Solid tumour CAR-T cells engineered with fusion proteins targeting PD-L1 for localized IL-12 delivery

Murad, John P.; Christian, Lea; Rosa, Reginaldo; Ren, Yuwei; Buckley, Alyssa J.; Lee, Eric Hee Jun; Lopez, Lupita S.; Park, Anthony K.; Yang, Jason; Yamaguchi, Yukiko; Trac, Candi; Adkins, Lauren N.; Chang, Wen-Chung; Martinez, Catalina; June, Carl H.; Forman, Stephen J.; Ishihara, Jun; Lee, John K.; Stern, Lawrence A.; Priceman, Saul J.

doi:10.1038/s41551-025-01509-2

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Published: 01 October 2025

Solid tumour CAR-T cells engineered with fusion proteins targeting PD-L1 for localized IL-12 delivery

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Abstract

Chimeric antigen receptor (CAR)-T cell efficacy in solid tumours is limited due in part to the immunosuppressive tumour microenvironment (TME). To improve antitumour responses, we hypothesized that enabling CAR-T cells to secrete bifunctional fusion proteins consisting of a cytokine modifier such as TGFβ^trap, IL-15 or IL-12, combined with an immune checkpoint inhibitor such as αPD-L1, would provide tumour-localized immunomodulation to improve CAR-T cell functionality. Here we engineer CAR-T cells to secrete TGFβ^trap, IL-15 or IL-12 molecules fused to αPD-L1 scFv and assess in vitro functionality and in vivo safety and efficacy in prostate and ovarian cancer models. CAR-T cells engineered with αPD-L1–IL-12 are superior in safety and efficacy compared with CAR-T cells alone and those engineered with αPD-L1 fused with TGFβ^trap or IL-15. Further, αPD-L1–IL-12 engineered CAR-T cells improve T cell trafficking and tumour infiltration, and localize IFNγ production, TME modulation and antitumour responses, with reduced systemic inflammation-associated toxicities. We believe our αPD-L1–IL-12 engineering strategy presents an opportunity to improve CAR-T cell clinical efficacy and safety across multiple solid tumour types.

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Main

Chimeric antigen receptor (CAR)-T cell therapy for solid tumours has been limited by the suppressive solid tumour microenvironment (TME), which results in poor trafficking of T cells to tumours, T cell exhaustion, inadequate T cell persistence and restricted engagement of endogenous antitumour immune responses^1,2,3,4,5. To address these therapeutic challenges, combination strategies with immune checkpoint inhibitors (ICIs; for example, αPD-1, αCTLA-4 and αPD-L1) and use of immunomodulatory factors (for example, IL-2, IL-7, IL-12, IL-15 and αTGFβ) have been investigated. ICIs are successful in treating certain malignancies including melanoma, kidney cancers and non-small-cell lung cancers^6,7,8,9,10. However, immunologically ‘cold’ cancers, such as metastatic castration-resistant prostate cancer, ovarian cancer and pancreatic cancer, have shown limited responses to ICIs. Delivery issues, poor biodistribution and systemic toxicities further complicate the use of ICIs and their combinations in many disease settings. Further, combining CAR-T cell therapies with ICIs, which is an obvious combinatorial option, has not produced consistently desirable outcomes^11,12,13. Interleukins play a critical role in both innate and adaptive immune responses, enhancing T and natural killer (NK) cell activity and improving antitumour responses in preclinical models^14,15,16,17. However, clinical applications of many cytokines including IL-12 and IL-15 have been limited due to tumour non-specific activities, raising concerns with cytokine release syndrome (CRS) and other unwanted systemic toxicities^{18,19,20,21,22,23}. These results highlight the need for unique tumour-restricted approaches to combine these therapies to improve outcomes with solid tumour immunotherapies.

Research by our group and others have developed CAR-T cells with integrated ICI and cytokine support for tumour-restricted activity^{24,25,26,27,28,29}. Our group recently engineered CAR-T cells with a membrane-bound form of IL-12 cytokine (mbIL-12), increasing local tumour IFNγ and enabling cooperation between cytokine modulation and TME changes²⁹. However, restricted expression of cytokines by adoptively transferred cells may lead to less pronounced TME modifications than with soluble or secreted IL-12. Similar studies have been applied to IL-15 and other cytokines^{16,26,27,30,31,32,33,34,35,36,37,38,39}. Approaches such as T cell activation-dependent cytokine secretion or anchoring cytokines to immune-related or extracellular matrix proteins showed promise but still may be subject to systemic toxicity concerns or would require exogenous intratumoural delivery^14,18,40,41. We reasoned that tumour-targeting CAR-T cells engineered to secrete bifunctional αPD-L1–cytokine fusions would enhance IFNγ induction, subsequently increasing immune checkpoint ligands, such as PD-L1, and provide an intratumoural ligand for further sequestering cytokine to the local TME to improve the therapeutic index for antitumour efficacy.

In the current study, we designed and evaluated CAR-T cells that are engineered to secrete bifunctional fusion proteins. We assessed safety and efficacy of this approach using multiple solid tumour CAR-T cells engineered with bifunctional proteins comprising an αPD-L1 blocking antibody fused with stimulatory molecules (IL-12, IL-15/Ra or TGFβ^trap). These armoured CAR-T cells exhibit functional PD-L1 binding, improved antitumour activity and antigen-specific expansion in tumour co-culture assays. In immunologically ‘cold’ syngeneic prostate and ovarian cancer mouse models^42,43,44, mice receiving CAR-T cells engineered with αPD-L1–IL-12 fusions show superior antitumour activity compared with αPD-L1–IL-15 and αPD-L1–TGFβ^trap. Further, we demonstrated that a combination of monotherapies, or CAR-T cells secreting non-PD-L1 binding controls (αPD-L1^mut–IL-12) elicit greater systemic IFNγ and systemic inflammation while failing to achieve durable antitumour responses, as compared to CAR-T cells secreting αPD-L1–IL-12 fusions. In contrast, PD-L1 ligation safely facilitated IL-12-mediated increases in intratumoural IFNγ, increased CAR-T cell presence within the TME and remodelling of the suppressive TME. We also generated a human version of the αPD-L1–IL-12 fusion, which showed binding to PD-L1 and presentation of IL-12, along with functional benefits to engineered human CAR-T cells. We believe that our CAR-T cell engineering strategy to locally secrete and sequester αPD-L1–IL-12 fusion proteins within the TME has clinical application across a variety of solid tumour CAR-T-cell therapies to safely and dramatically improve therapeutic outcomes.

Results

CAR-T cells secrete bifunctional fusion proteins and exhibit PD-L1 binding and cytokine modulation in vitro

Murine splenic T cells were retrovirally transduced to express murine prostate stem cell antigen (PSCA)–CAR (previously described), CD19t⁴⁵ and bifunctional fusion proteins (Fig. 1a). T cell conditions include untransduced (UTD) T cells, PSCA–CAR only, PSCA–CAR/αPD-L1, PSCA–CAR/αPD-L1^mut–TGFβ^trap, PSCA–CAR/αPD-L1–TGFβ^trap, PSCA–CAR/αPD-L1^mut–IL-15, PSCA–CAR/αPD-L1–IL-15, PSCA–CAR/αPD-L1^mut–IL-12 or PSCA–CAR/αPD-L1–IL-12 (Supplementary Fig. 1a). All constructs showed comparable CAR-T cell transduction as measured by surface expression of mCD19t (Fig. 1b)^29,45, CD8/CD4 phenotypes (Supplementary Fig.1b) and bifunctional fusion proteins and relevant controls as measured by intracellular flow cytometry of the ΔCH2 modified Fc fragment in each fusion protein (Fig. 1c). Binding of each fusion protein and controls was assessed by an in vitro PD-L1 binding assay. The murine prostate cancer cell line, RM9, was induced to express PD-L1 via conditioned media obtained from tumour cell:CAR-T cell co-cultures. Following induction, supernatants from engineered CAR-T cells were incubated with tumour cells and assessed by flow cytometry for surface PD-L1 binding (Fig. 1d). Conditioned media resulted in significant induction of PD-L1 mean fluorescence intensity (MFI) (Fig. 1e). Supernatants from CAR-T cells engineered with PD-L1 fusion proteins, but not αPD-L1^mut proteins, significantly competed for binding, effectively blocking tumour surface PD-L1 and reducing its MFI in our flow cytometry assay (Fig. 1e). We also confirmed concurrent surface detection of bound TGFβRII, IL-15/Ra and IL-12(p40) via flow cytometry on PD-L1-induced tumours (Fig. 1f).

We next assessed differences in murine T cell functionality, including tumour cell killing, T cell expansion and activation, and cytokine production by CAR-T cells engineered with bifunctional fusion proteins in a repetitive tumour challenge assay in vitro. PSCA–CAR-T cells engineered with indicated fusion protein or controls were co-cultured with PSCA-positive murine prostate cancer cell line, PTEN–Kras hPSCA, at an effector:tumour (E:T) ratio of 1:2. Every 48 h, co-cultures were analysed by flow cytometry for tumour cell killing, cell phenotypes, as well as IFNγ secretion by ELISA. In addition, every 48 h, T cells were rechallenged with increasing numbers of PTEN–Kras hPSCA cells for a total of 4 tumour rechallenges (Fig. 1g). IL-15- and IL-12-containing fusions (αPD-L1 and αPD-L1^mut) sustained tumour cell killing at nearly 100% (relative to UTD) over each rechallenge as compared to PSCA–CAR only, PSCA–CAR/αPD-L1 or PSCA–CAR/αPD-L1–TGFβ^trap constructs (Fig. 1h). While IL-15 fusions induced the highest total T cell expansion (Fig. 1i) and CAR-T cell expansion at each tumour challenge (Supplementary Fig. 1c), IL-12 fusions induced an enrichment of CAR percentage over time (Fig. 1j). PSCA–CAR-T cells engineered with either αPD-L1^mut–IL-12 or αPD-L1–IL-12 outperformed the remaining constructs over each tumour challenge with higher 4-1BB MFI (Fig. 1k) and an increase in TIM-3 expression, which is a known downstream target of IL-12 signalling (Fig. 1l)⁴⁶. Total percentage of 4-1BB and other markers of exhaustion (PD-1, LAG3) were relatively unchanged (Supplementary Fig. 1d–f). While TGFβ^trap, IL-15 and IL-12 fusions induced higher initial secretion of IFNγ, only IL-12 fusions endowed a sustained increase in IFNγ secretion over the entire tumour challenge assay (Fig. 1m). The results indicated that CAR-T cells engineered to secrete cytokine modifiers IL-15 or IL-12 enhanced CAR-T cell function over CAR-T cells alone, but were relatively indistinguishable from their αPD-L1^mut counterparts, which suggested that more extensive in vivo assessment was required to determine whether simultaneous PD-L1 blockade with the IL-12 fusion protein conferred CAR-T cell non-autonomous benefits.

CAR-T cells engineered with αPD-L1–IL-12 fusions exhibit safe and durable antitumour responses in a syngeneic murine cancer model

We next assessed the therapeutic efficacy of CAR-T cells engineered with various bifunctional fusion proteins in vivo. hPSCA-KI mice (previously described)⁴⁵ were subcutaneously (s.c.) injected with PTEN–Kras hPSCA mouse prostate tumour cells, followed by treatment with Cy (100 mg kg⁻¹, intraperitoneal (i.p.)) and indicated CAR-T cells (1.0 × 10⁶, intravenous (i.v.)). Tumour growth was monitored by calipers, and peripheral blood was collected for flow cytometry and serum cytokine analysis (Fig. 2a). PSCA–CAR/αPD-L1–IL-12 T cells showed the greatest antitumour response in this model, which was 100% curative in mice treated relative to PSCA–CAR alone or other conditions. Interestingly, PSCA–CAR/αPD-L1–IL-12 T cells also outperformed PSCA–CAR/αPD-L1^mut–IL-12 T cells, which achieved a 50% curative response (Fig. 2b,c). PSCA–CAR-T cells engineered with TGFβ^trap and IL-15 fusions resulted in little to no therapeutic benefit over CAR-T cells alone. Importantly, while mice treated with PSCA–CAR/αPD-L1–IL-12 T cells showed no significant body weight changes, mice treated with either PSCA–CAR + soluble IL-12 (sIL-12) or PSCA–CAR/αPD-L1^mut–IL-12 T cells exhibited significant weight loss indicating systemic toxicity (Fig. 2d).

**Fig. 2: CAR-T cells engineered with αPD-L1–IL-12 fusion exhibit safe and durable antitumour responses in a syngeneic murine prostate cancer model.**

Mice treated with PSCA–CAR/αPD-L1^mut–IL-12 T cells had significantly higher systemic IFNγ levels relative to PSCA–CAR/αPD-L1–IL-12 (Fig. 2e). These findings suggest that PD-L1 binding in the TME resulted in reduced systemic IL-12 effects. Serum analysis of AST and ALT showed modest changes across treatment groups, but a trend of increased BUN, which may suggest mild impairment in kidney function in PSCA–CAR/αPD-L1^mut–IL-12 T cells and PSCA–CAR-T cells + sIL-12-treated mice (Supplementary Fig. 2a–c). Flow cytometric analysis of blood lymphoid populations showed that CAR-T cells engineered with IL-15 fusions resulted in significantly higher frequencies of circulating CD8 T cells and elevated 4-1BB expression (Fig. 2f,g and Supplementary Fig. 2d) but without improved antitumour response or survival. Mice treated with PSCA–CAR-T cells and sIL-12 or engineered with either αPD-L1^mut–IL-12 or αPD-L1–IL-12 showed a significant increase in peripheral blood CD4 T cells (Fig. 2h). For all indicated treatments, circulating T cells were not CAR positive (Supplementary Fig. 2e), as we have observed previously⁴⁵. For non-CAR CD8⁺ T cells in circulation, we saw minor increase in PD-1 expression in αPD-L1^mut–IL-12 and αPD-L1–IL-12 treatment, increases in TIM-3 expression for sIL-12, αPD-L1^mut–IL-12 and αPD-L1–IL-12, and slightly lower LAG3 expression relative to PSCA–CAR-T cell treatment alone (Supplementary Fig. 2f–h). Analysis of circulating myeloid cells in mice showed modest changes in total CD11b⁺ cells in all treatment groups (Fig. 2i). Mice treated with PSCA–CAR/αPD-L1–IL-12 T cells showed significant reduction in PD-L1 expression among the circulating Ly6C^−/lo monocytic myeloid cells (Fig. 2k and Supplementary Fig. 2i–k), which probably represent competitive fusion protein binding on tumour-experienced and -egressed cells^47,48. These results suggest that PSCA–CAR-T cells engineered with αPD-L1–IL-12 exhibit distinct local tumour delivery and subsequent binding to PD-L1-positive cells.

TAG72–CAR-T cells engineered with αPD-L1–IL-12 promote durable antitumour responses and exhibit tumour-restricted effects in a syngeneic murine ovarian cancer model

To expand our in vivo therapeutic efficacy studies, we evaluated TAG72–CAR-T cells engineered with bifunctional fusion proteins in our established ID8 ovarian cancer peritoneal metastasis model (Fig. 3a). Mice were i.p. injected with an aggressive, ex vivo passaged (Supplementary Fig. 3), TAG72-positive ID8 murine ovarian cancer cell line expressing firefly luciferase (ID8-mSTn/ffluc). At 14 days following injection, mice were treated i.p. with either engineered non-targeting (NT)-CAR/αPD-L1–IL-12 T cells, TAG72–CAR-T cells alone, or TAG72–CAR-T cells engineered with αPD-L1^mut–IL-12 or αPD-L1–IL-12. Non-invasive imaging of tumours showed that TAG72–CAR/αPD-L1–IL-12 had a dramatic and sustained antitumour response (Fig. 3b–d) resulting in 100% survival rate beyond 150 days post tumour injection (Fig. 3e). In contrast to αPD-L1–IL-12, mice treated with TAG72–CAR/αPD-L1^mut–IL-12 showed heterogeneous antitumour responses and reduced overall survival (Fig. 3e). Mouse weight changes were minimal among all treatment groups (Fig. 3f), probably due to the regional delivery of CAR-T cells compared with systemic delivery as well as the regional tumour development versus local disease in the prostate model (Fig. 2). To further validate the efficacy of αPD-L1–IL-12 bifunctional fusion, we compared this to αPD-L1–IL-15 and αPD-L1–TGFβ^trap using TAG72–CAR-T cells in the ovarian cancer peritoneal metastasis model (Supplementary Fig. 4a). TAG72–CAR/αPD-L1–IL-12 T cells again showed the greatest antitumour responses in this model (Supplementary Fig. 4b), which was curative in 50% of treated mice relative to other groups (Supplementary Fig. 4c). Interestingly, CAR-T cells engineered with αPD-L1–TGFβ^trap did not improve antitumour responses despite confirmed expression of TGFβ in relevant tumour cell lines (Supplementary Fig. 5). In contrast to our prostate cancer model, TAG72–CAR-T cells engineered with αPD-L1–IL-15 or αPD-L1^mut–IL-15 exhibited modest increases in antitumour responses relative to CAR-T cells alone, but not to the level observed with αPD-L1–IL-12. These data, using a second tumour model, confirm the benefits provided by engineering CAR-T cells with αPD-L1–IL-12 fusion.

**Fig. 3: TAG72–CAR-T cells engineered with αPD-L1–IL-12 promote durable antitumour responses and exhibit tumour-restricted effects in a syngeneic murine ovarian cancer model.**

Analysis of blood plasma collected from mice showed a trend of higher systemic inflammation with higher IFNγ in TAG72–CAR/αPD-L1^mut–IL-12 relative to TAG72–CAR/αPD-L1–IL-12 T cell treatment (Fig. 3g, left), similar to our in vivo prostate model observations. In a selection of mice collected on day 6 post treatment, solid tumour masses found within the upper omental region of the peritoneum were digested to single cells and magnetic bead isolated for CD45⁺ immune cells. Local IFNγ secretion from tumour-isolated CD45⁺ cell supernatants in ex vivo culture were significantly higher in mice treated with TAG72–CAR/αPD-L1–IL-12 T cells in contrast to TAG72–CAR/αPD-L1^mut–IL-12 or TAG72–CAR-T cells alone (Fig. 3g, right). Representative flow cytometry analysis of the same isolated CD45 cells showed similarly increased CD3⁺ T cells in TAG72–CAR/αPD-L1^mut–IL-12 and TAG72–CAR/αPD-L1–IL-12 groups (Fig. 3h), with the highest CAR-T cell percentages observed in the TAG72–CAR/αPD-L1–IL-12 group. Analysis of immune cell populations isolated from the peritoneal ascites from mice collected on day 12 post T cell treatment in mice treated at day 35 post tumour injection (Supplementary Fig. 6a) showed similar trends in IL-12-related increased T cell frequencies, but interestingly, unlike within the tumour itself, there was a non-significant presence of CAR⁺ T cells (Supplementary Fig. 6b–d). Within the ascites of TAG72–CAR/αPD-L1–IL-12 T cell treated mice, total myeloid (CD11b⁺) cells were relatively unchanged but exhibited decreased PD-L1 MFI (Supplementary Fig. 6e,f) as a result of fusion protein αPD-L1 competitive binding blockade. Further, a majority of these Ly6G⁻ Ly6C⁺ myeloid cells are F4/80^−/lo which also exhibit a decrease in PD-L1 expression relative to αPD-L1^mut–IL-12, and this trend is similarly found in F4/80^high subsets (Supplementary Fig. 6g–j). These results highlight sustained local tumour IFNγ production with αPD-L1–IL-12 engineered CAR-T cells, which in turn enhances tumour infiltration of CAR and non-CAR-T cells, contributing to the durable antitumour responses observed in this model.

CAR-T cells engineered with αPD-L1–IL-12 fusion elicit distinct changes in the TME

To assess TME changes following CAR-T cell treatment, we employed immunohistochemistry (IHC) and Nanostring GeoMx spatial proteomic analysis on tumours collected 7 days after T cell treatment in the syngeneic ovarian cancer model as previously described. Tumours from TAG72–CAR/αPD-L1^mut–IL-12 and TAG72–CAR/αPD-L1–IL-12 T cell groups showed increased intratumoural CD3, CD4 and CD8 T cells relative to TAG72–CAR-T cells alone, with low total frequencies of Foxp3⁺ regulatory T cells relative to total immune cells (Supplementary Fig. 7). Furthermore, tumour PD-L1 was markedly increased in αPD-L1^mut–IL-12 and αPD-L1–IL-12 groups as a likely consequence of increased local IFNγ. To further interrogate changes in the TME, Nanostring GeoMx spatial proteomic analysis was performed. A total of 12 tumour-specific regions of interest (ROI) from three replicate mice per group were selected (Fig. 4a and Supplementary Fig. 8). ROI values were z-score transformed from signal-to-noise ratio (SNR) adjusted counts of detected proteins. A heat map subdivided into major cell types, phenotype or status was then generated, and unsupervised clustering was applied to organize treatment groups. In mice treated with TAG72–CAR/αPD-L1–IL-12 and TAG72–CAR/αPD-L1^mut–IL-12 T cells, unbiased clustering tightly associated with treatment type (red treatment group and dark grey treatment ROIs, respectively), indicating consistent effects per treatment and per ROI (Fig. 4b).

In the TAG72–CAR/αPD-L1–IL-12 T cell treatment group, there was a marked enrichment of proteins related to lymphoid lineage, immune checkpoint and T cell activation status. This increase was less prominent in TAG72–CAR/αPD-L1^mut–IL-12 T cell, TAG72–CAR/αPD-L1 T cell or TAG72–CAR-T cell only groups (Fig. 4b). Moreover, TAG72–CAR/αPD-L1–IL-12 T cell treatment resulted in significantly greater intratumoural lymphoid cell and activation status via measures of CD3e, CD4, CD8a, CD28, PD-1, perforin, VISTA and Ki-67 in comparison to TAG72–CAR/αPD-L1^mut–IL-12 (Fig. 4c). All treatment groups had relatively similar counts in total CD11b; however, we found that TAG72–CAR/αPD-L1–IL-12 showed significantly reduced counts in myeloid CD14 positivity^49,50, but higher dendritic cells as measured by CD11c (Fig. 4d). Further, TAG72–CAR/αPD-L1–IL-12 T cell groups showed significantly lower counts of suppressive M2-like marker CD163 and significantly higher counts of MHC class II (Fig. 4d)^49,50,51. Of note, only the TAG72–CAR/αPD-L1–IL-12 T cell treatment group induced a significant increase in measured total PD-L1 protein (intracellular and extracellular) within the tumour ROIs selected, supporting the higher observed levels of local IFNγ. Lastly, total proliferative marker Ki-67 and cell apoptotic marker cleaved-caspase-3 were increased in TAG72–CAR/αPD-L1–IL-12 (Fig. 4e).

To gain a better understanding of co-expression patterns within tumour tissues for all treatment groups, a Spearman correlation analysis was performed. Statistically significant and strong positive or negative correlations were assessed and imply stronger intratumoural T cell activation responses in the TAG72–CAR/αPD-L1–IL-12 T cell group. We observed that CD8a T cell presence strongly correlated with both T cell co-stimulatory activation marker CD28 (Spearman R = 0.83, P = 0.0017) and T cell V-domain immunoglobulin suppressor of T cell activation (VISTA; R = 0.82, P = 0.002) relative to all other treatment groups (Fig. 4f,g). Furthermore, in the TAG72–CAR/αPD-L1–IL-12 T cell group, we observed a negative correlation between greater perforin expression (commonly associated with T cell killing) and lower M2-like suppressive myeloid CD163 expression (Fig. 4h; R = −0.7, P = 0.015)⁵¹. We also observed a strong correlation of MHC class II expression with CD11c in the TAG72–CAR/αPD-L1–IL-12 T cell group compared with the other groups (Fig. 4i). Together, these data support a cell-extrinsic mechanism of local TME modulation by CAR-T cells engineered with αPD-L1–IL-12 fusions, resulting in improved immune T cell function, shifts in myeloid cell status and other immune correlates.

Improved safety of CAR-T cells engineered with αPD-L1–IL-12 compared with drugging αPD-L1 antibody and IL-12 cytokine

Thus far, we have demonstrated the feasibility, safety and robust efficacy of engineering CAR-T cells with PD-L1–IL-12 fusion proteins against solid tumours. We have also demonstrated the therapeutic benefits of engineering CAR-T cells with αPD-L1–IL-12 versus αPD-L1^mut–IL-12, supporting our initial hypothesis that sequestering the cytokine in PD-L1⁺ tumours is optimal in reducing potential toxicities and enhancing therapeutic activity of IL-12. It is unclear, however, whether engineering αPD-L1 and IL-12 in CAR-T cells is beneficial over treatment with these modalities using biologics. To this end, we compared CAR-T cells engineered with αPD-L1–IL-12 to CAR-T cells in addition to αPD-L1 antibody and IL-12 cytokine injections in the syngeneic ovarian cancer model. To mitigate the significant body weight loss that we observed with IL-12 in our previous studies and to focus more heavily on the impact of systemic inflammation following therapy²⁹, we reduced the IL-12 dose when injected either alone or in combination with αPD-L1. Mice were treated (i.p.) with indicated NT-CAR or TAG72–CAR-T cells alone, or TAG72–CAR-T engineered with αPD-L1, αPD-L1^mut–IL-12 or αPD-L1–IL-12 fusions, or TAG72–CAR co-treated with either Avelumab (which is capable of binding both human and mouse PD-L1)⁵² and/or soluble IL-12 (sIL-12) (Fig. 5a). Tumours were monitored by non-invasive imaging for 7 days, showing a trend of the most potent antitumour activity with TAG72–CAR/αPD-L1–IL-12 T cells, followed by TAG72–CAR/αPD-L1^mut–IL-12 T cells and TAG72–CAR-T cells combined with Avelumab and sIL-12 (Fig. 5b). Interestingly, PD-L1 blockade alone, either engineered in CAR-T cells or CAR-T cells in combination with Avelumab, showed little to no benefit over CAR-T cells alone, but showed synergy in combination with IL-12.

Mice were subsequently euthanized on day 7 post treatment and assessed for systemic inflammatory responses. Mice treated with TAG72–CAR-T cells in combination with sIL-12, sIL-12 + Avelumab, and engineered TAG72–CAR/αPD-L1^mut–IL-12 had significant increases in spleen weight relative to TAG72–CAR/αPD-L1–IL-12 (Fig. 5c). Serum and peritoneal ascites fluid were assessed by ELISA and multiplex cytokine analysis. Treatment with TAG72–CAR/αPD-L1^mut–IL-12 T cells showed significantly higher levels of serum and ascites IFNγ relative to TAG72–CAR/αPD-L1–IL-12 (Fig. 5d,e) and correlated with IL-12 levels (Fig. 5f,g). Remarkably, while TAG72–CAR-T cells in combination with Avelumab and sIL-12 also showed elevated serum levels of IFNy and IL-12, these cytokines were dampened in the ascites fluid compared with TAG72–CAR/αPD-L1^mut–IL-12 T cell treatment. TAG72–CAR/αPD-L1^mut–IL-12 T cell treatment showed a marked increase in locoregional levels of multiple factors associated with chronic or acute inflammation (Fig. 5h), including IL-10, IL-6, CXCL10, and in triple treatment with sIL-12, an increase in MIP1a, all known to be associated with cytokine release syndrome (Fig. 5i–l). In contrast, TAG72–CAR-T cells engineered with αPD-L1–IL-12 fusion resulted in potent antitumour responses and reduced systemic and locoregional cytokine levels, thereby restricting systemic inflammatory responses compared with engineering with αPD-L1^mut–IL-12 or by drugging αPD-L1 and IL-12. These data further support our CAR-T cell engineering strategy for the safe and effective treatment of solid tumours.

Human CAR-T cells engineered to secrete human bifunctional αPD-L1–Fc–hIL-12 fusion protein exhibit improved in vitro functional activity

To expand on our translational efforts, we generated and tested human versions of our αPD-L1–Fc–hIL-12 and relevant controls (αPD-L1–Fc and hIL-12–Fc). To test secretion and PD-L1 binding efficacy, we collected supernatants from lentivirus-transduced Jurkat human T cells engineered with bifunctional fusion proteins (Fig. 6a). Briefly, PC3 human prostate tumour cells were induced to express PD-L1 via recombinant IFNγ overnight before being cultured with transduced Jurkat supernatants (Fig. 6b) from either untransduced (UTD), αPD-L1–Fc–hIL-12, αPD-L1–Fc or hIL-12–Fc lentiviruses. Human αPD-L1 containing fusion constructs significantly competed for binding, effectively blocking PC3 tumour surface PD-L1 and reducing MFI in our flow cytometry-based assay (Fig. 6c). We also confirmed appropriate surface detection of bound Fc and human IL-12 (hIL-12) via flow cytometry showing binding to PD-L1-induced PC3 cells (Fig. 6d,e).

We also assessed our αPD-L1–Fc–hIL-12 fusion approach in modifying human TAG72-targeting CAR-T cells engineered with bifunctional fusion proteins in in vitro co-culture assays. Using two independent lentiviral vectors (CAR and fusions), we single or dual-transduced human T cells to generate UTD, TAG72–CAR, TAG72–CAR/αPD-L1–Fc, TAG72–CAR/hIL-12–Fc or TAG72–CAR/αPD-L1–Fc–hIL-12. These cells were then co-cultured with the TAG72-positive human ovarian cancer cell line, PEO4-STn, at an E:T ratio of 1:2. After 48 h, co-cultures were analysed by flow cytometry to assess tumour cell killing, T cell expansion, activation and IFNγ cytokine production by ELISA. Dual transduction achieved ~40% TAG72–CAR expression and ~30% intracellular hIL-12 (hIL-12–Fc and αPD-L1–Fc–hIL-12) (Fig. 6f). Of note, detection of Fc was not used to detect αPD-L1–Fc in dual transduction with CAR, as the same Fc region is used in our human CAR-T cell constructs (Supplementary Fig. 9a,b), but detected as single transduced controls (Fig. 6g).

Functional analysis after a 48-h co-culture with TAG72-positive PEO4-STn cancer cells shows that hIL-12-containing constructs show moderate but significant increase in tumour cell killing (Fig. 6h). Total TAG72–CAR-T cell counts were higher in the TAG72–CAR/αPD-L1–Fc–hIL-12 condition relative to controls, and a significant increase in CAR+ T cell activation by CD25 (MFI) was also observed (Fig. 6i,j). Further, both TAG72–CAR/hIL-12–Fc and CAR/αPD-L1–Fc–hIL-12 engineered conditions resulted in significant increases in IFNγ (Fig. 6k). TAG72–CAR-T cells induced increases in tumour and T cell surface PD-L1, which were significantly reduced by culturing with TAG72–CAR/αPD-L1–Fc and CAR/αPD-L1–Fc–hIL-12 (Fig. 6l,m). While reduction in PD-L1 surface expression was modest in tumour cells compared with that in T cells, we reason that this may be due to the greater relative magnitude of PD-L1 induction (by MFI) on tumour cells in this system. We confirmed the presence of surface Fc and/or IL-12 on both tumour cells and T cells (Fig. 6n,p). In summary, we generated and functionally validated in vitro activity of a human version of αPD-L1–IL-12 fusion secreted by human CAR-T cells for further clinical translation.

Discussion

Our study addresses a substantial challenge in engineering safe and effective combinations of cytokine modification, immune checkpoint inhibition and CAR-T cell therapy for solid tumours. Our initial aim was to identify whether combining immune checkpoint inhibition of PD-L1 signalling with various immune modulating cytokines (IL-12, IL-15 or TGFβ^trap) is synergistic when engineered into solid tumour-targeting CAR-T cells. Using multiple in vivo models, we show that our CAR-T cells engineered to secrete αPD-L1–IL-12 fusion allow for sequestering IL-12 in PD-L1⁺ tumours, which improves safety, efficacy and TME landscape changes. Our study also highlights the synergistic antitumour activity of combining PD-L1 blockade and IL-12, compared with IL-15 or TGFβ^trap. Further, we observed an enhanced safety profile using our engineered αPD-L1–IL-12 fusion as compared with CAR-T cells combined with co-administration of exogenous PD-L1 blocking antibody and recombinant IL-12 cytokine.

Research exploiting combinations of ICI with cytokine modification as single agents or fusion molecules have been recently reported. As a therapeutic, αPD-L1–IL-15 fusions have shown mixed results in syngeneic mouse models in vivo^{15,27,31,32,53,54,55}, and suggest that perhaps other immune checkpoints such as αPD-1 or αCTLA-4 may work more favourably in concert with IL-15, specifically to aid in engaging T cell or NK cell effector function and immune memory⁵³. While we do see that αPD-L1–IL-15 fusion and the αPD-L1^mut–IL-15 control both elicited increased peripheral total T cell proliferation and higher peripheral CD8 T cells as well as 4-1BB+ T cells (Fig. 2 and Supplementary Fig. 2d) in our syngeneic prostate cancer model, these changes did not result in improved antitumour responses. While CAR-T cells secreting αPD-L1–IL-15 did show therapeutic benefit in our ovarian cancer model compared with CAR-T cells alone (Supplementary Fig. 4a–c), again CAR-T cells engineered with αPD-L1–IL-15 fusion did not reach the level of therapeutic activity observed in CAR-T cells engineered with αPD-L1–IL-12 fusion. With regards to TGFβ signalling inhibition, preclinical data also support engaging tumour PD-L1 in combination with TGFβ-‘trapping’ to alleviate immune suppression and boost immune and cell-based therapeutics^16,37,56,57. Clinically, this αPD-L1–TGFβ^trap approach so far has faced safety challenges in phase 3 trials^19,58. Our current data show that αPD-L1–TGFβ^trap provides moderate early increases in IFNγ (Fig. 1m) supportive of its function in vitro. However, we show no benefit of this fusion in our PTEN–Kras syngeneic prostate model (Fig. 2b,c) or our syngeneic ovarian cancer model (Supplementary Fig. 4a–c). While αPD-L1–IL-15 and αPD-L1–TGFβ^trap fusions performed suboptimally in our model systems, there may still be use for secreting these molecules from CAR-T cells in other models and warrants further investigation.

While effective in preclinical models as druggable agents, one major drawback of cytokine/ICI fusions is the undefined pharmacokinetics, biodistribution and requirement of repeated administration. This also raises concerns over tumour specificity despite attempting to anchor to checkpoints, or to non-immune tumour-specific targets including cell-surface vimentin, fibronectin or collagen binding domain^14,18,59. To improve tumour specificity, ex vivo loading of IL-12 anchored to CD45 expressed on antigen-specific T cells before infusion was recently described³³. Their results are most closely related to our approach in that anchoring IL-12 to tumour-targeting T cells can improve antitumour responses, improve safety and elicit modifications of the TME. Our approach uses a single dose of engineered CAR-T cells as a tumour-guided cellular delivery vehicle for αPD-L1–IL-12 fusions, enabling optimal local tumoural effects and potential for eliminating the need for repeated administration.

Our previous work established that CAR-T cells engineered with membrane-bound IL-12 (mbIL-12) stimulated greater IFNγ production and sequestered effects in vivo²⁹. While productive IFNγ signalling is crucial for effective tumour responses as well as MHC-I changes in the TME facilitated by CAR-T cells and IL-12 signalling, it can also induce compensatory PD-L1 expression^6,20. Recent findings suggest that improved immune function in immunologically ‘cold’ cancers, such as ovarian cancer, can be achieved when combining ICIs such as αPD-1 and αCTLA-4 with IL-12 (ref. ⁶⁰). These studies show that these combinations drive much greater amounts of IFNγ, which benefits immune function and tumour targeting via upregulation of MHC molecules. Our unique engineered fusion also showed enhanced IFNγ production, T cell–dendritic cell crosstalk and immune effector status, thereby presenting an opportunity to enhance the therapeutic index of ICIs in combination with CAR-T cells. Importantly, we believe that this IFNγ-driven increase in PD-L1 is not restricted to tumour cells, but may also apply to other immune cells within the TME including myeloid cells and T cells, which can also express PD-L1 resulting in greater immune suppression^61,62,63,64. Our data show that our combinatorial approach highlights the synergistic activity of IL-12 and PD-L1 blockade combined with CAR-T cell therapy, as evidenced by studies combining exogenous IL-12 and PD-L1 blockade as well as our engineered αPD-L1–IL-12 fusion. While we aimed to consolidate our treatment into a single administration, one could envision combining our mbIL-12-engineered CAR-T cells with PD-L1 blockade. Here we used a dual viral vector system to engineer the CAR and the fusion protein, which allows flexibility in manufacturing to control levels of each gene. This approach could be modified to single or bicistronic vectors if warranted^65,66.

In addition to safety and efficacy with systemic administration of CAR-T cells engineered with αPD-L1–IL-12 fusions, we showed use in both systemic and locoregional T cell delivery, suggesting applicability in other regional malignancies including brain and pleural cancers⁶⁷. In addition, we observed that regional intraperitoneal delivery of soluble IL-12 (with or without Avelumab) with CAR-T cells led to greater systemic IL-12 accumulation, while fusion-engineered CAR-T cells reduced systemic IL-12 leakage and lowered systemic inflammatory effects. To further improve tumour specificity of IL-12, alternative approaches may include structural attenuation of cytokines such as IL-12 (refs. ^17,68), localized proteasomal masked cytokines^21,34 or logic-gating strategies for regulatable activity outside the TME⁶⁹. While PD-L1 blockade in the presence of IL-12 promoted greater antitumour effects, we also observed a correlation of higher intratumoural CD8⁺ T cells with greater amounts of immune checkpoint VISTA in mice treated with TAG72–CAR/αPD-L1–IL-12 T cells. Our spatial data also showed higher levels of other immune checkpoints including TIM-3, LAG3 and PD-1, indicating additional potential resistance mechanisms to support combinations with traditional immune checkpoint blockade to offset T cell exhaustion and prolong therapeutic action of these engineered T cells.

Overall, our study highlights a clinically translatable platform for CAR-T cell engineering with localized delivery of immune modulatory αPD-L1–IL-12 bifunctional fusions, validated in both in vivo mouse and in vitro human systems. This approach has broad applicability to multiple solid tumours, and cellular delivery holds potential across not only CAR-T cells, but CAR-engineered NK cells, macrophages and non-conventional T cells, in disease settings that warrant immune checkpoint blockade and cytokine delivery to improve therapeutic responses^70,71,72.

Methods

Cell lines

The mouse prostate cancer cell line PTEN^−/− Kras (PTEN–Kras) derived from 10-week K-ras G12D;PTEN deletion mutant prostate cells was a kind gift from David Mulholland (The Icahn School of Medicine at Mount Sinai)⁴². The Ras/Myc transformed prostate cancer line, RM9, was a kind gift from Timothy C. Thompson (MD Anderson Cancer Centre) and has been previously described⁴⁵. ID8, a cell line originated from C57BL/6 mouse ovarian surface epithelial cells, was a kind gift from Karen Aboody (City of Hope National Medical Center). Cell lines were cultured in complete Dulbecco’s modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum (FBS, Hyclone), 2 mM l-glutamine (Fisher Scientific) and 25 mM HEPES (Irvine Scientific) (cDMEM). PC3 human prostate cancer cell line (ATCC, CRL-1435) and PEO4 human ovarian cancer cell line (Sigma Aldrich, 10032309) were cultured in RPMI-1640 medium (Lonza) containing 10% FBS (Hyclone), 2 mM l-glutamine (Fisher Scientific) and 25 mM HEPES (Irvine Scientific) (complete (c)RPMI). Virus titres were checked with either HT1080 (ATCC; CCL-121, cDMEM) and/or Jurkat cells (ATCC TIB-152; clone E6-1, cRPMI). Human embryonic kidney cell line 293T (ATCC; CRL-3216) was cultured in cDMEM. To generate a more aggressive ID8 tumour line in vivo, the parental ID8 cells were engrafted in vivo in the peritoneal cavity of mice for ~30 days. Unsorted cells collected from murine ovarian model peritoneal ascites were processed for red blood cell (RBC) lysis and analysed immediately as described. Passaged ID8 cells were then engrafted into tumour-naïve mice confirming increased aggressiveness relative to parental cells.

DNA constructs, lentiviral production and transduction, and retrovirus production

Tumour cells were engineered to express firefly luciferase (ffluc) by transduction with epHIV7 backbone lentivirus carrying the ffluc gene under the control of the EF1α promoter. PTEN–Kras cells were transduced to express human PSCA (hPSCA) (human PSCA, accession no. NM_005672.5), and ID8 and PEO4 cells were engineered to express tumour-associated glycoprotein-72 (TAG72) via transduction with epHIV7 lentivirus carrying the St6galnac-I gene (STn; human, accession no. NM_018414.5, or murine mSTn, accession no. NM_011371), under the control of the EF1α promoter. STn is the unique sialyltransferase responsible for generating surface expression of aberrant glycosylation sialyl-Tn (TAG72)⁷³. The single-chain variable fragment (scFv) sequence used in the TAG72–CAR construct was obtained from a murine monoclonal antibody clone CC49 that targets TAG72 (ref. ⁷⁴). The extracellular spacer domain included the 129-amino acid middle-length CH2-deleted version (ΔCH2) of the murine IgG1 Fc spacer and a murine CD28 transmembrane domain⁷⁴. The scFv sequence from the murine anti-hPSCA antibody (clone 1G8) was used to develop the murine PSCA–CAR construct. The extracellular spacer domain included the murine CD8 hinge region, followed by a murine CD8 transmembrane domain⁴⁵. Both TAG72–CAR and PSCA–CAR had a 4-1BB intracellular co-stimulatory signalling domain. The murine CD3ζ cytolytic domain was previously described^29,45. The CAR sequence was separated from a truncated murine CD19 gene (mCD19t) by a T2A ribosomal skip sequence and cloned in a pMYs retrovirus backbone under the control of a hybrid MMLV/MSCV promoter (Cell Biolabs) as previously described^29,45,74. Bifunctional fusion proteins or controls contain αPD-L1 scFv derived from Avelumab, or mutein non-PD-L1 binding variant (αPD-L1^mut) as described previously⁵⁵. Sequences from full-length murine IL-12(p40p35) (murine IL-12a and IL-12b, NCBI accession nos. NM_001159424.3, NM_001303244.1)⁶⁸, human IL-15/Ra (NCBI human IL15 and IL15RA, accession nos. NM_000585.5, NM_001243539)^30,55 and TGFβRII (US Patent US9676863B2)³⁰ were used to construct their respective fusions. Murine fusions contained linkers using a CH2-deleted version (ΔCH2) of human IgG1 Fc spacer (NCBI genomic DNA accession no. NG_001019.6) (Supplementary Fig. 1a). Human bifunctional fusion proteins and controls contain αPD-L1 scFv derived from Avelumab (which is cross reactive with human and mouse). Human fusions contained linkers using a CH2-deleted version (ΔCH2) of human IgG4 Fc spacer (NCBI accession no. NG_001019.6). Sequences from full-length human IL-12(p35p40) (NCBI accession no. IL-12A, NM_000882.4; IL-12B, NM_002187.3) were used, with the p40 subunit distal in all instances. All constructs were ordered as complete gene fragments (Genscript) and cloned into appropriate epHIV7 lentiviral or pMYs retroviral backbones as previously described^{29,45,74,75,76}. Bifunctional murine and human fusion and CAR construct illustrations are detailed in Supplementary Figs. 1 and 9. Amino acid sequences for murine CARs and bifunctional fusions (human and murine) are detailed in Supplementary Table 1, and human bifunctional fusions are also available in US Provisional Patent Application No. 63/772,399. Amino acid sequence for the human TAG72–CAR is detailed in US Patent US20210308184A1.

Retrovirus was produced by transfecting the ecotropic retroviral packaging cell line, PLAT-E (Cell Biolabs), with addition of murine PSCA–CAR and murine TAG72–CAR retrovirus backbone plasmid DNA using FuGENE HD transfection reagent (Promega). Viral supernatants were collected after 24, 36 and 48 h, pooled and stored at −80 °C in aliquots for future T cell transductions⁴⁵. Lentivirus was generated by plating 293T cells in T-225 tissue culture flasks 1 day before transfection with packaging plasmids and desired CAR lentiviral backbone plasmid. Supernatants were collected after 3–4 days, filtered and centrifuged to remove cell debris, and incubated with 2 mM magnesium and 25 U ml⁻¹ Benzonase endonuclease (EMD Millipore) to remove contaminating nucleic acids. Supernatants were combined and concentrated via high-speed centrifugation (6,080 × g) overnight at 4 °C. Lentiviral pellets were then resuspended in phosphate-buffered saline (PBS)-lactose solution (4 g lactose per 100 ml PBS), aliquoted and stored at −80 °C for later use. Lentiviral titres were determined by transducing Jurkat or HT1080 cells. Flow cytometry was used to quantify expression of either intracellular αFc (fusion constructs) or surface CD19t (CAR+), respectively.

Murine and human T cell isolation, transductions and ex vivo expansion

Murine T cell activation and transduction was performed as described previously^29,75. Briefly, for murine studies, splenocytes were obtained by manual digestion of spleens from male heterozygous hPSCA-KI or female C57BL/6j mice. Enrichment of T cells was performed using the EasySep mouse negative T cell isolation kit following manufacturer protocol (StemCell Technologies). Murine PSCA–CAR or murine TAG72–CAR were achieved with single retrovirus transductions. Dual transduction required mixing of desired murine CAR retrovirus and secondary helper/fusion protein retrovirus to generate CAR-expressing fusion-positive T cells. Subsequent murine T cell expansion was performed as previously described²⁹.

Human leukapheresis products were obtained from consenting healthy donors under City of Hope IRB-approved protocol no. 09025. Peripheral blood mononuclear cells (PBMCs) were isolated via Ficoll-Paque (GE Healthcare) density gradient centrifugation, washed in PBS/EDTA and cryopreserved in CryoStor (StemCell Technologies). T cell activation and transduction was performed in part as described previously^29,76. Briefly, thawed PBMCs (2.0 × 10⁶ CD3⁺) were cultured in X-VIVO medium (Lonza) containing 100 U ml⁻¹ recombinant human IL-2 (rhIL-2, BioTechne) and 0.5 ng ml⁻¹ recombinant human IL-15 (rhIL-15, BioTechne). T cells were activated via CD3/CD28 TransAct (Miltenyi Biotec, 130-111-160) at 1:500 dilution, following manufacturer-suggested protocol, in 24-well tissue culture plates (Corning). Primary human TAG72–CAR transduction was performed in the presence of protamine sulfate (APP Pharmaceuticals), cytokines and human TAG72–CAR-T cell lentivirus as described previously^29,76. Secondary transduction with human helper viruses (αPD-L1–Fc, αhIL-12–Fc or αPD-L1–Fc–hIL-12) involved restimulation with TransAct and cytokine replenishment 24 h following primary transduction. All viruses were used at an MOI of 1. Cells were continuously cultured with cytokine-enriched X-VIVO, refreshed every 2–3 days. At 10 days post transduction, CAR-T cell purity and phenotype were assessed by flow cytometry, detecting surface CAR (hCD3⁺hCD19⁺) and intracellular helper proteins via Fc-linker and hIL-12 expression. At day 14, cells were used for in vitro co-culture assays or cryopreserved in CryoStor CS5 for future use.

Flow cytometry

For flow cytometric analysis, cells were resuspended in FACS buffer (Hank’s balanced salt solution without Ca²⁺, Mg²⁺ or phenol red (HBSS^−/−), Life Technologies) containing 2% FBS and 1× antibiotic-antimycotic (GIBCO, FACS buffer). Single-cell suspensions from mouse tissues or tumours were incubated for 20 min on ice with mouse Fc Block at a dilution of 1:50 (BD, 553140). Cells were then incubated with primary antibodies for 30 min at 4 °C in the dark with either Brilliant Violet 510 (BV510), Brilliant Violet 570 (BV570), Brilliant Violet 605 (BV605), Brilliant Violet 650 (BV650), fluorescein isothiocyanate (FITC), phycoerythrin (PE), peridinin chlorophyll protein complex (PerCP), PerCP-Cy5.5, PE-Cy7, allophycocyanin (APC), or APC-Cy7 (or APC-eFluor780), eFluor506, PE/Dazzle 594, PerCP-eFluor 710, BD Horizon Red 718 (R718), Alexa Fluor 488 (AF488), or PE-Cy5-conjugated antibodies. Antibodies against mouse CD3 (BD Biosciences, 563109, Clone: 17A2), mouse CD4 (ThermoFisher, 340443, Clone: RM4-5), mouse CD8a (BioLegend, 347313, Clone: 53-6.7), mouse CD19 (BD Biosciences, 557835, Clone: 1D3), mouse CD45 (BioLegend, 103145, Clone: 30-F11), mouse CD137 (ThermoFisher, 25-1371-82, Clone: 17B5), mouse NK1.1 (BioLegend, 108733, Clone: PK163), mouse PD-1 (BioLegend, 69-9985-80, Clone: J43), mouse LAG3 (BioLegend, 125227, Clone: C9B7W), mouse TIM-3 (BioLegend, 119704, Clone: RMT3-23), mouse CD11b (BioLegend, 101237, Clone: M1/70), CD44 (BD Biosciences, 103010, Clone: IM7), CD62L (BioLegend, 104412, Clone: MEL-14), CD80 (BD Biosciences, 740130, Clone: 16-10A1), mouse I-A/I-E (MHC Class II) (Biolegend, 64-5321-80, Clone: M5/114.15.2), mouse CD274 (PD-L1) (BioLegend, 124312, Clone: 10F.9G2), Ly6-C (BioLegend, 128029, Clone: HK1.4), mouse CD11c (BioLegend, 117316, Clone: N418), mouse Ly-6G (Biolegend, 127623, Clone: 1A8), mouse CD103 (BioLegend, 121426, Clone: 2E7), mouse F4/80 (BioLegend, 123127, Clone: BM8), mouse IL-12/IL-23 p40 (ThermoFisher, 12-7123-41, Clone: 17.8), human IL-15/Ra (R&D Systems, FAB10900R, Clone: 2639B, 1:20 dilution), human TGFβ Receptor II (BioLegend, 399703, Clone: W17055E), purified non-conjugated anti-human TGFβ Receptor II antibody (BioLegend, 399702, Clone: W17055E), anti-human CD45 (BD Biosciences, 563204; Clone: HI30), anti-human CD25 (Invitrogen, 46-0259-42, Clone: BC96), anti-human CD19 (BD Pharmigen, 557835, Clone: SJ25C1), purified APC anti-human PD-L1 antibody (BD Biosciences, 563741, Clone: MIH1) and PE mouse anti-human IL-12 (p40/p70) (BD Bioscienes, 554575, Clone: C11.5). All antibodies (anti-mouse and anti-human) used for flow cytometery were used at a dilution of 1:100 unless stated otherwise. Cell viability was determined using 4′, 6-diamidino-2-phenylindole (DAPI, Sigma). When necessary, secondary staining of cells was performed by washing twice before a 30-min incubation at 4 °C in the dark. Flow cytometry was performed on a MACSQuant Analyzer 16 (Miltenyi Biotec), and the data were analysed with FlowJo software (v.10.8). Representative flow cytometry gating strategy for lymphoid and myeloid populations can be found in Supplementary Fig. 10.

For intracellular flow cytometry, BD GolgiStop (51-2092KZ) was added to CAR-T cells for blocking intracellular protein transport and incubated for 3–4 h at 37 °C. Cells were transferred to a 96-well plate. Reagents and buffers for flow cytometry processing were pre-chilled on ice unless otherwise stated. Cells were washed with FACS buffer and then fixed in 1× BD Cytofix/Cytoperm (51-2090KZ) at 4 °C for 20 min. Following washing with 1× BD Perm/Wash buffer (51-2091KZ) twice, cells were stained with intracellular antibody: FITC polyclonal goat anti-human Fc (Jackson ImmunoResearch, 109-096-008) for 30 min at 4 °C. Data were acquired on a MACSQuant Analyzer 16 cytometer (Miltenyi) and analysed with FlowJo software (v.10.8).

In vitro PD-L1 binding, tumour killing and functional assays

For murine PD-L1 blocking/binding experiments, PD-L1 expression was first induced on RM9 cells for 4 h via conditioned media collected from CAR-T cell:tumour cell co-cultures. PD-L1-induced tumours were then co-cultured for 1 h at room temperature with supernatants collected from supernatants of dual-transduced CAR-T cells engineered to secrete bifunctional fusion proteins (Fig. 1d). Levels of PD-L1 blockade, from indicated CAR-T cell supernatants or relevant positive and negative controls, were measured in a flow cytometry-based competitive binding assay using a fluorescently conjugated competitively binding αPD-L1 antibody (BioLegend, Clone: 10F.9G2). Simultaneously, PD-L1-induced and fusion protein-blocked tumours were measured for detection of surface-bound fusion cytokines IL-15/Ra, IL-12 and TGFβRII. For TGFβRII detection, tumours were first blocked with cold unconjugated anti-TGFβRII antibody to remove tumour receptor background before adding CAR-T cell supernatants.

For human PD-L1 blocking/binding experiments, PD-L1 expression was first induced on PC3 prostate tumour cells overnight in 20 ng ml⁻¹ recombinant human IFNγ (BioLegend, 570202). PD-L1-induced PC3 tumour cells were then co-cultured for 1 h at room temperature with supernatants collected from supernatants of transduced human Jurkat T cell tumour cells engineered to secrete human bifunctional fusion proteins or controls (Supplementary Fig. 9a). Levels of PD-L1 blockade, from indicated CAR-T cell supernatants or relevant positive and negative controls, were measured in a flow cytometry-based competitive binding assay using an APC fluorescently conjugated competitively binding anti-human PD-L1 antibody (BD Biosciences, 563741, Clone: MIH1; 1:100 dilution). Simultaneously, PD-L1-induced, αPLD1–Fc–IL-12 protein, single-sided controls or untransduced control Jurkat T cell supernatant blocked tumours, were measured for detection of surface-bound fusion cytokine human IL-12 (BD Bioscienes, Clone: C11.5; PE mouse anti-human IL-12 (p40/p70), 554575; 1:100 dilution) and/or Fc-linker protein using goat anti-human Fc-FITC (Jackson ImmunoResearch, 109-096-008; 1:100 dilution).

For mouse tumour cell killing assays, UTD or PSCA–CAR mouse T cells engineered with or without αPD-L1, αPD-L1–TGFβ^trap, αPD-L1^mut–IL-15, αPD-L1–IL-15, αPD-L1^mut–IL-12 or αPD-L1–IL-12 were co-cultured at a primary 1:2 E:T ratio against PTEN–Kras hPSCA mouse prostate tumour cells in complete RPMI medium without cytokines in 96-well plates (Fig. 1g). At indicated timepoints (every 48 h) co-cultures were analysed by flow cytometry as described. At these timepoints, replicate plates were rechallenged with an additional 20,000 PTEN–Kras hPSCA tumour cells every 2 days for a total of 5 tumour challenges. For human tumour cell killing assays, UTD or TAG72–CAR-T cells engineered with or without human αPD-L1–Fc, hIL-12–Fc or αPD-L1–Fc–hIL-12 were co-cultured in triplicate at a primary 1:2 E:T ratio against PEO4-STn, human ovarian cancer cells transduced to express TAG72, in complete X-VIVO media without cytokines in 96-well plates (Fig. 1g). After 48 h, co-cultures were analysed by flow cytometry as described. For both murine and human co-culture assays, tumour cell killing by CAR-T cells was calculated by comparing CD45-negative DAPI-negative (viable) cell counts relative to targets co-cultured with UTD. In addition to flow cytometry measures of co-cultures, cell supernatants were collected from each timepoint to quantify IFNγ (human or mouse) by ELISA.

ELISA cytokine assays

Supernatants from tumour cell killing assays were collected at indicated times and frozen at −80 °C for future analysis. Supernatants from all timepoints were thawed and analysed for murine IFNγ using the murine IFNγ ELISA Ready-SET-GO! kit (mIFNγ; Invitrogen, 88-7314-88), or for human IFNγ using the human IFNγ ELISA kit (huIFNγ; Invitrogen, 88-7316-88) following manufacturer protocol. Mouse serum, plasma, peritoneal ascites or tumour CD45⁺ isolated cell supernatant cytokine levels of IFNγ and IL-12 were measured using murine IFNγ and IL-12 ELISA Ready-SET-GO! ELISA kits (IL-12; Invitrogen, BMS616), following manufacturer protocols. Plates were read at 450 nm using a Cytation3 imaging reader with Gen5 microplate software v.3.05 (BioTek). Mouse Luminex Discovery Assay kit (15-Plex; R&D Systems, LXSAMSM-15) was used to evaluate multiple analytes on collected peritoneal ascites from in vivo ovarian cancer models. Multiplex cytokine expression data were log₂ transformed and displayed in balloon plots created using the gglot2 v.3.5.1 R package.

RT–qPCR

Complementary DNA was prepared from 1 μg of total RNA (matched from RNA-seq analyses) using SuperScript IV reverse transcriptase (ThermoFisher). Quantitative PCR was performed in triplicate using SsoAdvanced Universal SYBR Green Supermix (BioRad). Data were analysed using the comparative threshold method, and gene expression was normalized to murine GAPDH expression on a CFX RT-PCR instrument and CFX Maestro software (BioRad, v.2.3). Murine forward (F) and reverse (R) gene primers: murine gapdh (F: GTCAAGCTCATTTCCTGGTATGACA, R: GTTGGGATAGGGCCTCTCTTG) and murine tgfβ (F: AGCTGCGCTTGCAGAGATTA, R: AGCCCTGTATTCCGTCTCCT) were used.

In vivo studies

All animal experiments were performed under protocols approved by the City of Hope Institutional Animal Care and Use Committee (protocol no. 21025). For all animal studies, mice were housed with 12 h light and 12 h dark cycle at a temperature range of 68–75 °F and humidity between 30–70%. For subcutaneous prostate tumour studies, PTEN–Kras hPSCA cells (1.0 × 10⁶) were prepared in a final volume of 100 μl HBSS^−/− and injected under the skin of the abdomen of 6–8-week-old male heterozygous hPSCA-KI C57BL/6J mice as previously described⁴⁵. Tumour growth was measured using calipers (length × width × height = mm³). For prostate tumour studies, mice were treated by i.p. administration with soluble murine recombinant IL-12 (sIL-12; 1 μg, PeproTech, 210-12) once daily for 5 days starting on the day of T cell treatment. For in vivo i.p. ovarian tumour studies, ID8-mStn (ffluc expressing) cells (5.0 × 10⁶) were prepared in a final volume of 400 μl HBSS^−/− and engrafted in >6-week-old female C57BL/6J (Jackson Laboratories) mice by i.p. injection. Tumour burden was measured via non-invasive bioluminescence imaging (LagoX, Spectral Imaging), and flux signals were analysed with Aura software (v.4.0, Spectral Imaging). Where indicated, mice were treated i.p. with 100 mg kg⁻¹ cyclophosphamide (Cy). Mice received i.v. treatment with indicated T cells (1.0 × 10⁶ CAR+ T cells) in 100 μl final volume 24 h after Cy pre-conditioning. For ovarian tumour studies, mice were i.p. treated at 14 (or 35) days post tumour engraftment as indicated, with CAR+ T cells (3.0–5.0 × 10⁶) in 400 μl final volume without Cy pre-conditioning. For indicated ovarian tumour studies, mice were co-treated i.p. with CAR-T cells and either soluble murine recombinant sIL-12 (0.5 μg i.p.) or with Avelumab (200 μg i.p.; anti-PD-L1, Bavencio, EMD Serono) every other day for a total of 3 doses starting on the day of T cell treatment. For all studies, mice were euthanized upon reaching s.c. tumour volumes exceeding 1,000 mm³ or i.p. tumours showing signs of distress such as a distended belly due to peritoneal ascites, laboured or difficulty breathing, apparent weight loss, impaired mobility or evidence of being moribund.

Peripheral blood was collected from isoflurane-anaesthetized mice by retro-orbital (r.o.) bleed through heparinized capillary tubes (Chase Scientific) into polystyrene tubes containing a heparin/PBS solution (1,000 units ml⁻¹, Sagent Pharmaceuticals). Total volume of each blood draw (~120 μl per mouse) was recorded. RBCs were lysed with 1× Red Cell Lysis buffer (Sigma) according to manufacturer protocol, and then washed, stained and analysed by flow cytometry as described above. When applicable, tumour, liver and spleen were collected from euthanized mice. Spleen weights were measured for analysis of splenomegaly. Serum from syngeneic prostate and ovarian cancer mouse studies was collected via r.o. bleed in non-heparinized capillary tubes as described above. Blood was kept at room temperature for 30 min, centrifuged at 6,000 × g for 10 min at 4 °C, aliquoted and frozen at −80 °C until used for serum cytokine ELISA or chemistry analyses. Serum chemistry analysis was performed by running samples on a VETSCAN VS2 Chemistry Analyzer (Zoetis), using the phenobarbital chemistry panel rotor (Zoetis) for BUN, ALT and AST quantification as described in the manufacturer protocol. At pre-determined timepoints or at moribund status, mice were euthanized, and tissues and/or peritoneal ascites were collected and processed for flow cytometry and immunohistochemistry. Peritoneal ascites was centrifuged at 336 × g for 10 min at 4 °C, aliquoted and frozen at −80 °C until used for cytokine ELISA or Mouse Luminex Discovery Assay multiplex analysis per manufacturer protocol (R&D Systems). For the study of immune cells collected from ovarian i.p. tumour masses, mice from each group were randomly selected and killed at day 6 or 7 post T cell injection. Peritoneal solid tumour masses were physically minced and enzymatically digested using a Miltenyi mouse tumour digestion kit, and CD45 positive selection was performed using magnetic mouse CD45 MicroBeads following manufacturer protocols (Miltenyi Biotec). Isolated CD45⁺ cells were analysed by flow cytometry, and individual replicate supernatants of overnight-cultured isolated CD45⁺ cells from each treatment group were measured for murine IFNγ secretion via ELISA as described above.

Immunohistochemistry and Nanostring GeoMx digital spatial profiling (DSP) analysis

Tumour tissues were fixed for up to 3 days in 4% paraformaldehyde (Boston BioProducts) and stored in 70% ethanol until further processing. Immunohistochemistry was performed by the solid tumour pathology core at City of Hope. Briefly, paraffin-embedded sections (10 μm) were stained with haematoxylin and eosin (H&E, Sigma Aldrich), CD3 (Ventana, Clone: SGV6), PD-L1 (Invitrogen, Clone: JJ08-95), CD4 (Abcam, Clone: EPR19514), CD8 (Cell Signaling, Clone: D4W22) and FOXP3 (Abcam, Clone: EPR22102-37). Images were obtained using the Nanozoomer 2.0HT digital slide scanner and the associated NDP.view2 software (Hamamatzu). For Nanostring GeoMx DSP, similarly prepared tissues and slides were sent for multiplex protein profiling with spatial context. The Nanostring GeoMx system-provided data centre software was used for generating the raw SNR data. Briefly, after quality control of the initial data, the segment and target data were filtered, followed by generation of SNR data for downstream analysis. SNR count protein expression data from 12 tumour ROIs per treatment group were processed and analysed using R v.4.4.0. The normality of data was assessed using the Shapiro–Wilk test. Protein expression values were z-score transformed and displayed in heat maps using the ComplexHeatmap R package (v.2.20.0)⁷⁷. ROIs from different treatment groups were clustered using complete-linkage hierarchical clustering on expression data of 39 proteins subgrouped by cell type or phenotype. The strength and direction of the relationship between the expression of defined pairs of proteins were evaluated using Spearman’s rank correlation method. Unless otherwise specified, a P-value threshold of P < 0.05 was used to determine statistical significance.

Statistical analysis

In this study, we evaluated CAR-T cells for the treatment of solid tumours using in vitro T-cell functional assays, as well as syngeneic tumour models in mice. We engineered CAR-T cells to secrete bifunctional fusion proteins and evaluated therapeutic efficacy in these model systems. All in vitro assays were performed with at least duplicate samples and were repeated in at least two independent experiments. In vivo studies were performed using 6–8-week-old C57BL/6 or C57BL/6 background huPSCA-KI transgenic mice, using at least three mice per group for all in vivo studies to ensure statistical power. Mice were randomized on the basis of tumour volume or bioluminescence imaging to ensure evenly distributed average tumour sizes across each group. In vivo experiments were repeated at least twice. For subcutaneous tumour models, survival was based on the maximum tumour size allowed (~1,000 mm³ maximum volume). Excel (Microsoft, v.16.97) and GraphPad Prism 10 (GraphPad Software) were used to generate bar plots and graphs. Data are presented as mean ± s.e.m., unless otherwise stated. Unless otherwise indicated, P values for pairwise comparisons were generated using an unpaired two-tailed Student’s t-test with assumption of unequal variance, where *P < 0.05, **P < 0.01, ***P < 0.001 and ****P < 0.0001.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The main data supporting the results in the study are available within the paper and its Supplementary Information. Source data are provided with this paper.

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Acknowledgements

We thank the staff of the following cores at the Keck School of Medicine of the University of Southern California (USC) and the Norris Comprehensive Cancer Center: Animal Facility, and the Beckman Research Institute at City of Hope (COH) Comprehensive Cancer Center: Animal Facility, Pathology, and Small Animal Imaging for excellent technical assistance. Research reported in this publication was supported by a Prostate Cancer Foundation Tactical Award (2022TACT3835, to S.J.P. and C.H.J.), a Department of Defense Idea Expansion Award (W81XWH-2110354, to S.J.P.), the Mike and Linda Fiterman Foundation fund (to S.J.P.), the Camiel Family Foundation fund (to S.J.P.), and the Norman and Mary Pattiz Foundation fund (to S.J.P.). Work performed was supported by the National Cancer Institute of the National Institutes of Health under grant number P30CA014089 (USC) and P30CA033572 (COH). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

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Keck School of Medicine (KSOM)/Norris Center for Cancer Cellular Immunotherapy Research (CCCIR), Division of Medical Oncology, Department of Medicine, Keck School of Medicine of USC, Los Angeles, CA, USA
John P. Murad, Reginaldo Rosa, Yuwei Ren, Alyssa J. Buckley, Lupita S. Lopez, Yukiko Yamaguchi, Lauren N. Adkins, Catalina Martinez & Saul J. Priceman
Department of Hematology and Hematopoietic Cell Transplantation, City of Hope, Duarte, CA, USA
John P. Murad, Lea Christian, Anthony K. Park, Jason Yang, Candi Trac, Wen-Chung Chang, Stephen J. Forman & Saul J. Priceman
Wallace H. Coulter Department of Biomedical Engineering, Georgia Tech College of Engineering and Emory School of Medicine, Atlanta, GA, USA
Eric Hee Jun Lee
Center for Cellular Immunotherapies, Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, USA
Carl H. June
Department of Bioengineering, Imperial College London, London, UK
Jun Ishihara
Division of Hematology/Oncology, Department of Medicine, David Geffen School of Medicine at the University of California Los Angeles, Los Angeles, CA, USA
John K. Lee
Department of Chemical, Biological, and Materials Engineering, University of South Florida, Tampa, FL, USA
Lawrence A. Stern

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Contributions

S.J.P., along with J.P.M., conceived and designed the study. S.J.P., J.P.M., L.C., R.R., Y.R., A.J.B., E.H.J.L, L.S.L., A.K.P., J.Y., Y.Y., C.T., L.N.A., W.-C.C., J.I., J.K.L. and L.A.S. designed the experimental procedures, data analysis, and/or interpretation. J.P.M., L.C., R.R., Y.R., A.J.B., E.H.J.L, L.S.L., A.K.P., J.Y., Y.Y. and C.T. performed experiments. S.J.P., J.P.M. and L.C. wrote the manuscript. C.M., C.H.J., S.J.F., J.I. and J.K.L. assisted in data interpretation and writing/editing the manuscript. S.J.P. supervised the study. All authors reviewed the manuscript.

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Correspondence to Saul J. Priceman.

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Competing interests

S.J.P. is a scientific advisor and/or receives royalties from Imugene Ltd, Adicet Bio, Port Therapeutics, and Celularity. S.J.P., J.P.M., E.H.J.L. and S.J.F. are listed as co-inventors on a patent on the development of TAG72-targeted CAR-modified T cells for the treatment of TAG72-positive tumours, and S.J.P. and S.J.F. are listed as co-inventors on a patent on the development of PSCA-targeted CAR-modified T cells for the treatment of PSCA-positive tumours, owned by COH. S.J.P. and J.P.M. are listed as co-inventors on a patent on the development of PD-L1-targeted IL-12 fusion proteins, co-owned by USC and COH. The other authors declare no competing interests.

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Murad, J.P., Christian, L., Rosa, R. et al. Solid tumour CAR-T cells engineered with fusion proteins targeting PD-L1 for localized IL-12 delivery. Nat. Biomed. Eng (2025). https://doi.org/10.1038/s41551-025-01509-2

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Received: 27 September 2024
Accepted: 13 August 2025
Published: 01 October 2025
Version of record: 01 October 2025
DOI: https://doi.org/10.1038/s41551-025-01509-2