CISH, a key intracellular checkpoint, in comparison and combination to existing and emerging cancer immune checkpoints

Abstract

Over the past decade, Immuno-Oncology has largely focused on blocking inhibitory surface receptors like PD-1 to enhance T cell anti-tumor activity. However, intracellular immune checkpoints such as CISH, which function independently of tumor-expressed ligands, offer powerful and previously untapped therapeutic potential. As a downstream regulator of TCR signaling, CISH controls T cell activation, expansion, and neoantigen reactivity. Though historically considered undruggable, recent advances in CRISPR engineering have enabled functional interrogation of these targets. We demonstrate that CISH deletion enhances T cell activation and anti-cancer functions more effectively than other emerging intracellular checkpoints. In CAR-T cells, CISH inactivation significantly increased sensitivity to tumor antigen, enabling robust recognition and killing even at low antigen levels, conditions that often lead to treatment failure with conventional T cell therapies, mirroring antigen escape scenarios seen in solid tumors. Our findings further validate CISH as a potent and druggable intracellular checkpoint capable of boosting anti-tumor T cell responses across diverse cancer types, independent of PD-L1 status. The underlying mechanisms of CISH inhibition may help explain the positive outcomes reported in recent clinical studies of this approach in solid tumor immunotherapy.

Functional interrogation of the intracellular checkpoint CISH sheds light on its superior role in boosting anti-tumor T cell responses compared to other emerging targets across diverse cancer types.

Introduction

Cancer immunotherapies that boost the immune system’s ability to target tumors have transformed oncology, significantly extending survival in patients with advanced cancers. Among these, immune checkpoint inhibitors (ICIs) have become standard care for various malignancies^1,2. These therapies block inhibitory receptors like PD-1 and CTLA-4, which normally suppress T cell activity to maintain self-tolerance, with numerous inhibitors now commercially available^3–8. PD-1-targeting monoclonal antibodies are approved for several cancers including metastatic melanoma, non-small cell lung cancer (NSCLC), lymphoma and colorectal cancer (CRC)^5,8. However, their efficacy largely depends on PD-L1 expression in the tumor microenvironment, a factor that varies between patients and can decrease during treatment. Consequently, 30–60% of patients fail to respond to PD-1/PD-L1 blockade^5,9,10.

To improve clinical outcomes, a new class of intracellular immune checkpoints (ICs) are being explored both preclinically and clinically. Unlike conventional surface ICs that suppress T cell activity through ligand interactions, intracellular ICs act within the cell to inhibit TCR and cytokine signaling, offering potential for broad anti-tumor effects regardless of tumor type or biomarker status. We and others have identified CISH (cytokine-induced SH2 protein) as a key intracellular IC that is induced in CD8⁺ T cells following TCR stimulation, impairing anti-tumor function by targeting PLC-γ1 for proteasomal degradation^11–13. It is markedly upregulated in tumor-infiltrating lymphocytes (TILs) compared to matched peripheral blood lymphocytes (PBLs)¹¹. CISH depletion in patient-derived, neoantigen-selected TILs enhances proliferation, cytokine secretion, polyfunctionality, and tumor neoantigen recognition¹¹. Notably, combining Cish knockout with PD-1 blockade in a preclinical murine model led to synergistic tumor regression¹².

Historically, intracellular ICs have been difficult to target but advances in T cell gene-editing, such as CRISPR/Cas9, have opened new therapeutic possibilities. The anti-cancer potential of CISH depletion was recently tested in a first-in-human clinical trial for metastatic colorectal cancer (NCT04426669)¹⁴, using CRISPR-mediated CISH deletion. This study reported an exceptional complete response attributed to CISH knockout, underscoring CISH’s promise as a novel checkpoint addressing this unmet need and raising the prospect of small molecule inhibitors to broaden clinical access¹⁵.

Building on these findings, we further investigated CISH and other emerging intracellular ICs using multiplexed gene-editing of primary human T cells, followed by functional assays in cancer antigen-specific in-vitro systems.

Results

Unlike PD-1, enhanced T cell effector and cytolytic functions by CISH inactivation are tumor cell surface ligand-independent

CISH differs from conventional immune checkpoints like PD-1, which rely on tumor ligand interactions, by acting intracellularly to suppress TCR signaling independently of target cell ligands¹¹. Inhibiting CISH could therefore enhance anti-tumor responses across tumor types and biomarker statuses, crucial since many patients do not respond to PD-1/PD-L1 blockade^5,9,10. Unlike CISH deletion, knockout of the PDCD1 gene encoding PD-1 did not enhance T cell effector function or memory formation following αCD3 stimulation, suggesting that PD-1 inactivation alone does not confer benefit when TCR engagement occurs in the absence of its ligand, PD-L1. (Fig. 1a–c, Supplementary Fig. 1a). (See Supplementary Fig. 1b for exhaustion markers in unstimulated CISH and PD-1 depleted CD8⁺ cells). To compare their functional impact, we evaluated CISH- and PD-1-deficient T cells in a KRAS^G12D cancer model using CD8⁺ T cells engineered to express a KRAS^G12D-specific TCR via TRAC locus knock-in^16,17 (Fig.1d, Supplementary Fig. 1c). We selected the KRAS^G12D antigen because mutations in the KRAS oncogene are common and play a key role in the initiation and progression of many human cancers, making it a clinically relevant target for assessing T cell responses. The enhanced effector phenotype from CISH deletion persisted upon cognate antigen stimulation, either by peptide-pulsed H-1975 cells or direct TCR activation, even at low antigen levels (e.g.: 0.1 µg/mL), as shown by cytokine profiling (Supplementary Fig. 1d, e). This enhanced effector phenotype was evident even in response to low levels of pulsed KRAS^G12D peptide (e.g.: 0.1 ug/ml) (Supplementary Fig. 1e). Finally, we assessed cytolytic activity of KRAS^G12D-TCR⁺ CD8⁺ cells against PD-L1/2-expressing H-1975 targets (Supplementary Fig. 1f). CISH-deficient cells demonstrated superior antigen-specific killing compared to PD-1 KO cells, and this effect was independent of PD-L1/2 expression. (Fig. 1d, Supplementary Fig. 1f–i).

Fig. 1. Unlike PD-1, enhanced T cell effector and cytolytic functions by CISH inactivation are tumor cell surface ligand-independent.

a, b Effector cytokine production (a) and polyfunctionality (b) of CD8⁺ T cells following TCR stimulation via αCD3 stimulation (1 ug/ml), as measured by ICS. c T cell memory formation measured by flow cytometry after TCR stimulation. (T_{Naive =} CD45RO^-, CD62L⁺; T_CM = CD45RO⁺, CD62L⁺; T_Eff = CD62L^-). d KRAS^G12D antigen-specific cytolysis: Kinetic impedance assay using xCELLigence enables real-time measurement of cytolysis of KRAS^G12D expressing H-1975 cells by edited KRAS^G12D-TCR⁺ CD8⁺ T cells. (4:1 Effector: Target ratio). a–d: Statistical significance was determined by either One or Two-way ANOVA with multiple comparison test for repeated measures. P values shown in graphs. Not shown = not significant, All data are representative of three independent experiments. Data are mean ± SEM.

CISH inactivation enhances T cell function more effectively than the inactivation of other prominent intracellular immune checkpoints

To compare the immune-enhancing effects of CISH knockout with other emerging intracellular immune checkpoints (Fig. 2a), we developed a CRISPR/Cas9-based multiplex gene-editing strategy for primary human CD8⁺ T cells, followed by phenotypic analysis after TCR stimulation (Supplementary Fig. 3a). We assessed whether CISH deletion increases T cell sensitivity to low-level TCR stimulation, mimicking low antigen expression in tumors, as previously observed in KRAS^G12D-TCR⁺ CD8⁺ cells (Supplementary Fig. 1e). CISH-deficient cells showed significantly elevated production of effector cytokines IFNγ, TNFα, and IL-2 (Fig. 2b–f, Supplementary Fig. 3c–e), including higher IFNγ expression levels and frequency (Fig. 2b–d), as well as increased cytokine polyfunctionality (Fig. 2f). These effects were most pronounced at low αCD3 stimulation and sustained after 21 days in culture (Fig. 2b–e, Supplementary Fig. 3c, d). No significant differences were observed in the exhaustion phenotype of unstimulated (naïve) WT vs. CISH depleted cells (Supplementary Fig. 2a-b). As controls, CISH depleted CD8⁺ cells were also tested in an αCD3/CD28 bead (Supplementary Fig. 2c) and a repetitive antigen (αCD3) challenge (Supplementary Fig. 2d) protocol where CISH depletion was also able to enhance T cell effector function, consistent with previous reports¹⁷.

Fig. 2. CISH inactivation enhances T cell function more effectively than the inactivation of other leading intracellular immune checkpoints.

a Intracellular immune checkpoint signaling in human T cells showing the unique and non-redundant roles of select checkpoint targets in the context of antigen receptor signaling. b Effector Cytokines: IFNγ production at day 21 (D21) in CD8⁺ cells after increasing levels of αCD3 stimulation, as measured by intracellular cytokine staining (ICS). Statistics shown vs. Control at each αCD3 dose. c Evaluation of IFNγ production in CD8⁺ cells upon 0.5 ug/ml αCD3 stimulation, over time post editing. Statistics shown vs. Control at each time point. d Intensity of intracellular IFNγ staining in stimulated immune checkpoint knockout (KO) cells (0.5 ug/ml αCD3 stimulated cells, at day 21). e Evaluation of TNFα and IL-2 production at day 21 upon 0.5 ug/ml αCD3 stimulation. f Ratio of polyfunctional CD8⁺ T cells after TCR stimulation via αCD3 stimulation (0.5 ug/ml, Day 21). Statistics shown vs. Control at each category. g Proportion of CD8⁺ cells with an effector memory phenotype at day 21 after editing upon 0.5 ug/ml αCD3 stimulation. Statistical significance was determined by One-way ANOVA vs. Control in each population. h CD8⁺ T cells activation status upon 0.5 ug/ml αCD3 stimulation (Day 21). i Expression of inhibitory receptor PD-1 in T cells at low αCD3 stimulation (0.5 ug/ml), Day 21. b–i: Statistical significance was determined by ANOVA vs. Control (unless otherwise stated); P values shown in graphs or *P ≤ 0.05; **P ≤ 0.01; ***P ≤ 0.001; ****P ≤ 0.0001; not shown = not significant. All data is representative of at least three independent experiments, N = 6. Data represents mean ± SEM.

CISH knockout induced a stronger program of T cell activation (Fig. 2h), effector function (Fig. 2b–f), and effector memory T cell formation (Fig. 2g) than knockouts of other intracellular ICs, including RASA2, CBLB, SOCS1, REGNASE1 and HPK1, under low αCD3 stimulation. Notably, unlike RASA2 and CBLB knockouts, PD-1 expression at low αCD3 stimulation remained comparable between control and CISH-deleted cells (Fig. 2i), suggesting a distinct regulatory profile. These data demonstrate that CISH deletion more effectively enhances T cell activation upon TCR ligation than other intracellular immune checkpoints tested.

CISH functions as a non-redundant regulator of antigen-specific T cell cytolysis, and its inactivation can synergize with other intracellular immune checkpoints

The complexity of TCR signaling suggests that multiple non-overlapping pathways are governed by distinct intracellular ICs (Fig. 2a). To explore potential synergistic effects, we expanded our CRISPR multiplex editing platform in primary human T cells to test combinations of CISH depletion with other ICs (Supplementary Fig. 4a). Using low αCD3 stimulation (0.5 μg/ml), we observed an additive increase in cytokine production, notably IFNγ, TNFα, and IL-2, only when CISH was co-deleted with SOCS1, HPK1, or RASA2 (Fig. 3a, b), supporting non-redundant roles in regulating T cell effector functions. However, no additive effects were seen in the frequency of effector memory T cells or activation status with these combinations (Supplementary Fig. 4b, c).

Fig. 3. CISH functions as a non-redundant regulator of antigen-specific T cell cytolysis, and its inactivation can synergize with other intracellular immune checkpoints.

a T cell effector function when CISH is depleted in CD8⁺ cells in combination with other intracellular immune checkpoints. Effector cytokines production at day 21 post editing upon 0.5 ug/ml αCD3 stimulation, measured by ICS. b Polyfunctionality of CD8⁺ T cells after TCR stimulation via αCD3 stimulation when CISH is depleted in combination with other intracellular immune checkpoints. a, b: Statistical significance was determined by One-way ANOVA vs. CISH KO. P values shown in graphs or *P ≤ 0.05; **P ≤ 0.01; ***P ≤ 0.001; ****P ≤ 0.0001; not shown = not significant. All data is representative of at least three independent experiments, N = 6. Error bars represent mean ± SEM. c Antigen-specific killing in a model of KRAS^G12D solid tumor when CISH is depleted in combination with other intracellular immune checkpoints: Cytolysis of KRAS^G12D expressing H-1975 cells by edited KRAS^G12D-TCR⁺ CD8⁺ T cells. (2:1 Effector: Target ratio). Statistical significance was determined by One-way ANOVA vs. CISH KO at 24 h timepoint. P values shown in graph; not shown = not significant. All data is representative of at least three independent experiments. Error bars represent mean ± SEM.

In the same KRAS^G12D antigen-specific solid tumor model, CISH deletion enhanced tumor cell killing by KRAS^G12D-TCR⁺ T cells (Fig. 3c), and this effect was further potentiated by co-deletion with RASA2 or SOCS1. The observed synergy with SOCS1 is consistent with its role as a member of the same SOCS family as CISH. Notably, recent work by Schlabach et al.¹⁸ (KSQ-001EX) demonstrated that SOCS1 inactivation enhances anti-tumor activity in PD-1–refractory models and preclinical TIL studies, supporting the rationale for dual CISH/SOCS1 checkpoint inhibition in cancer therapy.

We also evaluated HPK1, a kinase under active clinical investigation as a small molecule checkpoint inhibitor^19–21. Although co-deletion of CISH and HPK1 increased effector cytokine production (Fig. 3a, b), it did not enhance antigen-specific cytolysis (Fig. 3c), suggesting a partial functional synergy.

Additionally we assessed PTPN1 and PTPN2, phosphatases targeted by novel small-molecule inhibitors currently in clinical trials (NCT04777994²², NCT04417465²³). CISH deletion alone outperformed PTPN1, PTPN2, and combined PTPN1/2 knockouts in terms of T cell activation, cytokine production, polyfunctionality, and cytolytic activity (Supplementary Fig. 5a–e). Co-deletion with CISH did not yield additive benefits in cytolysis, indicating that CISH operates through distinct, dominant pathways in regulating anti-tumor T cell function.

CISH inactivation enhances CD19-specific CAR⁺ T cell cytotoxicity against low-antigen expressing cancer cells

Adoptive cell therapies (ACT) using tumor-specific T cells have shown strong clinical efficacy in certain hematological malignancies^24,25, but success in solid tumors remains limited^26,27. To assess the translational relevance of CISH deletion, we extended our prior work with KRAS^G12D-TCR⁺ T cells to a CD19-CAR⁺ T cell model targeting hematologic cancers. Primary human CD8⁺ T cells were retrovirally transduced with a second-generation CD19-specific CAR incorporating a CD28 costimulatory domain^28,29 (Supplementary Fig. 6b). These CD19-CAR⁺ T cells were then co-cultured with NALM6 B-cell leukemia cells engineered to express varying levels of CD19 (Supplementary Fig. 6a), and cancer cell killing was measured by luciferase activity.

CISH knockout significantly enhanced tumor cell lysis (Fig. 4a, b, Supplementary Fig. 6c) and antigen-specific IFNɣ production (Fig. 4c), including at low CD19 expression levels, recapitulating the enhanced sensitivity observed previously (Fig. 2b, Supplementary Fig. 1e). Similar results were obtained in a non-B cell line (H-1975) expressing CD19, confirming that CISH inactivation broadly boosts antigen-specific CAR-T cytotoxicity (Supplementary Fig. 6d, e).

Fig. 4. CISH inactivation enhances CD19-specific CAR T cell cytotoxicity against low-antigen expressing cancer cells.

a NALM6 (GFP⁺ FF-Luc⁺) target cell killing by CD8⁺ CD19-CAR⁺ T cells in the presence (Control) or absence of CISH (CISH) at effector: target ratio 1:2. Cytolysis was measured by remnant luciferase activity assay after 18 h of co-culture. b Left, CD19 expression (colored) on engineered NALM6 target cells compared with unstained cells (gray). Right, CD19-CAR⁺ T cell killing of NALM6 cells expressing varying levels of CD19. c Effector cytokine (IFNγ) levels in CD19 stimulated CD19-CAR⁺ CD8⁺ cells, as measured by ELIZA. a–c: Statistical significance was determined by Two-way ANOVA vs. Control: Not shown = not significant, P values shown in graphs or *P ≤ 0.05; **P ≤ 0.01; ***P ≤ 0.001; ****P ≤ 0.0001. All data is representative of at least three independent experiments. N = 10 donors. Error bars represent mean ± SEM. d–f Cytokine profile of CISH depleted CD19-CAR⁺ T cells after overnight co-culture with CD19 expressing NALM6 cells (as measured by nELISA). d Pathway analysis by Reactome.org. Graph shows significantly regulated pathways; p ≤ 0.05 (x-axis: Pathway Hierarchy/ Go Biological Process; y-axis; -Log₁₀(p-value)). e Volcano plot (Log₂FC vs. -Log₁₀(p-value); Left panel) and bar graph (Log₂FC ≥ 1.2); Right panel) of differentially regulated secreted factors in CISH KO vs. Control CD19-CAR⁺ CD8⁺ cells after overnight co-culture with CD19^WT NALM6 cells. f Significantly downregulated factors in CISH depleted CD19 stimulated CD19-CAR⁺ CD8⁺ cells. d–f Statistical significance was determined by either multiple t-test or Two-way ANOVA vs. Control in each treatment. Not shown = not significant, P values shown in graphs or *P ≤ 0.05; **P ≤ 0.01; ***P ≤ 0.001; ****P ≤ 0.0001. N = 3 donors. Data are mean ± SD.

To further characterize this phenotype, we performed a nELISA cytokine screen (Nomic Bio) after overnight co-culture of CD19-CAR⁺ T cells with CD19⁺ NALM6 cells (Fig. 4d–f). CISH-deficient CAR-T cells exhibited elevated production of IFNɣ, TNFα, IL-5, CCL4, and GM-CSF, cytokines associated with T cell activation, tumor infiltration, and immune recruitment (Fig. 4d, e). Interestingly, CISH deletion also led to reduced secretion of immunosuppressive or tumor-promoting factors (Fig. 4e, f), including: Galectins 1 and 3, which negatively regulate T cells and induce apoptosis^30,31; Soluble 4-1BB (sCD137), known to attenuate co-stimulatory signaling^32,33; IL-1α, a DAMP molecule shown to inhibit CD8⁺ memory formation and IFNɣ production^34,35; and EMMPRIN (CD147), a tumor-associated glycoprotein that promotes immunosuppressive cytokine release and correlates with T cell exhaustion^36–38.

Collectively, these findings suggest that CISH inactivation enhances CAR-T cell function in part by increasing antigen sensitivity and effector function while reducing exposure to immunosuppressive factors. This makes CISH targeting a compelling strategy to improve T cell performance, particularly for tumors with low antigen density or immunosuppressive microenvironments.

Discussion

While single and combination therapies targeting cell surface immune checkpoints, such as PD-1 and CTLA-4 have produced notable clinical responses, a significant proportion of patients remain unresponsive or ultimately relapse. In this study, we demonstrate that genetic ablation of CISH enhances antigen sensitivity and sustains effector function in engineered human T cells. Unlike conventional surface checkpoints, CISH belongs to a novel class of intracellular regulators that act downstream of the TCR and function independently of ligand engagement. This ligand-independent mechanism offers a strategic advantage, as targeting CISH may overcome multiple upstream inhibitory signals simultaneously, regardless of tumor type or biomarker expression. These findings support the therapeutic promise of CISH inhibition as a broadly applicable and potentially superior approach to current immune checkpoint blockade strategies.

Several emerging intracellular immune checkpoints have been proposed as promising targets to enhance T cell-based immunotherapies^18,39–43. Notably, Carnevale et al. demonstrated that RASA2 depletion in NY-ESO-1-specific TCR-T cells, as well as in CD19- and EphA2-targeting CAR-T cells, significantly improved tumor control and survival in both hematologic and solid tumor models³⁹. Similarly, Wei et al. identified REGNASE1 as being highly expressed in tumor-infiltrating lymphocytes (TILs) and showed that its deletion enhanced adoptive cell therapies (ACTs) by promoting the development of long-lived effector T cell phenotypes⁴². While these regulators act at distinct nodes of the T cell activation network, their functional roles partially overlap or converge on shared signaling cascades, such as MAPK, JAK-STAT, and NF-κB, resulting in redundancy in maintaining immune homeostasis and preventing overactivation. In direct comparison, CISH ablation induced a more robust program of T cell activation, marked by enhanced antigen-specific cytolysis, effector cytokine production, and effector memory T cell formation. These improvements were consistently superior to those observed with other intracellular ICs, including RASA2, CBLB, SOCS1, REGNASE1, HPK1, and PTPN1/2, particularly under conditions of low antigen expression, a common feature of tumor escape.

Of note, Wiede et al.⁴⁴ describes PTPN2 knockdown as sufficient to upregulate cytokine expression by HER-2-CAR⁺ T cells. The discrepancy between Wiede et al.⁴⁴’s findings and our studies’ experimental setup may be attributed to the following factors. 1) Signal Strength: while in Wiede et al.⁴⁴ HER-2-CAR⁺ T cells were stimulated with 24JK sarcoma cells expressing HER2 antigen, a strong stimulation, in our experimental system CD8⁺ cells were stimulated with low levels of αCD3 (0.5ug/ml), a weak stimulation (Supplementary Fig. 5). 2) Differences in downstream signaling cascades: the HER2-CAR used in Wiede et al.⁴⁴ is a second-generation CAR that engaged CD28 signaling, which significantly enhances T-cell signaling; whilst in our experimental system, CD8⁺ cell activation relied solely on low αCD3 stimulation and did not engage CD28 signaling, leading to distinct downstream signaling cascade activation landscapes.

Additionally, our findings demonstrate that the concurrent inactivation of CISH and select ICs (RASA2 and SOCS1) can act in a non-redundant synergistic manner to significantly further improve the tumor-specific cytolytic potential of T cells, offering the compelling prospect of a dual-therapeutic approach. Future in-vivo studies will strengthen these findings. Importantly, CISH appears to exhibit a particularly broad suppressive role, integrating both cytokine and TCR signaling, and may offer a more comprehensive strategy to enhance T cell efficacy compared to single-pathway regulators.

The finding that CISH ablation enhances T cell effector and cytolytic functions downstream of distinct antigen receptors, including both chimeric antigen receptors and T cell receptors (e.g.: KRAS^G12D-specific), further supports the receptor-independent nature of CISH biology. This highlights the broad therapeutic potential of CISH inhibition to enhance the anti-tumor activity of diverse T cell modalities, whether engineered CAR-T cells, patient-derived tumor-infiltrating lymphocytes, or endogenous peripheral tumor antigen-specific T cells. Most importantly, CISH deletion significantly increased T cell sensitivity to antigen, a benefit observed in in-vitro models using CAR- and TCR-engineered T cells. In these contexts, CISH-deficient T cells were able to recognize and kill tumor cells expressing even very low levels of antigen, conditions that closely mirror tumor antigen escape, a key mechanism by which cancers evade conventional T cell therapies.

CISH ablation in CD19-CAR⁺ T cells also showed significantly reduced levels of secreted Galectins 1 and 3, even in the absence of CD19 stimulation, perhaps due to impact on cytokine receptor signaling. Galectins (Gal) are a family of mammalian β-Galactoside-binding proteins that play a key role in T cell homeostasis in the TME by acting as negative extracellular checkpoints of T cell function, inducing lymphocyte deactivation, and promoting T cell death^30,31. Indeed, both Gal-1 and Gal-3 inhibition significantly reduces tumor growth, increases activated CD8⁺ infiltration into the TME, and increases IFNɣ secretion from CD8⁺ TILs improving their cytotoxicity^45–48. Increasing attention has been paid on Gal-1 and Gal-3 inhibitors which have shown enormous potential in tumor therapy with Gal-1 inhibition boosting anti-PD1 therapy responses^31,45.

Notably, CISH-depleted CD19-CAR⁺ T cells also exhibited significantly elevated levels of IL-8 and IL-5 (Fig. 4e) in NALM6 co-cultures. While IL-8 has been shown to exert pro-tumorigenic effects within the tumor microenvironment, and IL-5 may be linked to tumor metastasis, further research would be needed to elucidate CISH’s role in this context.

The most compelling evidence supporting the therapeutic potential of targeting CISH comes from the first-in-human clinical trial employing CRISPR-mediated knockout of CISH in T cells administered to patients with metastatic colorectal cancer (NCT04426669¹⁴). In this study, CISH-knockout Tumor-Infiltrating Lymphocytes were used as a model system to evaluate the safety and efficacy of CISH inactivation. Remarkably, the trial reported an ongoing and durable complete response lasting over two years in a young adult patient with colorectal cancer that was refractory to multiple lines of chemotherapy and immunotherapy. This outcome is particularly noteworthy given the grim prognosis typically associated with metastatic colorectal cancer, where treatment options are extremely limited, and survival outcomes are almost uniformly poor. The durable complete response in such a heavily pretreated patient provides strong clinical validation of the potential of CISH as a therapeutic target.

Given this landmark clinical result, together with mounting preclinical evidence demonstrating the broad utility of CISH targeting across both hematological malignancies and solid tumors, next-generation small molecule drugging strategies aimed at modulating CISH are now in advanced stages of development.

Methods

Cell lines

H-1975 (CRL-5908) and Phoenix-AMPHO (CRL-3213) cells were obtained from ATCC. H-1975 cells stably expressing mutant KRAS^G12D were sourced from Horizon Discovery (HD 120-001). The NALM6 FFLuc/GFP reporter cell line was purchased from Creative Bioscience (CSC-RR0360). All cell lines were cultured under recommended media formulations and growth conditions, maintained at sub-confluent densities, and routinely tested for mycoplasma contamination.

Generation of CD19-expressing NALM6 and H-1975 cells

NALM6 CD19^KO cells were generated using a Synthego CRISPR knockout kit targeting the CD19 gene. To create NALM6 cell lines with varying levels of CD19 expression, CD19^KO cells were transduced with a lentiviral vector encoding PGK-driven expression of truncated CD19 (lacking the intracellular signaling domain). Individual clones exhibiting distinct CD19^Low/Med expression were selected and sorted for uniform expression profiles. For H-1975 cells expressing CD19, the parental H-1975 cell line (CRL-5908) was transduced with a lentiviral vector encoding EF1A-driven expression of truncated CD19.

Isolation and culture of primary T cells from healthy donors

All ethical regulations relevant to human research participants were followed. Leukopaks were purchased from the NHS Blood and Transplant Bank (NHSBT) from anonymized healthy individuals and handled and stored in accordance with the Human Tissue Authority UK regulations. PBMCs were isolated by density-gradient centrifugation using lymphocyte separation medium (Corning) and cryopreserved until ready for use. Total CD8 + T cells were isolated from unfractionated PBMCs using either the EasySep™ or MojoSort™ Human CD8 + T Cell Isolation Kit (Stem Cell Technologies, Biolegend) and following manufacturer’s guidelines. Isolated human CD8 + T cells were cultured in X-VIVO-15 Basal Media (Lonza) supplemented with 10% Human AB Serum Heat Inactivated (Sigma), 300IU/ml Recombinant Human IL-2 (unless otherwise stated), 5 ng/ml Recombinant Human IL-7, and 5 ng/ml Recombinant Human IL-15 (all Peprotech) and 10mM N-Acetyl-L-cysteine (Sigma) (henceforth referred to as complete T cell media) and cultured in a 37 °C, 5% CO2 culture incubator. Media was replaced every 2-3 days with fresh complete media including cytokines.

Immune checkpoint edited CD8⁺ T cells using CRISPR/Cas9

CD8⁺ T cells were stimulated using TransAct (1:100, Miltenyi Biotec, 130-128-758) in complete T cell media and under normal growth conditions for 48–72 h prior to electroporation. T cells (1-2 million) were electroporated with Cas9–sgRNA–RNP (60 pmol Cas9: 2–4 ug sgRNA) using Lonza’s P3 Primary Cell Nucleofector® Kit (V4SP-3096) program EH-115, following manufacturer’s protocol. Unless otherwise stated, control-edited T cells were targeted with the AAVS1 sequence. All sgRNA sequences are listed in Table S1. All guide RNAs were ordered as Alt-R CRISPR-Cas9 sgRNAs from Integrated DNA Technologies (IDT) except for PTPN1 and PTPN2, which were purchased as ready Gene-Knock out kits from EditCo.

Analysis of Gene Knockout Efficiency on DNA Level

Primers for PCR were designed to amplify a 600–900 basepair region surrounding the sgRNA target site. Following a minimum of 48 h after electroporation, genomic DNA was extracted from CD8⁺ T cells and target regions were amplified by PCR using the GoTaq G2 PCR mastermix (Promega). Correct and unique amplification of the target regions was verified by agarose gel electrophoresis before purifying PCR products using the QIAquick PCR Purification Kit (Qiagen). For analysis by TIDE, PCR amplicons were Sanger sequenced and paired ab1 files of control versus edited samples were analyzed using Synthego’s ICE tool (https://ice.synthego.com). All primer sequences used are listed in Table S1.

Production of AAV-mediated TCR-Knock-in and checkpoint-knockout CD8⁺ T cells using CRISPR/Cas9

Isolated CD8⁺ cells were thawed, activated and electroporated as described above, followed by a resting period at room temperature for 15 min to allow for cell recovery. After recovery, cells were resuspended in prewarmed complete T cell media. To achieve targeted KRAS^G12D-TCR knock-in, rAAV6 was added to CD8⁺ T cells 2–5 h after electroporation at an MOI of 1 × 10⁶. Viral rAAV6 particles were produced by Signagen. Electroporated T cells were recovered in complete T cell media at a density of 1×10⁶ cells per ml and allowed to rest for 48 h before subsequent analysis.

Flow cytometry analysis of T cell phenotypes

For flow cytometric analyses of the CRISPR-edited T cell phenotypes and cell surface marker expression, cells were harvested from culture plates and washed using FACS Buffer containing PBS with 0.5% Bovine Serum Albumin (Thermo Scientific), followed by staining with fluorophore-conjugated monoclonal antibodies against cell surface markers. All antibodies used for flow cytometric analyses are listed in Table S2. Live/Dead Fixable Dead Cell Stains (Invitrogen) were included in all experiments to exclude dead cells. All samples were acquired on an Cytek Aurora cytometer (Cytek^® Biosciences), and data was analyzed using FlowJo 10 software (BD Biosciences). All antibodies used for flow cytometric analyses are listed in Table S2.

Intracellular cytokine staining

Cells were stimulated for a total of 6 h with anti-hCD3 (OKT3) or anti-mTCRb (H57) (as stated in figure legend). After one hour of stimulation GolgiStop solution (BD Bioscience) was added (a total of 5 h block of intracellular protein transport). As a positive control for cytokine production, a pool of T cells was stimulated for 6 h with 50 ng/ml PMA and 1ug/ml Ionomycin (Sigma). T cells were then harvested and washed with FACS Buffer and stained for surface markers followed by fixation and permeabilization using BD Cytofix/Cytoperm Fixation/Permeabilization Solution (ThermoFisher) before proceeding with intracellular cytokine staining. Live/Dead Fixable Dead Cell Stains (Invitrogen) were included in all experiments prior to staining to exclude dead cells.

Realtime Cytolysis Assay (xCELLigence)

Cytolysis assays were performed using the xCELLigence RTCA SP platform (Agilent), which measures electrical impedance to generate a cell index (CI). Background readings were recorded with media alone prior to seeding. Adherent H-1975 tumor cells were plated in 96-well RTCA View plates at a density optimized to enter the linear growth phase after 14–18 h of incubation at 37 °C and 5% CO₂ in complete growth medium. The following day engineered T cells (with indicated gene edits) were added at specified effector-to-target (E:T) ratios. Assays were run undisturbed for up to 90 h, with impedance measurements taken every 2–10 min. Data were analyzed using RTCA software and expressed as % cytolysis, calculated as: % Cytolysis = [(CI_target only – CI_with effectors) ÷ CI_target only] × 100. Controls included: Background (media only); Negative control (target cells only); Positive control (target cells treated with 2.5% Triton X-100 for maximum cytolysis).

CD19-CAR⁺ T production

Preparation of retrovirus: CD19-CAR retrovirus (Yescarta^28,29) was generated by transient transfection of Phoenix-AMPHO packaging cells. Briefly, 5 million Phoenix-AMPHO cells were plated per 10-cm dish 24 h before transfection in DMEM + 10% FBS and transfected with 20ug of CD19-CAR plasmid (Yescarta^28,29) using Lipofectamine Plus following manufacturer’s instructions. Viral supernatant was collected after 48 h and loaded onto RetroNectin-coated (10 ug/ml, Takara Bio) non-TC 24-well plates and centrifuged at 2000 g for 2 h at 32 °C.

Engineering of CISH depleted CD19-CAR⁺ T cells

CD8⁺ cells from healthy donors’ PBMCs were stimulated with TransAct (1:100, Miltenyi Biotec, 130-128-758) for 48 hours before KO/transduction. Stimulated cells were then CRISPR-engineered to knock-out immune checkpoints (ICs). Preformed Cas9–sgRNA–RNP (60 pmol Cas9: 2-4 ug sgRNA) and CD8⁺ cells were gently resuspended in P3 buffer with supplement (Lonza’s P3 Primary Cell Nucleofector® Kit, V4SP-3096) at 2 million cells per 20 μl, and pulsed with code EH-115. After electroporation, the 4D-Nucleofector cuvette was placed at room temperature for 15 min to allow for cell recovery. After recovery, cells were resuspended in prewarmed X-vivo media supplemented with 300IU/ml IL-2, 5 ng/ml IL-7 and 5 ng/ml IL-15. Cells were then transduced with CD19-CAR viral supernatant by transferring them onto the RetroNectin/virus coated plates (see above), and spun at 32 °C for 30 mins at 300 rpm. Cells were then transferred to 37 °C incubator to rest overnight. Cells were assessed for transduction efficiency after 3–4 days by measuring surface expression of CD19-CAR by flow cytometry. If needed, CD19-CAR⁺ CD8⁺ cells were enriched using the MACSprep™ CD19 CAR MicroBead Kit following manufacturer’s instructions (Miltenyi Biotec, 130-127-866).

CD19-CAR⁺ T cells in-vitro Cytolysis Assay

CD19-CAR⁺ CD8⁺ T cells were co-cultured with pre-plated GFP⁺ FF-Luc⁺ NALM6 tumor cells in a 96-well flat bottom plate starting at a 1:2 E:T ratio, then with a log₂ serial dilution in triplicates. After 18 h, NALM6 viability was measured by luciferase.

Statistics and Reproducibility

Appropriate statistical tests were used to analyze data, as described in each figure legend. Statistical differences between two sample groups, where appropriate, were analyzed by a standard Student’s two-tailed, non-paired, t-test and between three or more sample groups using two-way or three-way ANOVA using GraphPad Prism Software version 10. Significance was preset at P < 0.05. P and N values are indicated in the figures where statistical analyses have been carried out.

Author contributions

Conceptualization: F.C., T.H., M.C., C.C., E.L., and A.C. Methodology and Investigation: In-vitro CRISPR validation studies were performed by M.C. and C.C.; in-vitro characterization of effects of ICs manipulation in T cells was performed by A.B.B., M.C., C.C., R.K.C.V., and P.M.; KRASG12D-TCR-T cell experiments were performed by A.B.B, R.K.C.V., and P.M.; CAR-T cell experiments were performed by O.P., E.B., P.M., and F.C. Experiments were done in collaboration with the preclinical therapeutics’ cores at UMN (N.S., M.J., B.R.W., B.S.M., and E.L.) and MSKCC (C.A.K.). Data curation/Analysis: F.C.; C.A.K. contributed to the data interpretation. Project Supervision: T.H. and M.Ch. Funding acquisition: M.Ch. Writing (original draft preparation, review & editing): F.C., T.H., and M.Ch.

Peer review

Peer review information

Communications Biology thanks Nicholas Huntington and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Primary Handling Editors: Gloryn Chia and Johannes Stortz.

Data availability

The data supporting the findings from this study are available within the manuscript and its supplementary information. Values for all the graphs in the figures and Supplementary Figs. are provided in the Supplementary_Data file. All other data are available from the corresponding author (or other sources, as applicable) on reasonable request.

Competing interests

C.A.K. is an inventor on patents related to TCR discovery and public neoantigen-reactive TCRs unrelated to the present manuscript that have been licensed to Intima Biosciences and is recipient of licensing revenue shared according to MSKCC institutional policies. C.A.K. has consulted for or is on the scientific advisory boards for Achilles Therapeutics, Affini-T Therapeutics, Aleta BioTherapeutics, Bellicum Pharmaceuticals, Bristol Myers Squibb, Catamaran Bio, Cell Design Labs, Chronara Biosciences, Decheng Capital, G1 Therapeutics, Klus Pharma, Merck, Obsidian Therapeutics, PACT Pharma, Roche/Genentech, Royalty Pharma, Stereo Biotherapeutics, and T-knife. C.A.K. is a scientific co-founder and equity holder in Affini-T Therapeutics. The other authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at 10.1038/s42003-026-09579-x.