated the chemoattractant _Ccl2_, which has been associated with

promoting monocyte and macrophage infiltration of the tumor site, and _Irf1_ and _IL-1ß_, which enhance M1 polarization in macrophages (Fig. 6h and Supplementary Data 6). Thus, the

carboplatin-induced CAF phenotype orchestrates a coordinated anti-tumorigenic response, which facilitates the polarization of macrophages to a pro-inflammatory M1 phenotype, enables

infiltration of phagocytes to the tumor site, and alters the ECM to promote immune cell trafficking. Taken together, carboplatin induces multiple changes that could overcome the

immunosuppressive TME. CAR T cells gain entry to the tumor site through the process of transendothelial migration, and tumor-associated high endothelial venule (TA-HEV) cells have been

described to facilitate this trafficking30. Three weeks post-carboplatin treatment there was a reduction in the proportion of endothelial cells in the PDXs (Fig. 6b). However, multiple

phenotypic changes were observed. Firstly, endothelial cells showed decreased expression of mouse _Angpt2_ and _Egln1_ genes, which typically promote tumor cell metastasis and the

dysregulation of vascular stability (Fig. 6i and Supplementary Data 7). Furthermore, carboplatin-treated endothelial cells had increased expression of _Sele_ and _Selp_, which encode key

selectins involved in the initial “capture and rolling” of leukocytes along the endothelial layer (Fig. 6i and Supplementary Data 7). Downstream in the signaling pathway, carboplatin-treated

endothelial cells also upregulated endothelial adhesion molecules _Icam1_, as well as _Vcam1_ gene expression, both of which may enhance that typify the TA-HEV phenotype and could be

expected to enhance CAR T cells traversal through the endothelial layer (Fig. 6i and Supplementary Data 7). Collectively, we describe a cascade of anti-tumorigenic alterations in cell

content and gene expression in response to low-dose carboplatin in PDX-287R that demonstrate how carboplatin effectively primes the TME and promotes LeY CAR T efficacy (Fig. 7). In the

absence of carboplatin sensitivity, including upregulation of hFAS and immune cell infiltration, CAR T cell entry to the tumor is limited and the response to LeY CAR T cell treatment is

dampened. DISCUSSION Collectively, these studies demonstrate that LeY CAR T cells are effective at targeting prostate cancer cells when co-cultured in vitro and can eliminate a prostatic PDX

tumor in vivo when preceded by a single dose of carboplatin. Mechanistically, carboplatin induced a cascade of events following the initial apoptosis of tumor cells, resulting in a

pro-inflammatory, anti-tumor immune response that enables CAR T cells to enter the tumor site and exert cytotoxic effector functions on LeY-expressing tumor cells. Failure of carboplatin to

induce apoptosis at the time of CAR T cell infusion and decreased T cell recruitment into the tumor dampens, but does not abolish, the response to treatment. Therefore, carboplatin improves

the efficacy of CAR T cell treatment, with the extent of the response dependent on changes induced within the tumor. Trafficking of CAR T cells to the tumor is a major barrier to CAR T cell

therapy in all cold, immune excluded solid tumors, and our data further supports this notion. In PDX-287R, carboplatin treatment initially induced apoptosis within the tumor cells,

accompanied by a pro-inflammatory response, increased cGAS-STING signaling and increased expression of chemokines, including CXCR3 ligands for T cell chemotaxis. At this time, there was an

influx of mouse myeloid cells into the tumor, and enhanced recruitment and activation of human CAR T cells following carboplatin treatment. In contrast, carboplatin failed to increase tumor

cell apoptosis, myeloid cell recruitment and T cell infiltration in PDX-224R-Cx. Interestingly, the T cells present within the PDX-224R-Cx tumors still showed increased activation following

carboplatin, presumably accounting for the decrease in tumor burden in carboplatin-LeY CAR T cell treated grafts compared to treatment with LeY CAR T cells alone. Collectively, this

demonstrates that pro-inflammatory changes induced by carboplatin are mandatory to enable the CAR T cells to gain unfettered access to the LeY-positive tumor cells and eliminate the tumor.

Whilst tumor cell sensitivity to carboplatin is clearly a feature, the transcriptomic changes within the TME following carboplatin treatment strongly suggest that pro-inflammatory effects

are not solely dependent on cancer cell death. A single dose of carboplatin induced a cascade of events to sustain a pro-inflammatory microenvironment and T cell persistence. The increase in

macrophages initially observed at 1 week post-treatment was sustained, with a shift to the M1 pro-inflammatory phenotype by 3 weeks post-treatment. Indeed, carboplatin is known to activate

mouse macrophages31,32, and consequently, the increased production of macrophage-derived factors, _Ccl5_ and _Cxcl10_, likely mediated the recruitment of T cells through known mechanisms,

including TCR3 receptor recruitment33. The evidence from our animal studies strongly suggests that the myeloid compartment plays a key role in the mechanism of action of carboplatin. Whether

the response will be similar in humans is unknown, but it is of high relevance to the clinical translation of this combination therapy. In addition to the activation of macrophages,

carboplatin induced a range of downstream modifications to the TME, including the induction of an iCAF phenotype, involving the downregulation of immunosuppressive genes such as _Tgfβ-1_ and

_Vegfa_, and decreased expression of _LOXL_ genes and increased expression of _MMP_ genes, leading to ECM degradation. We also demonstrated the presence of tumor-associated high endothelial

venule (TA-HEV) cells that are known to effectively traffic lymphocytes to tumors30. Collectively, this cascade of dynamic alterations to the TME (Fig. 7), observed after carboplatin

treatment, resulted in the induction of a pro-inflammatory microenvironment and provides a mechanism by which human T cell efficacy is mediated in this immune-compromised setting. The

chemotherapeutic agents docetaxel and cyclophosphamide have been shown to induce immunogenic cell death, dampen the suppressive TME, and improve the efficacy of immunotherapy, including CAR

T cells and checkpoint blockade, in preclinical studies for prostate cancer5,7,21. Furthermore, docetaxel is currently in clinical trials in combination with immunotherapy for

chemotherapy-naïve patients with advanced prostate cancer, where it is well-tolerated and showing anti-tumor activity34. Here, we demonstrate that the response to chemotherapies as a

modulating agent is likely dependent on the response of individual tumors, with TME modifications at the transcriptional level likely playing a major role in CAR T cell infiltration and

tumor response. In contrast to carboplatin, docetaxel failed to induce apoptosis and myeloid cell infiltration in PDX-287R at 1 week post-treatment, and had no effect on tumor growth when

combined with CAR T cells. It is likely that a multimodal approach is required to enhance the efficacy of immunotherapy in prostate cancer, and a pro-inflammatory gene signature induced by

agents such as carboplatin may be helpful in identifying patients that may benefit from CAR T cell therapy. Interestingly, PDX-287R has a mutation in the mismatch repair gene mutS homologue

2 (_MSH2_), which may be associated with response to immune checkpoint inhibitors in a range of solid tumors35. This may have contributed to the response seen here and needs to be

investigated in other prostate tumors with mismatch repair deficiency. The sequencing and timing of combination therapies may also influence response, and future studies are required to

optimize their roles in enhancing CAR T cell therapy, for example, the use of real-time analyses to provide more dynamic insights into the spectrum of responses across time. In this study,

the action of carboplatin was assessed in immunosuppressed NSG mice, which lack T cells, B cells, and natural killer cells. This model allows us to compare the response of individual patient

tumors to treatment, and thus it is more reflective of tumor heterogeneity seen in the clinic. In addition, a potentially unappreciated benefit of this immunosuppressed model is that it

allows the focus to be on the interaction between the tumor and human CAR T cells without a cognate effector immune response being induced through the murine lymphoid compartment. This lack

of lymphoid cells was advantageous as it did not obscure the major shift we saw in the phenotypes of host myeloid cells, fibroblasts, ECM, and endothelial cells in response to carboplatin.

This is likely to be a critical facet of the mechanism of action and is highly relevant to the human setting. However, this model is limited by the lack of crosstalk between immune cells

within the TME, including the dynamic interaction between immune activation and suppression, and future studies need to be extended into an immunocompetent syngeneic setting. In this study,

we examined the effect of bulk transduced T cells using IL-2; however, future studies should also consider the effectiveness of selected CAR T cell products with defined subsets of T cells

that possess increased memory capability36. Evidence is emerging to show that maintaining CAR T cells in an early, less-differentiated state in vitro results in superior persistence,

proliferation, and anti-tumor effects in vivo36,37. Although effector memory T cells have strong cytolytic capacity, less differentiated T cells, such as stem cell memory T cells, are

critical for in vivo expansion and long-term persistence37,38. LeY is not a predicted target antigen for prostate cancer treatment. Unlike other cell-surface prostate adenocarcinoma targets,

such as prostate stem cell antigen (PSCA), prostate-specific membrane antigen (PSMA), and six-transmembrane epithelial antigen of the prostate (STEAP-1)7,39, LeY is expressed on a broad

spectrum of tumors. In general, expression of PSCA, PSMA and STEAP-1 is limited to AR-positive adenocarcinoma tumors and is rarely detected in AR-null or neuroendocrine prostate cancer,

limiting their clinical application. Here, we demonstrate that LeY expression is detected in both adenocarcinoma and neuroendocrine prostate cancer, and is independent of AR or PSMA, and

thus LeY CAR T cells could be applicable for the treatment of a diverse range of prostate tumors. Here, PDX-224R, a de novo neuroendocrine tumor, had a weaker response to combination therapy

compared to PDX-287R. However, this is more likely dependent on chemotherapy response given that both adenocarcinoma and neuroendocrine tumors were sensitive to LeY CAR T cells in vitro. A

larger panel of PDXs is required to determine whether this response is phenotype dependent. This is clinically relevant as LeY CAR T cells may provide another treatment option for patients

who would not benefit from AR- or PSMA-directed therapies, or who had progressed following these therapies. Overall, carboplatin induces a cascade of changes within the tumor to increase the

efficacy of CAR T cell treatment, with the extent of the response dependent on TME modifications to promote T cell infiltration, activation and persistence. These data provide preclinical

proof-of-concept evidence for the use of carboplatin as a modulating agent for CAR T cell immunotherapy in the treatment of prostate cancer. METHODS PATIENT-DERIVED XENOGRAFTS

Serially-transplantable PDXs were established and characterized by MURAL, as previously described23. Original specimens of localized prostate cancer tissue were collected from patients

undergoing radical prostatectomy or transurethral resection of the prostate, and metastatic prostate cancer tissue was obtained from biopsy, palliative surgery, or rapid autopsy through the

CASCADE program40. Informed, written consent was obtained from patients prior to tissue collection according to human ethics approval from the Cabrini Institute (03-14-04-08), Monash

University (1636), and Peter MacCallum Cancer Centre (15/98, 97_27), and patient details, including age and treatment history, have been previously published23. All PDX experiments were

performed in 6–8-week-old male non-obese diabetic severe-combined immune-deficient (NSG) mice (RRID:IMSR_JAX:005557)41. NSG mice were purpose bred at Monash Animal Research Laboratories

(Monash University, Monash Breeding Colony approval number MMCA 209/25BC and 15160). All animal care and procedures for the establishment and maintenance of PDXs were performed in accordance

with Monash University animal ethics approvals (MARP/2014/085, MARP/2014/119, MARP/2016/016, 17963, 28911). All mice were bred and housed in a PC2-specific pathogen-free facility under

controlled temperature (22 °C) and lighting (12:12 h light-dark cycle), and were fed a chow diet _ad libitum_. Experimental mice and control mice were co-housed. NSG mice either had a 5 mm

testosterone pellet implanted subcutaneously to supplement host testosterone levels, or were castrated to mimic androgen deprivation. Routine validation of PDX authenticity included DNA and

RNA-sequencing, STR profiling and pathological assessment including biomarker expression for androgen receptor (AR), prostate specimen membrane antigen (PSMA) and the neuroendocrine markers

CD56, synaptophysin and chromogranin A, as previously described23. Genomic alterations in PDXs, determined by targeted DNA sequencing, was previously published23. IMMUNOHISTOCHEMISTRY FOR

LEWIS Y LeY expression was analyzed in 709 primary prostate cancer specimens on the Case PSA Progression tissue microarray (obtained from the Prostate Cancer Biorepository Network) and 91

metastatic prostate cancer specimens from 32 patients in our own clinical collection. LeY expression was also analyzed in 49 PDXs obtained from MURAL derived from 14 high-risk localized and

35 metastatic tumors. Detection of the LeY antigen was performed by immunohistochemistry on formalin-fixed tissue using a humanized monoclonal IgG1 antibody m3S193 (1.9 µg/ml; provided by

Ludwig Institute for Cancer Research) on the Dako Autostainer Link 48 with EnVison FLEX, High pH Link visualization system reagents42. The staining protocol was adapted and validated from a

previously published study43. Briefly, antigen retrieval was performed at 10 oC for 20 min, peroxidase blocking for 10 min, antibody or negative control reagent for 30 min at 1.9 µg/ml,

EnVision Flex HRP polymer for 30 min, DAB for 5 min and finally sections were counterstained using Mayer’s Hematoxylin. Normal human lung, NSCLC squamous, and adenocarcinoma control tissues

were used for staining quality control throughout the study. ESTABLISHMENT OF ORGANOIDS FROM PDX TISSUES Organoids were established from PDX tissues as previously described23,44. Fresh or

frozen-thawed single PDX cells were seeded in growth factor reduced, phenol red-free, ldEV-free Matrigel (Corning), with 1–1.5 × 105 cells seeded per 20 µl of Matrigel in 48 well plates.

Organoids were cultured in organoid media, consisting of advanced DMEM/F-12 media (Thermo Fisher) containing 1% penicillin-streptomycin, 2 mM Glutamax, 1 nM DHT, 1.25 mM _N_-acetylcysteine,

50 ng/ml EGF, 500 nM A83-01, 10 mM nicotinamide, 10 μM SB202190 (Sigma), 2% B27 (Life Technologies), 100 ng/ml noggin (Peprotech), 10 ng/ml FGF10 (VWR), 5 ng/ml FGF2, 1 μM prostaglandin E2

(Tocris), and 10% R-spondin 1 conditioned media. 10 μM Y-27632 dihydrochloride (Selleck Chemicals) was added to culture medium during organoid establishment and the first subsequent passage.

To assess LeY expression, organoids were fixed in 10% formalin for at least 1 h, and then embedded in paraffin before performing immunohistochemistry for LeY, as described above. CAR T CELL

PRODUCTION The anti-LeY scFv-CD3ζ-CD28 CAR construct and retroviral packaging cell line PG13-LeY CAR was used in this study as described previously10. Human PBMCs were isolated from normal

donor buffy coats (Australian Red Cross Blood Service) by density gradient centrifugation (Ficoll-Paque, GE Healthcare Life Science). PBMCs were stimulated with anti-human CD3 (OKT3 30 

ng/ml, Miltenyi) and 600 units/ml IL-2 (Peprotech) for 48 h and were transduced on day 3 and 4 with viral supernatant produced by PG13-LeY CAR cells on a Retronectin (Takara Bio, Otsu,

Japan) coated 6-well plates as per the manufacturer’s instructions. After 4 h incubation at 37 °C, viral supernatant was removed, and 2.5 × 106  T cells in 5 ml of fresh retroviral

supernatant with IL-2 (600 units/ ml) were added. 24 h after the second transduction, T cells were transferred to fresh RPMI 1640 (Gibco) supplemented with 10% FBS, penicillin/streptomycin,

Glutamax, and 600 units/ml IL-2. The media was changed every two days until day 13. In the LeY-CAR construct, a CD34 epitope was embedded for purification. During the CAR-T cell production,

the transduced T cells were purified using the EasySep™ Human CD34 Positive Selection Kit (STEMCELL Technologies) on day 4–5 post transduction. The purified cells were further expanded for

infusion. The percentage of CAR+ cells in the final product was no <98%. The gating strategy for CAR T cells is shown in Supplementary Fig. 6a. IN VITRO CAR T CELL KILLING ASSAY Once

organoids had reached approximately 50 μM in size, organoids were recovered from Matrigel using Cell Recovery Solution (Corning, USA #354253) for 1 h on ice. 50–100 organoids were plated in

10 µl of Matrigel to form a Matrigel dome in µ-Plate 96 well ibiTreat (ibidi GMbH, Germany #89626) with organoid media. In all, 3 × 104 LeY CAR T cells or control ev-T cells were added to

triplicate organoid wells with 300 units/ml IL-2. Live CAR T/ev-T cells can infiltrate into the dome, but the dead ones are excluded. Propidium iodide (PI, Sigma-Aldrich) was used as an

indicator for cell death (2 μg/ml) and 1% Triton-X100 was used to obtain maximum cell death (positive control). Live cell images were captured by IX83 widefield live-imaging microscope

(Olympus) over a 48 h period. Briefly, 10 fields were imaged per well at ×10 magnification at one time point and 25 time points were taken with 2 h interval. Images were analyzed using

Fiji45 and Bio-formats46. Approximately 50–100 organoids were analyzed individually per well and the MFI of PI of each organoid at each time point was measured. Organoids were gated

frame-by-frame to determine the PI uptake by dead organoid cells. For mechanism study, granzyme B inhibitor Compound 20 (10 μM) and perforin inhibitor SN34960 (20 μM) were used during the

assay. ANALYSIS OF LEY EXPRESSION ON TUMOR CELL LINES To analyze the LeY expression on different prostate cancer cell lines, flow cytometry was used (Supplementary Fig. 6b). Cells were

stained with Live/Dead Near-IR fixable viability dye (Thermo Fisher) for 10 min at 4 °C, then followed by a 30-min surface staining with a murine monoclonal antibody against human LeY (clone

3S193) conjugated with Alex Fluor 488. Cells were washed and fixed in 2% paraformaldehyde then resuspended in FACS buffer before acquisition on a LSR Fortessa (Becton Dickinson). Flow

cytometry data was then analyzed using FlowJo v10.8 software. 51CHROMIUM RELEASE ASSAY IN TUMOR CELL LINES Target cells (LeY+ cell lines DU-145, PC-3, and 22Rv1 and LeY- cell line MDA-MB435)

were labeled with 50 μCi 51Chromium (51Cr, PerkinElmer) for 1 h at 37 °C. The labeled cells were washed and subsequently co-cultured with LeY-CAR-T cells or empty vector-transduced T cells

in triplicate wells at effector to target ratios ranging from 50:1 to 6.25:1. Target cells alone (spontaneous release) and target cells with 10% Triton X-100 (maximum release) served as

controls. After 4 h coculture, supernatants were collected, and the amount of 51Cr released (experimental release) was detected using a gamma counter (Wallac Wizard 1470, PerkinElmer). The %

lysis was calculated by [(experimental release – spontaneous release)/(maximum release – spontaneous release)] × 100. PDX EXPERIMENTS All animal care and procedures were performed in

accordance with Monash University animal ethics approval 20374 and Peter MacCallum Cancer Centre animal ethics approval E647. To assess the long-term in vivo response to LeY CAR T cells,

tissue from PDX-287R and PDX-224R-Cx was established subcutaneously (1 graft/mouse) in male NSG host mice (6–8 weeks old) until tumor volume reached approximately 50–100 mm3. Mice received a

single dose of irradiation (0.5 Gy) for lymphodepletion on day 0, before receiving three doses of 1 × 107 adoptively transferred LeY CAR T cells or empty vector T cells in PBS by

intravenous injection on days 1, 2, and 3. Mice also received eight doses of 50000 IU of IL-2 by intraperitoneal injection on days 1–5, 7, 9, and 11. A subset of host mice were pre-treated

with 200 µg/dose nivolumab (1 dose every 5 days by intraperitoneal injection starting 5 days before day 1 of CAR T cell transfer, 4 doses in total, dissolved in PBS), 10 mg/kg docetaxel (1

dose 7 days before day 1 of CAR T cell transfer, dissolved in 90% PBS with 5% Tween 80 and 5% ethanol), or 50 mg/kg carboplatin (1 dose 7 days before day 1 of CAR T cell transfer, dissolved

in water). The dose and dosing regimen of nivolumab has been used previously in NSG mice and is an equivalent safe dose for humans47,48. The doses of carboplatin and docetaxel are lower

doses than in clinical practice to investigate the modulatory effects of these agents. IL-2 was purchased from Peprotech (Cranbury, New Jersey, USA), and nivolumab, docetaxel and carboplatin

were purchased from Selleck Chemicals (Houston, Texas, USA). Between 5–21 mice were present per treatment group across 1–3 independent experiments (see figure legend). Tumors were measured

three times a week for up to 6 weeks post-adoptive transfer of CAR T cells using calipers. Tumor volume was determined from caliper measurements by length × width × height × 0.52, as

previously described49, and animals were culled when tumor volume reached the maximum ethical limit of 1000 mm3. At tumor collection, animals were placed under isoflurane for blood

collection by cardiac puncture and then humanely killed by cervical dislocation. The investigators involved in treating mice were not blinded to the treatment groups but were blinded to

tumor measurement data. Harvested tumor tissue was formalin-fixed for histological analysis. For early time point experiments, mouse PDXs were established as described above. Mice were

treated with a single dose of 50 mg/kg carboplatin 7 days before the CAR T cell infusion and were irradiated 1 day before infusion. CAR T cells were given on day 1 and day 2 with 50,000

IU/dose IL-2. On day 3, 48 h after the first CAR T cell infusion mice were placed under isoflurane for blood collection by cardiac puncture, and then humanely killed by cervical dislocation.

Blood, spleen, and tumors were collected for further analysis. For mechanistic single-cell RNA-sequencing studies, mice were treated with a single dose of 50 mg/kg carboplatin or vehicle

control and tumor tissues were harvested on day 21. IMMUNOHISTOCHEMISTRY ON PDX TISSUE Immunohistochemistry for phospho-histone H3 was performed on three representative sections per graft

using the Leica BOND-MAXTM automated system (Leica Biosystems, Mount Waverley, Victoria, Australia). The primary antibody was a rabbit antibody against phospho-histone H3 (Ser10) Antibody

(#9701), Research Resource Identifier (RRID): AB_331535 (Cell Signaling Technology) used at 8 µg/ml. The number of phospho-histone H3-positive cells were counted using Aperio ImageScope

Software (Leica Biosystems), with three sections analyzed per graft, and expressed as a percentage of the total number of cells counted. Immunohistochemistry for human CD3, human CD8, and

mouse F4/80 was performed manually. FFPE slides were dewaxed in xylene and antigens were retrieved at 125 °C for 10 min in sodium citrate pH6 buffer (for CD3), Tris EDTA pH9 buffer (for

CD8), or Tris EDTA pH8 buffer (for F4/80). Endogenous peroxidases were inactivated with 3% H2O2 (Merck), and the slides were blocked in PKI blocking buffer (Akoya Biosciences). Primary CD3

antibody (6.7 µg/ml; clone SP7; Abcam #ab16669; RRID AB_443425), CD8 antibody (1:100; clone 4B11; Invitrogen #MA1-80231; RRID AB_929437) or F4/80 antibody (1: 100; polyclone, Abcam #100790;

RRID AB_10675322) was incubated 30 min at room temperature, and the secondary anti-mouse ImmPress (Vector Laboratories, MP-7402) or anti-rabbit ImmPress (Vector Laboratories, MP-7401) was

added for 30 min at RT, followed by Dako Liquid DAB for 15 min. The slides were then counter-stained by Jung Autostainer (Leica) and scanned using the Olympus VS120 microscope at 20×

magnification. FLOW CYTOMETRIC ANALYSIS OF CAR T CELLS To analyze CAR T cells in blood and spleen in vivo, flow cytometry was used for absolute cell counting and cell phenotyping. Blood and

smashed spleen were treated with ACK buffer to remove the red blood cells. Cells were stained with Live/Dead Near-IR fixable viability dye (Thermo Fisher) for 10 min at 4 °C, then followed

by 30-min surface staining for CD3 (1:100; BD Biosciences #564001), CD4 (BioLegend #317438), CD8 (1:100; BD Biosciences # 612889), CAR (as indicated by Flag-tag, 1:200; BioLegend #637310),

and CD137 (1:100; BioLegend #309814), CD25 (1:200; BioLegend #302629). Before analyzing the cells on Symphony (BD Biosciences), cells were mixed with CountBright Absolute Counting Beads

(ThermoFisher) for absolute cell count according to the manufacture’s instruction. Flow cytometry data was then analyzed using FlowJo v10.8 software. The gating strategy is shown in

Supplementary Fig. 6a. ANALYSIS OF TUMOR-INFILTRATING T CELLS To analyze CAR T cells infiltrating tumors by flow cytometry, mice were euthanized 48 h-post first infusion, and tumors were

digested using RPMI-1640 (Gibco) supplemented with 1 mg/ml collagenase type IV (Sigma-Aldrich) and 0.02 mg/ml DNase (Sigma-Aldrich). Tumors were incubated for 1 h at 37 °C, rendered to

single-cell suspension, and then filtered twice (through 70-μm filter). Cells were then analyzed by flow cytometry directly for phenotyping and absolute cell counting using CountBright

Absolute Counting Beads (ThermoFisher). For analysis of cytokines, single-cell suspensions were incubated on anti-idiotype antibody (0.5 μg/ml) or anti-CD3 antibody (1:100; clone OKT3, 0.5 

μg/ml) coated-plate overnight with 25 units/ml IL-2 and were treated with GolgiStop (BD Biosciences) for 6 h before intracellular staining for IL-2 (1:50; BioLegend #500348), TNF-α (1:100;

BioLegend #502912) and IFN-γ (1:100; BioLegend #502546). The gating strategy is shown in Supplementary Fig. 6a. ANALYSIS OF PD-L1 ON ORGANOIDS To investigate CAR T cell-induced PD-L1

expression on organoids, cells from PDX-287R organoids were prepared on Matrigel with organoid growth media in 24-well plate as described above. LeY CAR T cells were co-cultured with

organoids for 24 h, then the supernatant was collected as CAR T cell-conditioned media. The fresh organoids or prostate tumor line DU-145 cells (as positive control) were incubated with

either conditioned media or fresh media containing 150 ng/ml human IFN-γ (PeproTech) for 24 h. The cells were trypsinized and resuspended in single-cell suspension for flow cytometry

analysis using anti-PD-L1 antibody (1:100; BioLegend #393608; Supplementary Fig. 6c, d). DISSOCIATION OF PDXS FOR SINGLE-CELL ANALYSIS PDXs were harvested from host mice and cut into 2 × 2 

mm pieces using a scalpel. Tumor pieces were digested in RPMI-1640, containing 0.65 U/ml Liberase TM (Roche) and 0.2 mg/ml DNase I (Roche), for 1 hour at 37 °C, following by lysis of red

blood cells (RBCs) using RBC Lysis Buffer (Sigma) for 1 minute. Cells were then resuspended in PBS, 1 mM CaCl2, with 2% FBS and underwent negative selection for viable cells using the Easy

Sep Dead Cell Removal kit (Miltenyi), according to the manufacturer’s protocol. Viable cells were passed through a 30 µM cell strainer (Miltenyi) to remove cell clumps, and then counted

using Trypan blue. Samples with cell viability >80% were resuspended in PBS with 2% BSA and proceeded to single-cell analysis. FLOW CYTOMETRIC ANALYSIS AND FLUORESCENCE-ACTIVATED CELL

SORTING (FACS) OF PDXS AND ORGANOIDS ANALYSIS OF LEWIS Y EXPRESSION ON PDX AND ORGANOIDS PDX-287R organoids were prepared on Matrigel with organoid growth media in 24-well plate as described

above. Organoids were treated with 10 µM carboplatin for 24 or 72 h in triplicate. PDX tissue or treated organoids cultures were digested into single-cell suspensions as above. Cells were

incubated with pre-titrated anti-human Lewis Y-Alexa Fluorochrome 647 (1:50–1:100; murine IgG3 monoclonal antibody, close 3S193) and IgG3-APC isotype control antibodies. Cells were incubated

on ice in the dark for 15 min, washed with FACS buffer, and centrifuged for 5 min at 200 g to collect cells. Cells were filtered into 5 ml round-bottom tubes with strainer cap (Falcon®)

prior to analysis. 100 ng/ml of propidium iodide (Sigma-Aldrich) was used to exclude dead cells. Samples were analyzed on the BD® LSR II Flow Cytometer. At least 2 × 103 live events were

recorded for each file for downstream analysis using FlowJoTM v10.8.1. The gating strategy is shown in Supplementary Fig. 6c. ANALYSIS OF PDXS FOLLOWING CARBOPLATIN TREATMENT PDX-287R and

PDX-224R mice were established in host mice before being treated with a single dose of 50 mg/kg carboplatin or vehicle control. The tumor was harvested 7 days after treatment and digested

into single-cell suspensions as above. Digested cells were incubated with pre-titrated antibodies diluted in staining buffer (1 x PBS with 10% FCS and 5 mM EDTA). Cells were pre-blocked for

5 min on ice with purified rat anti-mouse CD16/CD32 (25 mg/ml; clone 2.4G2, Cat #553141, BD Biosciences, USA) to block murine Fc receptors. Cells were then stained with the following

antibodies (all antibodies are purchased from BioLegend unless otherwise stated); anti-human EpCAM-PECy7 (1:200; clone 9C4), anti-human CD95-BUV395 (4 mg/ml; clone DX2, BD Biosciences),

anti-mouse MHC I-Ak-PE (0.2 mg/ml; clone 10-3.6), anti-mouse F4/80-APC (1 mg/ml; clone BM8), anti-mouse CD45-APCCy7 (1 mg/ml; clone 30-F11), anti-mouse ICAM1-BV421 (2 mg/ml; clone

YN1/1.7.4), anti-mouse CD31-BV510 (2 mg/ml; clone 390, BD Biosciences) or isotype controls; mouse IgG1k-BUV395 (4 mg/ml; clone X40, BD Biosciences), Rat IgG2bk-BV421 (2 mg/ml; clone

RTK4530). Cells were incubated on ice in the dark for 15 min, washed with FACS buffer, and centrifuged for 5 min at 200 g to collect cells. Cells were filtered into 5 ml round-bottom tubes

with strainer cap (Falcon®) prior to analysis. In all, 100 ng/ml of propidium iodide (Sigma-Aldrich) was used to exclude dead cells. Samples were analyzed on the BD® LSR II Flow Cytometer.

At least 2 × 103 live events were recorded for each file for downstream analysis with FlowJoTM v10.8.1. For FACS, samples were sorted on the BD FACSAria™ Fusion Flow Cytometer at no more

than 5 × 103 cells per second through a 100-micron nozzle at 20 psi. The following gating strategy was used to purified cell subsets; debris was excluded via FSC and SSC gate, PI was used to

excluded dead cells and single cells were gated based on FSC-H and FSC-A parameters (Supplementary Fig. 6e). FACS purification of; tumor cells (EpCAM+ CD45-), immune cells (EpCAM- CD45+),

stromal cells (EpCAM-, CD45-, CD31-) and endothelial cells (EpCAM-, CD45-, CD31+). Samples were collected in 30% (v/v) FCS in RPMI, recovered by centrifugation, counted and analyzed for

purity. Populations were sorted to greater than 90% purity. RNA ISOLATION Total RNA from FACS purified subsets were isolated using the RNAqueous Total RNA Isolation Kit (Ambion) according to

the manufacturer’s instruction. Total RNA was quantified in a Nanodrop ND-1000 spectrophotometer, checked for purity and integrity in a Bioanalyzer-2100 device (Agilent Technologies). BULK

RNASEQ ANALYSIS MULTIPLEX RNA-SEQUENCING Samples with a RIN (RNA Integrity Number) \(\ge\) 6, as determined by the Bioanalyzer-2100, were incorporated into the RNA library for associated

downstream Multiplex RNA-sequencing and analysis. The 3’ end of poly(A) primed transcripts were initially hybridized with a custom primer, 8 bp sample index and 10 bp unique molecular

identifier (UMI). Subsequently, during first complementary DNA (cDNA) strand synthesis, a modified reverse transcription reaction facilitated the addition of a template switching sequence to

the 5′ end of the RNA transcripts. The amplified cDNA libraries were subsequently tagmented utilizing PCR transposase. Indexed complementary DNA (cDNA) was selected for and amplified

utilizing Illumina P5 (5′ AAT GAT ACG GCG ACC ACC GA 3′) and P7 (5′ CAA GCA GAA GAC GGC ATA CGA GAT 3′)50. Multiplex RNA-sequencing was performed on the Illumina NextSeq550 by the Medical

Genomics Facility (Monash Health Translation Precinct). MULTIPLEX RNA-SEQUENCING ANALYSIS The raw Fastq files were quality checked using FastQC and low-quality reads were trimmed using

Cutadapt v1.7.1. Trimmed reads were aligned to both human hg38 and mouse mm39 reference genomes using STAR aligner v2.7.5b. XenofilteR v1.6 was used to select Human and mouse specific reads

and counts matrix were generated using HTSeq v0.11.2. EdgeR v3.28 was used for differential expression analysis. The normalized log-transformed counts per million (CPM) was used to calculate

single-sample gene set enrichment analysis score against specific MsigDB signature gene sets using R v4.2.0 package. Differentially expressed genes between treated vs untreated and its

significance (adjusted P-value < 0.05) were calculated using Wald’s T-test method implemented in the DESeq2 R package. SINGLE-CELL RNA TRANSCRIPTOME ANALYSIS (SCRNA-SEQ) SINGLE-CELL

RNA-SEQUENCING scRNA-Seq for dissociated PDXs was performed using the 10X Genomics Chromium Single Cell 3′ Library & Gel bead Kit V3.0, according to the manufacturers protocol (CG000183

Rev C). Briefly, ~5,000 PDX cells were used as input per sample. Cell encapsulation in microfluidic droplets yielded ~4000 recovered single-cell transcriptomes per sample. After reverse

transcription, barcoded-cDNA was purified using SILANE Dynabeads followed by 11 cycles of PCR-amplification. SPRIselect purification was performed on an Agilent Bioanalyzer High Sensitivity

chip to quantitate the fragment size and concentration of the amplified cDNA. Libraries were sequenced on an Illumina NovaSeq6000 with 151 bp paired-end reads. SINGLE-CELL RNA-SEQ ANALYSIS

XenoCell v1.0 was used to align transcripts to the GRCh38 human reference genome and mm10 mouse genome51, mouse-specific cellular barcodes were selected containing a minimum of 90% of

host-specific reads. Extracted mouse cells were then processed using Alevin tool (Salmon Software v1.3.0) to obtain unique molecular identifiers (UMIs; Supplementary Data 8) and generate a

cell by gene count matrix52, which was imported into Seurat (v3.2.0) for downstream analysis53. We excluded outlier cells that expressed in the range of <200–800 genes depending on the

sample type and had unusual gene count, transcript count, and mitochondrial gene fraction, according to sample-specific thresholds (Supplementary Data 8). We also excluded genes expressed in

fewer than 50 cells. Epithelial tumor cells (vehicle sample) identified were further down sampled to 100 cells, similar to the amount of carboplatin sample for all downstream analysis. For

the carboplatin-treated sample, we subsampled to 500 cells, similar to the amount of the vehicle sample. The SCTransform function from Seurat was then used to log-normalize and scale counts

to 10,000 transcripts per cell and detect highly variable features. Principal component analysis (PCA) was performed using the top 3000 most highly variable features and used to cluster

cells by Uniform Manifold Approximation and Projection (UMAP). Cell clusters were identified using Seurat FindClusters function. The ClusterTree (v0.4.3) R package was used to determine the

optimal resolution and number of clusters for each sample54. Expression of selected genes signatures were calculated per-cell using the AddModuleScore function from Seurat. Differentially

expressed markers were identified using Seurat FindMarkers function and significant genes were selected based on the FDR < 0.05. Gene set enrichment analysis was performed using escape

(v1.2.0) enrichIT function in R (v4.1.0)55. Cell type annotation was performed by a differential gene expression analysis with a ROC test using FindAllMarkers function. Markers were ranked

based on Log2Fold change and the ROC power score. Cross reference of the markers and literature was done to annotate each cell type25. Previously identified gene expression markers were used

to define the phenotypes of cells present in the PDX graft. These signature genes were discerned by cross-examining multiple primary research papers detailing markers of M1/M2

polarization26,27,28, the distinctions between an iCAF and myCAF phenotype25,29, and specialization of the endothelium30. STATISTICAL ANALYSIS Linear mixed model analyses were conducted

using SPSS Statistics (Version 27; International Business Machines Corporation, Armonk, New York, United States of America). All other statistical analyses were performed in Prism v9

software (GraphPad). A _p_-value ≤ 0.05 was considered statically significant. Unless otherwise state, data analyses were conducted using unpaired Student’s _t_ test to compare two data sets

or using one-way/two-way ANOVA when analyzing multiple sets of data. Data were presented as mean ± standard error of the mean (SEM). Statistically significant (FDR < 0.05) gene sets were

calculated using escape package (v1.2.0) in R (v4.1.0). REPORTING SUMMARY Further information on research design is available in the Nature Portfolio Reporting Summary linked to this

article. DATA AVAILABILITY The bulk RNA-sequencing, and single-cell RNA-sequencing data that support the findings of this study have been deposited in the NCBI dbGaP repository under

accession number: phs003369.v1.p1. Genomic alterations in PDXs, determined by targeted DNA sequencing, was previously published. The curated set of genomic alterations was based on data

downloaded from the cBioPortal resource Prostate Adenocarcinoma [https://www.cbioportal.org/study/summary?id=prad_p1000 and https://www.cbioportal.org/study/summary?id=prad_su2c_2015]. The

remaining data are available within the Article or its Supplementary Information. Source data are available as Source Data File. To request access to MURAL PDXs and/or biospecimens,

researchers should contact Dr. Melissa Papargiris, MURAL Project Manager ([email protected]) to initiate an Expression of Interest. Researchers would need to provide evidence of

institutional approval to experiment with human PDX tumors and research would be conducted under the conditions of a Materials Transfer Agreement. Source data are provided with this paper.

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members of the Prostate Cancer Research program, the patients, families, and consumers who support our research, and the members of the Melbourne Urological Research Alliance (MURAL). We

wish to thank the Prostate Cancer Biorepository Network (PCBN) for access to the tissue microarray, whose work is supported by the Department of Defense Prostate Cancer Research Program, DOD

Award No W81XWH-18-2-0013, W81XWH-18-2-0015, W81XWH-18-2-0016, W81XWH-18-2-0017, W81XWH-18-2-0018, and W81XWH-18-2-0019 PCRP PCBN. We acknowledge the association with the Bristol-Myers

Squibb (BMS) study #CA209-7TK and thank BMS for providing access to the LeY CAR T cell construct. We also thank Andrew Scott (Oliver Newton-John Cancer Research Institute) for access to the

anti-LeY antibody. This work was supported by Movember and MRFF (Upfront PSMA Prostate Cancer Research Alliance). Fellowship and grant support was obtained from the National Health and

Medical Research Council (GPR, APP1102752: PKD, APP1136680; JAT, APP1102752; RAT, LHP, Ideas grant, APP2011391), Cancer Council Victoria (JAT, LHP, TP834128), Department of Health and Human

Services acting through the Victorian Cancer Agency (RAT; MCRF15023), Prostate Cancer Foundation of Australia (LHP, PIRA YI-0322); and Monash University (LHP, Bridging Fellowship; LHP,

Platform Access Grant PAG2-9067770177). Infrastructure support was obtained from the EJ Whitten Foundation, Movember Foundation (Global Action Plan 1), the Peter and Lyndy White Foundation

and TissuPath Pathology. This research was supported by the Monash University Histology Platform, Monash University Animal Research Laboratories, Monash Biomedicine Discovery Institute

Organoid Program. This research was funded by a Prostate Cancer Research Alliance (PCRA) funded by Movember and the Medical Research Future Fund (MRFF). ProsTIC is additionally supported by

the Prostate Cancer Foundation (PCF). AUTHOR INFORMATION Author notes * These authors contributed equally: L. H. Porter, J. J. Zhu. * These authors jointly supervised this work: R. A.

Taylor, G. P. Risbridger. AUTHORS AND AFFILIATIONS * Prostate Cancer Research Group, Monash Biomedicine Discovery Institute, Cancer Program, Department of Anatomy and Developmental Biology,

Monash University, Clayton, VIC, 3800, Australia L. H. Porter, N. L. Lister & G. P. Risbridger * Cancer Immunology Program, Cancer Research Division, Peter MacCallum Cancer Centre,

Melbourne, VIC, 3000, Australia J. J. Zhu, P. J. Neeson, P. K. Darcy, J. A. Trapani, R. A. Taylor & G. P. Risbridger * Sir Peter MacCallum Department of Oncology, The University of

Melbourne, Parkville, VIC, 3010, Australia J. J. Zhu, S. Keerthikumar, D. L. Goode, R. Quezada Urban, A. Azad, M. S. Hofman, P. J. Neeson, P. K. Darcy, J. A. Trapani, R. A. Taylor & G.

P. Risbridger * Prostate Cancer Research Group, Monash Biomedicine Discovery Institute, Cancer Program, Department of Physiology, Monash University, Clayton, VIC, 3800, Australia S. G.

Harrison & R. A. Taylor * Cancer Research Division, Peter MacCallum Cancer Centre, Melbourne, VIC, 3000, Australia S. Keerthikumar, D. L. Goode & R. Quezada Urban * Computational

Cancer Biology Program, Peter MacCallum Cancer Centre, Melbourne, VIC, 3000, Australia S. Keerthikumar, D. L. Goode & R. Quezada Urban * Department of Pathology, Peter MacCallum Cancer

Centre, Melbourne, Victoria, Australia D. J. Byrne * Department of Medical Oncology, Peter MacCallum Cancer Centre, Melbourne, VIC, 3000, Australia A. Azad * Queensland Bladder Cancer

Initiative, School of Biomedical Science, Faculty of Health, Queensland University of Technology, Brisbane, QLD, 4102, Australia I. Vela * Australian Prostate Cancer Research Center, School

of Biomedical Science, Faculty of Health, Queensland University of Technology, Brisbane, QLD, 4102, Australia I. Vela * Department of Urology, Princess Alexandra Hospital, Brisbane, QLD,

4102, Australia I. Vela * Molecular Imaging and Therapeutic Nuclear Medicine, Peter MacCallum Cancer Centre, Melbourne, VIC, 3000, Australia M. S. Hofman * Prostate Cancer Theranostics and

Imaging Centre of Excellence (ProsTIC), Peter MacCallum Cancer Centre, Melbourne, VIC, 3000, Australia M. S. Hofman, R. A. Taylor & G. P. Risbridger Authors * L. H. Porter View author

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You can also search for this author inPubMed Google Scholar CONTRIBUTIONS G.P.R. and R.A.T. had full access to all the data in the study and takes responsibility for the integrity of the

data and the accuracy of the data analysis. Concept and design: G.P.R., R.A.T., J.A.T., P.K.D., and P.J.N. Acquisition of data: L.H.P., J.J.Z., N.L.L., S.G.H., S.K., D.L.G., R.Q.U., and

D.J.B. Analysis and interpretation of data: L.H.P., J.J.Z., N.L.L., S.G.H., S.K., D.L.G., R.Q.U., D.J.B., G.P.R., R.A.T., J.A.T., P.K.D., and P.J.N. Drafting of the manuscript: L.H.P.,

J.J.Z., N.L.L., S.G.H., S.K., R.Q.U., G.P.R., R.A.T., J.A.T., P.K.D., and P.J.N. Critical revision of the manuscript for important intellectual content: L.H.P., J.J.Z., N.L.L., S.G.H.,

D.J.B., A.A., I.V., M.S.H, P.J.N., P.K.D., J.A.T., R.A.T., and G.P.R. Statistical analysis: L.H.P., J.J.Z., N.L.L., S.G.H., S.K., and R.Q.U. Obtaining funding: G.P.R., R.A.T., J.A.T., and

L.H.P. Supervision: G.P.R., R.A.T., J.A.T., P.K.D., and P.J.N. Other: none. CORRESPONDING AUTHORS Correspondence to R. A. Taylor or G. P. Risbridger. ETHICS DECLARATIONS COMPETING INTERESTS

G.P.R. and R.A.T. (Research collaborations: Pfizer, Astellas, Zenith Epigenetics, AstraZeneca); P.K.D. (Research funding from Myeloid Therapeutics, Prescient Therapeutics, and Bristol Myers

Squibb); A.A. (Speakers Bureau: Astellas, Janssen, Novartis, Amgen, Ipsen, Bristol Myers Squibb; Merck Serono, Bayer; Honoraria: Astellas, Novartis, Sanofi, AstraZeneca, Tolmar, Telix, Merck

Serono, Janssen, Bristol Myers Squibb, Ipsen, Bayer, Pfizer, Amgen, Noxopharm, Merck Sharpe Dome; Scientific Advisory Board: Astellas, Novartis, Sanofi, AstraZeneca, Tolmar, Pfizer, Telix,

Merck Serono, Janssen, Bristol Myers Squibb, Ipsen, Bayer, Merck Sharpe Dome, Amgen, Noxopharm; Travel + Accommodation: Astellas, Merck Serono, Amgen, Novartis, Janssen, Tolmar, Pfizer;

Investigator Research Funding: Astellas, Merck Serono, AstraZeneca; Institutional Research Funding: Bristol Myers Squibb, AstraZeneca, Aptevo Therapeutics, Glaxo Smith Kline, Pfizer,

MedImmune, Astellas, SYNthorx, Bionomics, Sanofi Aventis, Novartis, Ipsen); M.S.H. (personal fees for lectures or advisory boards: Jannsen, Mundipharma, Astellas, Merck/MSD, Astra Zeneca,

Point Biopharma; Research support paid to institution: Endocyte and Advanced Accelerator Applications, both Novartis companies); I.V. (Honoraria: Astellas, Abbvie, Tolmar, Janssen;

Scientific Advisory Board: Astellas, AstraZeneca, Janssen, Bayer); P.J.N. (Research funding from BMS, Roche Genentech, MSD, Prescient Therapeutics, CRISPR Therapeutics, Allergan, Compugen);

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J.J., Lister, N.L. _et al._ Low-dose carboplatin modifies the tumor microenvironment to augment CAR T cell efficacy in human prostate cancer models. _Nat Commun_ 14, 5346 (2023).

https://doi.org/10.1038/s41467-023-40852-3 Download citation * Received: 06 September 2022 * Accepted: 11 August 2023 * Published: 02 September 2023 * DOI:

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