Ponatinib inhibits LCK and PI3K signaling and promotes CD8⁺ T stem cell memory cell development

Abstract

CD8⁺ T stem cell memory (T_SCM) cells show clinical promise for cancer immunotherapy, but T_SCM cell generation in clinical settings requires further optimization. Ponatinib is a tyrosine kinase inhibitor primarily targeting BCR-ABL1 and used for the treatment of chronic myeloid leukemia. Here, we investigate the effect of ponatinib on T cell activation and differentiation. Acting off-target, ponatinib inhibits LCK and PI3K signaling to enhance the transcriptional functions of TCF7 and FOXO1, thereby promoting CD8⁺ T_SCM cell differentiation. Mechanistically, stable and sustained, but not intermittent, inhibition of the LCK and PI3K pathways is essential for CD8⁺ T_SCM cell induction. In mouse tumor models, ponatinib treatment exhibits antitumor efficacy alone and in combination with PD-1 blockade. Furthermore, ponatinib increases chimeric antigen receptor (CAR) T_SCM cells by reducing CAR T cell exhaustion, resulting in durable antitumor efficacy. Our results thus implicate ponatinib as therapeutic immunomodulator, inducing T_SCM cells for improved antitumor T cell activity.

As a tyrosine kinase inhibitor targeting BCR-ABL, ponatinib has been used to treat chronic myeloid leukemia. Here, the authors identify LCK and PI3K signaling as additional ponatinib targets, with ponatinib inducing CD8⁺ T stem cell memory cells, thereby boosting immune checkpoint blockade and CAR T cell therapy.

Introduction

Cancer immunotherapies, such as immune checkpoint inhibitors (ICI), including PD-1 blockade and chimeric antigen receptor (CAR)-T cell therapy, have been applied in clinical settings. While cancer immunotherapies provide durable clinical benefit in some patients, even those with advanced disease, more than half of treated patients develop resistance to therapy¹. Therefore, many attempts have been made to identify biomarkers that can stratify responder patients and to develop new effective cancer immunotherapies. Reassuringly, among CD8⁺ T cells, which constitute a key effector cell population that kills tumor cells, CD8⁺ T stem cell memory (T_SCM) cells have been shown to contribute to controlling tumor growth in both ICIs and CAR-T cell therapy^2–5. However, strategies to induce T_SCM cells remain to be determined.

Ponatinib, a third-generation tyrosine kinase inhibitor (TKI), is approved for the treatment of patients with chronic myeloid leukemia (CML) who are resistant or intolerant to prior therapy or who have the T315I mutation in their CML cells, and patients with Philadelphia chromosome-positive acute lymphoblastic leukemia (Ph⁺ ALL)^6–8. While the primary mechanism of action of ponatinib is known to target BCR-ABL1, ponatinib also inhibits a broad spectrum of protein tyrosine kinases that are critical for the progression of several cancers^6,9. In addition, preclinical studies have demonstrated its antitumor efficacy in both hematologic malignancies and solid tumor models, in which these kinase pathways play key roles in survival and proliferation^10–12. Consequently, the therapeutic application of ponatinib has been investigated in multiple hematologic malignancies and solid tumors^13–15. However, the efficacy of molecularly targeted therapy, including TKIs, is frequently limited by various resistance mechanisms, such as tumor heterogeneity and the emergence of new genetic alterations in tumor cells^16,17. We previously reported that imatinib, a first-generation TKI, inhibits lymphocyte-specific protein tyrosine kinase (LCK), a T cell-specific signaling molecule, as an off-target effect and selectively impairs regulatory T (Treg) cells, resulting in increased antitumor immunity¹⁸. These observations raise the possibility that distinct kinase inhibitors may differentially modulate T cell subsets, prompting us to examine whether ponatinib exerts broader immunomodulatory effects beyond Treg cells.

Here, we investigate the immunomodulatory potential of ponatinib and show that it modulates T cell activation and differentiation to promote a T_SCM-like phenotype. Ponatinib acts via off-target inhibition of key T cell signaling pathways and enhances transcriptional programs associated with stemness and functional persistence. This effect is preferentially observed under stable and sustained, rather than intermittent, pathway inhibition. Together, these findings suggest that kinase inhibitors can be used to modulate antitumor T cell responses and to improve the quality of T cell products for adoptive immunotherapy. Given that ponatinib is clinically available, our results suggest that it may become a potential antitumor drug with dual functions: augmenting antitumor immunity and inducing direct tumor cell killing through inhibition of kinase activity.

Results

Ponatinib affects both CD8⁺ T cells and Treg cells in humans

Kinase inhibitors targeting multiple kinases, essential for T cell activation/differentiation, may be promising reagents for modulating tumor immunity. Among the clinically available TKIs, ponatinib possessed a broader range of target kinases, including LCK, FYN, ITK, and RAF1, than other kinase inhibitors (Supplementary Fig. 1a, b)^6,9, which prompted us to investigate the immunomodulatory effects of ponatinib on antitumor immunity. We first examined whether ponatinib affects the proliferation and viability of Treg cells and effector T cells, particularly CD8⁺ T cells from human peripheral blood mononuclear cells (PBMC). Purified CD8⁺ T cells and Treg cells were stimulated with anti-CD3/CD28 monoclonal antibodies (mAb) in the presence of graded concentrations of ponatinib (Fig. 1a). At 50 nM, ponatinib significantly inhibited Treg cell proliferation and induced apoptosis, while CD8⁺ T cell proliferation was not impaired (Fig. 1b). Since this concentration recapitulated the trough level (approximately 40 nM) observed in patients, the trough levels of ponatinib¹⁹ and imatinib²⁰, were employed in subsequent experiments to assess their effects on different T cell populations.

Fig. 1. Ponatinib limits CD8⁺ T cell differentiation and reduces Treg cells in human PBMCs.

a, b Proliferation inhibition and apoptosis induction by ponatinib. CD8⁺ T cells and regulatory T (Treg) cells purified from the peripheral blood mononuclear cells (PBMC) of healthy individuals were stimulated with anti-CD3/CD28 monoclonal antibodies (mAb) along with graded doses of ponatinib for 4 days. Proliferation and apoptosis were assessed by Carboxyfluorescein diacetate succinimidyl ester (CFSE) dilution and by Annexin V and Fixable Viability Dye staining, respectively. a Experimental scheme. Created in BioRender. https://BioRender.com/cxg8twg. b Representative histograms (left) and frequencies of proliferating cells and apoptotic cells (right) (n = 3 donors per group). The gray dashed lines indicate the trough level of ponatinib. c–f PBMCs purified from healthy individuals were cultured with dimethyl sulfoxide (DMSO; control), imatinib, or ponatinib and analyzed by flow cytometry on Days 3 and 9. c Experimental scheme. Created in BioRender. https://BioRender.com/cxg8twg. d Representative contour plots (CD45RA and FOXP3) of CD4⁺ T cells in the DMSO (left, black), imatinib (left, blue), and ponatinib (left, red) groups and cell numbers of effector Treg (eTreg, CD4⁺CD45RA⁻FOXP3^hi) cells (n = 6 donors per group). e Cell numbers of CD8⁺ T cells (n = 6 donors per group). f Representative contour plots (CCR7 and CD45RA) of CD8⁺ T cells treated with DMSO (left, black), imatinib (left, blue), or ponatinib (left, red) and frequencies of naive (CCR7⁺CD45RA⁺) and effector memory (T_EM: CCR7⁻CD45RA⁻) subsets among CD8⁺ T cells (right) (n = 6 donors per group). Data in (d–f) were normalized to those of the DMSO control (set to 1.0). Data are presented as the mean ± SEM (b, d–f). Significance was assessed using two-sided paired ttests (b, d–f). All experiments were performed independently at least two times with similar results.

Human PBMCs were cultured in the presence of dimethyl sulfoxide (DMSO, control), imatinib, or ponatinib, and their effects on T cells were assessed (Fig. 1c). Like imatinib, ponatinib reduced the number and frequency of Treg cells (Fig. 1d and Supplementary Fig. 2a). Despite the reduction of Treg cells that suppress CD8⁺ T cell proliferation and activation²¹, ponatinib increased CD8⁺ T cells with a naive phenotype and reduced those with an effector phenotype without altering their total number (Fig. 1e, f). Consistently, the expression of activation and exhaustion markers by CD8⁺ T cells was decreased after ponatinib treatment (Supplementary Fig. 2b–d), suggesting that ponatinib, unlike imatinib, may directly affect both Treg cells and CD8⁺ T cells.

To clarify the direct effects of ponatinib, Treg cells and CD8⁺ T cells were purified from human PBMCs and treated with imatinib or ponatinib (Supplementary Fig. 3a, 4a). The number of Treg cells was significantly reduced by imatinib and ponatinib treatment (Supplementary Fig. 3b). In addition, ponatinib, but not imatinib, decreased the expression of activation markers (CD25, CTLA-4, GITR, and ICOS) and essential transcription factors (BATF and IRF4²²) in Treg cells (Supplementary Fig. 3c, d). In CD8⁺ T cells, ponatinib reduced the proportion of those expressing activation and exhaustion markers (CD25, CD69, and PD-1) without altering their total numbers, accompanied by the expansion of naive and T_SCM (CCR7⁺CD45RA⁺CD62L⁺CD95⁺) cell subsets^23,24 (Supplementary Fig. 4b–e). These results suggest that ponatinib directly affects both CD8⁺ T cells and Treg cells, which prompted us to investigate whether ponatinib facilitates the induction of T_SCM cells.

Ponatinib promotes T_SCM cell generation by modifying the early differentiation of naive CD8⁺ T cells

Since the proliferation analysis suggested that ponatinib could influence cell division patterns (Fig. 1a, b), we next investigated the phenotype of CD8⁺ T cells at each division stage (Supplementary Fig. 5a). T_SCM cells were abundantly observed in the ponatinib group, particularly among the undivided cells. In addition, highly divided cells ( ≥ 5 divisions) significantly increased (Supplementary Fig. 5b). To determine their origin, naive and effector memory (T_EM) CD8⁺ T cells were sorted and subjected to cell division assays (Supplementary Figs. 5c, d, 6a). In the cell division assays with naive CD8⁺ T cells, although the overall division patterns were comparable between the control and ponatinib groups, T_SCM cells significantly increased in the ponatinib group, and these cells accumulated in the undivided fraction compared with the control group (Supplementary Fig. 5e, f). In contrast, little T_SCM cell induction was detected in either group in the T_EM cell division assays, indicating that ponatinib affects the initial stages of naive CD8⁺ T cell differentiation, thereby promoting T_SCM cell development. Ponatinib treatment also significantly increased the proportion of vigorously dividing CD8⁺ T cells without affecting the overall proportion of divided cells in the T_EM cell division assays (Supplementary Fig. 5g, h). Thus, the highly divided cells observed in bulk CD8⁺ T cell cultures treated with ponatinib were derived from T_EM cell populations. Taken together, ponatinib fine-tunes the T cell activation signal, promoting T_SCM cell induction upon the activation of naive T cells. The fine-tuning of the T cell activation signal also contributes to maintaining the proliferative capacity of T_EM cells.

Ponatinib directly affects CD8⁺ T cell activation/differentiation

To further investigate the effects of ponatinib on CD8⁺ T cells and antitumor immunity, we employed murine models to conduct detailed mechanistic studies. Murine CD3⁺ T cells were purified from splenocytes and stimulated with anti-CD3/CD28 mAbs in the presence of graded concentrations of ponatinib to determine the dose at which similar effects on human CD8⁺ T cells were observed (Fig. 2a). Ponatinib increased CD8⁺ T cells with a naive phenotype in a dose-dependent manner at concentrations above 5 nM, whereas CD8⁺ T cell activation was almost completely suppressed upon treatment with 20 nM (Supplementary Fig. 7a, b). On the basis of these results, we used a concentration of 10 nM ponatinib in subsequent experiments. At this concentration, the total number of CD8⁺ T cells remained unchanged, but the proportion of these cells, particularly those with a naive phenotype, increased (Fig. 2b, c and Supplementary Fig. 7c, d). In contrast, the number of Treg cells and their expression of CTLA-4 decreased (Fig. 2d). Moreover, the reduction of activation marker expression (CTLA-4, GITR, and OX-40) was also observed when purified Treg cells were stimulated with ponatinib (Supplementary Fig. 7e, f).

Fig. 2. Ponatinib directly targets murine CD8⁺ T cells to induce a T_SCM cell phenotype.

a–d Murine CD3⁺ T cells purified from splenocytes were stimulated with anti-CD3/CD28 monoclonal antibodies (mAb) for 48 h along with dimethyl sulfoxide (DMSO; control) or ponatinib and analyzed by flow cytometry. a Experimental scheme. Created in BioRender. https://BioRender.com/cxg8twg. b t-SNE plots were generated to reduce all the phenotypic data to 2 dimensions (left), and DMSO (right, black) and ponatinib (right, red) treatments were applied. tSNE: t-distributed stochastic neighbor embedding (n = 3 mice per group). c Representative contour plots (CD62L and CD44) of CD8⁺ T cells treated with DMSO (left, black) or ponatinib (left, red) and frequencies of naive (CD44⁻CD62L⁺), central memory (T_CM: CD44⁺CD62L⁺), and effector memory (T_EM: CD44⁺CD62L⁻) subsets among CD8⁺ T cells (right) (n = 3 mice per group). d Absolute numbers of regulatory T (Treg) cells (CD4⁺Foxp3⁺CD25^high) and frequencies of CTLA-4⁺ cells among Treg cells (right) (n = 3 mice per group). e–j Murine naive CD8⁺ T cells purified from splenocytes by magnetic separation were stimulated with anti-CD3/CD28 mAbs for 48 h along with DMSO or ponatinib and subjected to RNA sequencing analysis (RNA-seq) and flow cytometry analysis. e Experimental scheme. Created in BioRender. https://BioRender.com/cxg8twg. f Gene set enrichment analysis (GSEA) plots of gene sets upregulated in naive versus effector CD8⁺ T cells (left) and in memory versus effector CD8⁺ T cells (right) for the ponatinib-treated group compared with the DMSO group (n = 3 mice per group). ES: enrichment score; FDR: false discovery rate. g Heatmap of the expression of CD8⁺ T cell phenotype related genes, such as naive/memory, activation/effector, exhaustion, and cytotoxicity genes (n = 3 mice per group). h Gating strategy for murine CD8⁺ T stem cell memory (T_SCM: CD8⁺CD44⁻CD62L⁺Sca-1⁺) cell (left) and absolute numbers of T_SCM cells (n = 7 mice per group) (right). i Frequencies of IL-7Rα⁺ naive subset in CD8⁺ T cells (n = 6 mice per group). j Expression of Tcf1 and Tox on CD8⁺ T cells (n = 6 mice per group) MFI, mean fluorescence intensity. Data are presented as the mean ± SEM (c, d, h–j). Significance was assessed using two-sided unpaired t tests (c, d, h–j). All experiments were performed independently at least two times with similar results.

We next addressed how CD8⁺ T cells with a naive phenotype increased after ponatinib treatment. Naive (CD44⁻CD62L⁺), central memory (T_CM, CD44⁺CD62L⁺), and T_EM (CD44⁺CD62L⁻) cell populations of murine CD8⁺ T cells were purified and stimulated with or without ponatinib (Supplementary Figs. 6b, 8a, b). The activation/differentiation of CD8⁺ T cells was markedly inhibited in the naive population after ponatinib treatment, whereas dedifferentiation from the T_CM and T_EM populations to the naive population was not observed (Supplementary Fig. 8c–e). These findings indicate that ponatinib restrained the activation/differentiation of naive CD8⁺ T cells, resulting in the generation of T_SCM cells.

Ponatinib drives T_SCM cell associated transcriptional and phenotypic changes in CD8⁺ T cells

The direct effects of ponatinib on CD8⁺ T cells prompted us to further investigate their detailed phenotypes. Murine naive CD8⁺ T cells purified from splenocytes were stimulated with anti-CD3/CD28 mAbs in the presence or absence of ponatinib for 48 h and subjected to RNA-seq and flow cytometry analysis (Fig. 2e). Principal component analysis (PCA) of the RNA-seq data revealed a distinct expression profile in the ponatinib group (Supplementary Fig. 9a), with 3589 differentially expressed genes (1797 upregulated, 1792 downregulated; Supplementary Fig. 9b). GSEA showed significant enrichment of naive and memory T cell signatures, rather than the effector T cell signature, in ponatinib-treated CD8⁺ T cells (Fig. 2f, and Supplementary Fig. 9c). Notably, naive and memory-associated genes were upregulated, along with some activation and cytotoxicity-related genes (Fig. 2g). Thus, ponatinib directly affects CD8⁺ T cells, skewing them toward those with both naive and memory phenotypes while maintaining some activation pathways, which is consistent with previously reported T_SCM cells^23–27.

The T_SCM cell characteristics of ponatinib-treated murine CD8⁺ T cells were further explored with flow cytometry analyses (Fig. 2e). The number of T_SCM cells (CD8⁺CD44⁻ CD62L⁺Sca-1⁺)^27,28 and the frequency of naive CD8⁺ T cells expressing T_SCM cell-associated markers significantly increased in the ponatinib group (Fig. 2h, i, and Supplementary Fig. 9d). In addition, CD8⁺ T cells exhibited significantly increased expression of Tcf1, an essential transcription factor for T_SCM cell formation^23–26. Moreover, the ratio of Eomes to T-bet, which is important for memory formation²⁹, was also significantly increased, whereas Tox expression, which is related to the exhaustion phenotype^29,30, was decreased in the ponatinib group (Fig. 2j and Supplementary Fig. 9e). Therefore, ponatinib treatment directly affects CD8⁺ T cells and endows both human and murine CD8⁺ T cells with a T_SCM cell phenotype.

Ponatinib augments antitumor immune responses by inducing tumor antigen-specific T_SCM cells in the tumor microenvironment (TME)

The increase in T_SCM cells, which are reportedly associated with durable antitumor activity in cancer immunotherapy^23–25, and the decrease in immunosuppressive Treg cells led us to examine the potential of ponatinib to augment antitumor immune responses through dual mechanisms. Thus, wild-type C57B/6 J mice bearing MC38 or B16F10 cells were orally administered ponatinib and/or treated with an anti-PD-1 mAb (Fig. 3a). Ponatinib significantly augmented antitumor responses and prolonged the survival of the mice in both models. In addition, the combination of ponatinib with an anti-PD-1 mAb exhibited a stronger antitumor effect than either treatment alone (Fig. 3b and Supplementary Fig. 10a–d). The antitumor effect of ponatinib was abrogated in immunocompromised (RAG2 knockout and nude) mice, indicating that the antitumor effect of ponatinib was T cell dependent (Fig. 3c and Supplementary Fig. 10e, f). To further clarify the effector population, CD4⁺ T cells or CD8⁺ T cells were depleted with an anti-CD4 mAb or an anti-CD8β mAb, respectively. The antitumor effect of ponatinib was completely abrogated by CD8⁺ T cell depletion but not by CD4⁺ T cell depletion, indicating that the antitumor efficacy of ponatinib relies on CD8⁺ T cells (Fig. 3d and Supplementary Fig. 11a, b). In fact, after CD4⁺ T cell depletion, tumor growth in the control group was delayed, probably due to the depletion of Treg cells. Notably, even upon CD4⁺ T cell depletion, where Treg cells might have been eliminated, tumor growth in the ponatinib-treated group was slightly delayed compared with that in the control group. These findings suggest that the antitumor efficacy of ponatinib is attributed not only to Treg cell depletion but also to additional mechanisms. We assessed the involvement of Treg cells in the antitumor activity of ponatinib by evaluating its antitumor effects in Foxp3-DTR mice under Treg cell-deficient conditions³¹. Ponatinib treatment still significantly augmented antitumor activity in MC38-bearing Foxp3-DTR mice (Fig. 3e and Supplementary Fig. 11c, d). Therefore, the antitumor efficacy of ponatinib depends mainly on CD8⁺ T cell responses.

Fig. 3. Ponatinib augments antitumor immune responses in a CD8⁺ T cell dependent manner.

a, b Wild-type C57BL/6 J mice were subcutaneously (s.c.) inoculated with MC38 cells or B16F10 cells and treated with ponatinib or 1% 2-hydroxypropyl-beta-cyclodextrin (Dex; control) and/or an anti-PD-1 monoclonal antibodies (mAb). a Experimental scheme. p.o., oral administration. Created in BioRender. https://BioRender.com/cxg8twg. b Tumor growth curves of MC38 tumor-bearing mice (left) and B16F10 tumor-bearing mice (right) (MC38: Dex, n = 9; ponatinib, n = 9; Dex + anti–PD-1 mAb, n = 8; ponatinib + anti-PD-1 mAb, n = 8; B16F10: Dex, n = 8; ponatinib, n = 9; Dex + anti–PD-1 mAb, n = 9; ponatinib + anti-PD-1 mAb, n = 8; all n-values represent the number of individual mice). c RAG2 knockout (KO) mice or nude mice were subcutaneously inoculated with MC38 cells and treated with Dex or ponatinib. Tumor growth curves of RAG2 KO mice (left; Dex, n = 8 mice; ponatinib, n = 7 mice) and nude mice (right; Dex, n = 10 mice; ponatinib, n = 11 mice). d Wild-type C57BL/6 J mice were subcutaneously inoculated with MC38 cells and treated with Dex or ponatinib. Some mice received intraperitoneal injections of anti-CD8β mAbs (CD8⁺ T cell depletion model) or anti-CD4 mAbs (CD4⁺ T cell depletion model) (n = 5 mice per group). Tumor growth curves of mice depleted of CD8⁺ T cells (left) or CD4⁺ T cells (right). Dex + Anti-CD8β mAb, mice treated with Dex and anti- CD8β mAb; Ponatinib + Anti-CD8β mAb, mice treated with ponatinib and anti-CD8β mAb; Dex + Anti-CD4 mAb, mice treated with Dex and anti- CD4 mAb; Ponatinib + Anti-CD4 mAb, mice treated with ponatinib and anti-CD4 mAb. e Wild-type or Foxp3-DTR C57BL/6 J mice were subcutaneously inoculated with MC38 cells and treated with Dex or ponatinib in addition to diphtheria toxin (DT) administration (regulatory T (Treg) cell depletion model). Tumor growth curves of the Treg cell depletion model (n = 4 mice per group). Dex-Foxp3-DTR: Foxp3-DTR mice treated with Dex. Ponaitnib-Foxp3-DTR: Foxp3-DTR mice treated with ponatinib. Dex-WT: wild-type mice treated with Dex. Ponatinib-WT: wild-type mice treated with ponatinib. The average tumor size of the groups on a certain day is shown as a dot (b–e). Data are presented as the mean ± SEM (b–e). Significance was assessed using two-way ANOVA with Tukey’s multiple comparison test (b–e). All experiments were performed independently at least two times with similar results.

We then examined the immunological landscape in the TME. For the MC38 tumor model, the mice were euthanized on Day 8, and tumor-infiltrating lymphocytes (TIL) were collected and subjected to flow cytometry (Fig. 4a). Ponatinib treatment significantly increased T_SCM cells, the frequency of CD8⁺ T cells expressing Tcf1 and Bcl-2, and the ratio of Eomes to T-bet and decreased Tox expression (Fig. 4b–e). Consequently, the ratio of T_SCM cells to Treg cells was significantly increased in the ponatinib group (Fig. 4f). Furthermore, CD8⁺ T cells that produced effector cytokines (IFN-γ, TNF, and IL-2) and a cytotoxic molecule (granzyme B) were significantly increased in the ponatinib group (Fig. 4g, h, and Supplementary Fig. 12a). Moreover, in the group treated with the combination of ponatinib and the anti-PD-1 mAb, T_SCM cells were increased, and Treg cells were decreased, as observed in the ponatinib group, compared to the control group, resulting in a significant increase in the ratio of T_SCM cells to Treg cells (Supplementary Fig. 12b–e). The combination treatment also significantly increased the ratio of CD8⁺ T cells to Treg cells (Supplementary Fig. 12f). Thus, ponatinib exerts a durable antitumor effect by enhancing the generation of T_SCM cells in the TME, and the combination of ponatinib with an anti-PD-1 mAb further augments antitumor efficacy. Similar results were also observed in another mouse model, B16F10 (Supplementary Fig. 12g–j).

Fig. 4. Ponatinib induces T_SCM cells in the TME and augments antitumor immunity.

a–h Wild-type C57BL/6 J mice were subcutaneously (s.c.) inoculated with MC38 cells and treated with ponatinib or 1% 2-hydroxypropyl-beta-cyclodextrin (Dex; control). On Day 8, the mice were euthanized, and the tumor-infiltrating lymphocytes (TIL) were collected for the flow cytometry analysis. a Experimental scheme. p.o., oral administration. Created in BioRender. https://BioRender.com/cxg8twg. b Absolute numbers (per 1 g of tumor) of CD8⁺ TILs with the T stem cell memory (T_SCM) phenotype (CD8⁺CD44⁻CD62L⁺Sca-1⁺) (n = 12 mice per group). c Frequencies of Tcf1⁺ and Bcl-2⁺ cells among CD8⁺ TILs (n = 8 mice per group). d Ratios of Eomes to T-bet in CD8⁺ TILs (n = 8 mice per group). MFI, mean fluorescence intensity. e Frequencies of Tox⁺ cells in CD8⁺ TILs (n = 6 mice per group). f Ratios of T_SCM cells to regulatory T (Treg; CD4⁺CD25^hiFoxp3^hi) cells (n = 12 mice per group). g Representative contour plots (IFN-γ and TNF) of CD8⁺ TILs (left, Dex in black; right, ponatinib in red). h Frequencies of TNF- and IFN-γ-, TNF- and IL-2-, IFN-γ- and granzyme B (GZMB)-producing cells among CD8⁺ TILs (n = 11 mice per group). i–l Wild-type C57BL/6 J mice were subcutaneously inoculated with MC38-OVA cells and treated with ponatinib or Dex. The mice were euthanized on Day 8 (early phase) or when the tumor volume reached 1000 mm³ (late phase), and the TILs were collected for the flow cytometry analysis. i Experimental scheme. Created in BioRender. https://BioRender.com/cxg8twg. j Representative contour plots; left, CD8 and OVA dextramer (total CD8⁺ TILs); middle, CD62L and CD44 (antigen-specific CD8⁺ TILs (CD8⁺OVA dextramers⁺)); right, CD62L and Sca-1 (antigen-specific naive CD8⁺ TILs, CD8⁺OVA dextramers⁺CD44⁻CD62L⁺) in the DMSO group (black) and ponatinib group (red). k Absolute numbers (per 1 g of tumor) of antigen-specific T_SCM cells (CD8⁺OVA dextramers⁺CD44⁻CD62L⁺Sca-1⁺) in the early phase (Dex, n = 7 mice; ponatinib, n = 8 mice). l Representative contour plots (CD8 and OVA dextramer) of effector CD8⁺ TILs (CD8⁺CD44⁺) in the Dex (black) and ponatinib groups (red) (left) and absolute numbers (per 1 g of tumor) of antigen-specific effector CD8⁺ TILs (CD8⁺CD44⁺OVA dextramers⁺) in the late phase (n = 6 mice per group). In box plots (b, f, h), the center line indicates the mean, the box represents ± SEM, and whiskers indicate the minimum and maximum values. In dot plots (c–e), horizontal lines indicate the mean. In bar graphs (k, l), bars represent the mean ± SEM. Significance was assessed using two-sided Mann–Whitney U-test (b–f, h, k, l). All experiments were performed independently at least two times with similar results.

We further asked whether the increased T_SCM cells in the TME were tumor antigen-specific CD8⁺ T cells using MC38–ovalbumin (OVA) tumors. Tumors were collected at two different time points, namely, the early phase (Day 8 after tumor inoculation) and the late phase (when the tumor volume reached 1000 mm³), and analyzed by flow cytometry (Fig. 4i). Tumor antigen-specific T_SCM cells were significantly increased in the early phase and differentiated into an effector phenotype in the late phase in the ponatinib group (Fig. 4j–l). These results indicate that the antitumor effects of ponatinib are achieved by the initial increase of tumor antigen-specific T_SCM cells, followed by the generation of persistent effector CD8⁺ T cells in the TME.

Ponatinib improves the antitumor efficacy of adoptive cell transfer

Adoptive cell transfer (ACT) is a promising approach in cancer immunotherapy. However, several limitations, including the exhaustion and dysfunction of CD8⁺ T cells in the TME, hinder its broader application³², especially in solid tumors. T_SCM cells play a key role in the antitumor efficacy of ACT due to their self-renewal capacity, multipotency, and longevity^4,5. Given that ponatinib increased T_SCM cells, we next explored whether ponatinib could augment the antitumor efficacy of ACT. CD8⁺ T cells from OVA specific TCR Tg mice (OT-I) were cultured with OVA_257–264 peptide-pulsed antigen-presenting cells (APC) with or without ponatinib for 48 h and then adoptively transferred into SIINFEKL-expressing B16F10 tumor-bearing mice, and tumor growth and the number of transferred cells in the TME were examined (Fig. 5a). Prior to transfer, T_SCM cells were significantly increased in the ponatinib group (Fig. 5b). Consistently, mice receiving ponatinib-treated CD8⁺ T cells exhibited stronger antitumor responses, accompanied by an increased number of transferred cells in the TME (Fig. 5c, d and Supplementary Fig. 13a, b).

Fig. 5. Ponatinib augments the antitumor efficacy of ACT.

a–d Antigen-presenting cells (APC, Thy1.2⁻ cells) were purified from the splenocytes of wild-type C57BL/6 J and pulsed with the OVA_257–264 peptide overnight (OVA_257–264 peptide-pulsed APC), then cultured with naive CD8⁺ T cells purified from the splenocytes of OT-1 C57BL/6 J mice for 48 h in the presence of dimethyl sulfoxide (DMSO; control) or ponatinib. Prior to transfer, the phenotype of CD8⁺ T cells was confirmed by flow cytometry. Subsequently, OT-1 CD8⁺ T cells (1 × 10⁶) were transferred into SIINFEKL-expressing B16F10 (B16F10-SIINFEKL) tumor-bearing mice. On Day 8 after adoptive cell transfer (ACT), the mice were euthanized, and the tumor-infiltrating lymphocytes (TIL) were collected for the flow cytometry analysis. a Experimental scheme. s.c., subcutaneous administration; i.v., intravenous administration. Created in BioRender. https://BioRender.com/zxkr16g. b Prior to transfer, OT-1 CD8⁺ T cells were analyzed by flow cytometry. Representative contour plots (left, CD62L and CD44; middle, CD62L and Sca-1) of CD8⁺ T cells and naive CD8⁺ T cells in the DMSO (black) and ponatinib groups (red), and frequencies of T stem cell memory (T_SCM) cells (CD8⁺CD44⁻CD62L⁺Sca-1⁺) among CD8⁺ T cells (right) (n = 8 samples per group). c Tumor growth curves of mice receiving ACT (n = 10 mice per group). d Representative contour plots [CD45.2 (donor-derived) and CD45.1 (recipient-derived)] of CD8⁺ TILs from the DMSO (left, black) and ponatinib (left, red) groups and absolute numbers (per 1 g of tumor) of donor-derived CD8⁺CD45.2⁺ cells (right) (n = 3 mice per group). e–i For systemic ponatinib administration, OT-1 CD8⁺ T cells were activated for 48 h with OVA_257–264 peptide–pulsed APCs (without ponatinib), and OT-1 CD8⁺ T cells (1 × 10⁶) were transferred into B16F10-SIINFEKL tumor-bearing mice. The mice were orally administered ponatinib or 1% 2-hydroxypropyl-beta-cyclodextrin (Dex; control) once daily starting 1 day before transfer and continuing for 14 consecutive days. On Day 8 after ACT, the mice were euthanized, and the TIL were collected for the flow cytometry analysis. e Experimental scheme. p.o., oral administration. Created in BioRender. https://BioRender.com/zxkr16g. f Tumor growth curves of mice treated with ACT (n = 10 mice per group). Dex with ACT, oral administration of Dex with ACT; Ponatinib with ACT, oral administration of ponatinib with ACT. g Absolute numbers (per 1 g of tumor) of donor-derived CD8⁺CD45.2⁺ T cells (n = 4 mice per group). h Frequencies of PD-1⁺, TIM-3⁺, LAG-3⁺, and PD-1⁺TIM-3⁺ cells among donor-derived CD8⁺CD45.2⁺ T cells (n = 8 mice per group). i TOX expression in donor-derived CD8⁺CD45.2⁺ T cells (n = 8 mice per group). MFI, mean fluorescence intensity. The average tumor size of the groups on a certain day is shown as a dot (c, f). Data are presented as the mean ± SEM (b–d, f–i). Significance was assessed using two-sided Mann–Whitney U-test (b, g–i), two-sided unpaired t test (d) and two-way ANOVA with Tukey’s multiple comparison test (c, f). All experiments were performed independently at least two times with similar results.

Next, we assessed whether the systemic administration of ponatinib could increase the effectiveness of ACT. OVA-specific CD8⁺ T cells were stimulated with cognate peptide-pulsed APCs (without ponatinib) and then transferred into SIINFEKL-expressing B16F10 tumor-bearing mice. These mice were treated with ponatinib beginning one day before transfer (Fig. 5e). While ponatinib and ACT treatment alone inhibited tumor growth, the combination of ponatinib and ACT resulted in more profound tumor growth inhibition than ponatinib or ACT treatment alone did (Fig. 5f, and Supplementary Fig. 13c–e). Flow cytometry analysis of the T cells in the TME on Day 8 after transfer revealed that systemic ponatinib treatment accelerated the expansion of donor-derived CD8⁺ T cells and prevented their exhaustion (Fig. 5g–i). Thus, ponatinib enhances ACT efficacy by enriching T_SCM cells during ex vivo preparation and by promoting the expansion of the transferred cells without exhausting them in the TME upon systemic administration.

Ponatinib inhibits the PI3K pathway and alters metabolic programming in CD8⁺ T cells

To gain insights into the mechanism(s) by which ponatinib induces T_SCM cells, we reanalyzed the RNA-seq data of ponatinib-treated CD8⁺ T cells. The mTOR signaling pathway was significantly enriched in CD8⁺ T cells in the control group. Consistent with the activation of the PI3K‒mTOR pathway, which is involved in the enhancement of glycolysis³³, glycolytic metabolism was increased in CD8⁺ T cells in the control group (Fig. 6a and Supplementary Fig. 14a). By contrast, ponatinib-treated CD8⁺ T cells showed increased expression of OXPHOS-related genes, reflecting the T_SCM cell phenotype (Fig. 6b). The metabolic characteristics of CD8⁺ T cells were further examined using a Seahorse-based bioenergetics analysis. CD8⁺ T cells in the ponatinib group exhibited a significantly higher oxygen consumption rate (OCR)/maximal respiration and spare respiratory capacity (SRC) and lower extracellular acidification rate (ECAR), which indicated improved metabolic fitness and a greater reliance of CD8⁺ T cells in the ponatinib group on mitochondrial respiration (Fig. 6c–e). Given the essential role of the PI3K pathway in T cell metabolism³³, the phosphorylation levels of molecules involved in the PI3K pathway were assessed in both human and murine CD8⁺ T cells. The phosphorylation levels of signal transduction molecules related to the PI3K pathway decreased in the ponatinib group, whereas the level of phosphorylated ERK increased, suggesting possible compensatory activation of the MAPK pathway (Fig. 6f–h Supplementary Fig. 14b–d). This was consistent with RNA-seq results showing upregulation of genes related to activated and cytotoxic phenotypes in the ponatinib group (Fig. 2g). These results indicate that ponatinib induces T_SCM cells by inhibiting the PI3K pathway and altering the metabolic pathway.

Fig. 6. Ponatinib regulates metabolic pathways and induces T_SCM cells through inhibition of the LCK–PI3K pathway.

a Gene set enrichment analysis (GSEA) plots of the HALLMARK_MTORC1_SIGNALING (top) and HALLMARK_GLYCOLYSIS (bottom) gene sets (n = 3 mice per group). ES, enrichment score; FDR, false discovery rate. b Heatmap of the expression of OXPHOS-related genes. (n = 3 mice per group). c–e Murine naive CD8⁺ T cells purified from splenocytes were stimulated with anti-CD3/CD28 monoclonal antibodies (mAb) for 48 h with dimethyl sulfoxide (DMSO; control) or ponatinib and subjected to a Seahorse Mito Stress assay. c Oxygen consumption rate (OCR) (n = 8 samples per group). FCCP, carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone; R + A, Rotenone and Antimycin. d Spare respiratory capacity (SRC) (n = 8 samples per group). e Ratios of the OCR to extracellular acidification rate (ECAR) (n = 8 samples per group). f–i Human CD8⁺ T cells purified from peripheral blood mononuclear cells (PBMC) of healthy individuals were stimulated with anti-CD3/CD28 mAbs for 24 h in the presence of DMSO, imatinib, or ponatinib, and subjected to Western blot and immunocytochemistry analyses. f Experimental scheme. Created in BioRender. https://BioRender.com/cxg8twg. g Representative Western blot images. β-actin shown was run in the same experiment on a separate gel/blot with the same samples and is provided as a sample processing control. Uncropped Western blot images are provided in the Source Data file. h Quantification of the expression of the indicated molecules by Western blot analysis (n = 3 donors per group). i CD8⁺ T cells were stained with anti-phosphorylated Src family members (pSrc) antibodies (green), anti-CD28 antibodies (red), and Hoechst (blue) and analyzed by confocal microscopy. Multiple random fields of view were photographed, and the frequency of cells showing colocalization of pSrc and CD28 was determined. Representative images with or without colocalization (left) and frequencies of colocalized CD8⁺ T cells (right) are shown (n = 13 fields from one donor). Similar results were independently observed in samples from an independent donor. j Human CD8⁺ T cells purified from the PBMCs of healthy individuals were stimulated with anti-CD3/CD28 mAbs along with DMSO, imatinib, or ponatinib for 9 days and analyzed by flow cytometry. Expression levels of TCF1 and FOXO1 (TCF1, n = 5 donors; FOXO1, n = 4 donors) are shown. MFI, mean fluorescence intensity. k–m Human CD8⁺ T cells purified from the PBMCs of healthy individuals were stimulated with anti-CD3/CD28 mAbs in the presence of graded doses of an LCK inhibitor (LCKi), imatinib, or ponatinib. Absolute numbers of T stem cell memory (T_SCM; CD8⁺CCR7⁺CD45RA⁺CD62L⁺CD95⁺) cells analyzed on Day 9, and the phosphorylation of LCK (Y394) and AKT assessed by flow cytometry after 24 h. The experimental scheme is shown in Supplementary Fig. 14e. k Cell numbers of T_SCM cells (n = 3 donors per group). l Representative histograms (left) and expression level of phosphorylated AKT (p-AKT) (right) (n = 4 donors per group). m Representative histograms (left) and expression level of phosphorylated LCK (p-LCK, Y394) (right) (n = 3 donors per group). In dot plots (i), horizontal lines indicate the mean. Data are presented as the mean ± SEM (c–e, h, j–m). Data in (h) were normalized to DMSO (set to 1.0), those in (k) to LCKi 0 nM (set to 1.0), and those in (l, m) to ponatinib (set to 1.0). Significance was assessed using two-sided Mann–Whitney U-test (d, e, i) and two-sided paired t test (h, j–m). All experiments were performed independently at least two times with similar results.

Ponatinib induces T_SCM cells through inhibition of the LCK–PI3K pathway

Although ponatinib cannot directly inhibit the PI3K pathway⁶, we observed its inhibition in both mouse and human CD8⁺ T cells. In fact, the phosphorylation of Src family members was significantly decreased in the ponatinib group. Thus, ponatinib likely strongly inhibited Src family members, resulting in decreased phosphorylation of downstream CD28 and subsequent inhibition of the PI3K pathway in CD8⁺ T cells³⁴, leading to T_SCM cell induction. Therefore, we investigated the colocalization of phosphorylated Src family kinases with CD28 in CD8⁺ T cells stimulated with anti-CD3/CD28 mAbs in the presence of imatinib or ponatinib, using immunofluorescence staining (Fig. 6f). The frequency of CD8⁺ T cells in which phosphorylated Src family members colocalized with CD28 was significantly reduced in the ponatinib group compared with the control or imatinib groups, indicating that ponatinib inhibits CD28 phosphorylation by Src family members, thereby suppressing the downstream PI3K pathway (Fig. 6i). Accordingly, ponatinib treatment significantly increased the expression of TCF1 and FOXO1 in CD8⁺ T cells (Fig. 6j). Both transcription factors are negatively regulated by AKT downstream of PI3K and play key roles in the induction of T_SCM cells^{23–26,35–37}.

Since LCK is an essential member of the Src family kinases in T cells³⁸, we also investigated whether LCK inhibition alone could induce T_SCM cell differentiation. CD8⁺ T cells were stimulated with anti-CD3/CD28 mAbs in the presence of graded concentrations of an LCK inhibitor (Supplementary Fig. 14e). The inhibition of LCK alone induced T_SCM cells; the phosphorylation of AKT was decreased at concentrations that induced T_SCM cells (Fig. 6k, l). However, insufficient LCK inhibition failed to suppress AKT phosphorylation (Fig. 6l, m), suggesting that LCK inhibition by imatinib may be insufficient to induce T_SCM cells. To further examine the role of LCK, we investigated whether restoring LCK activity could counteract the effects of ponatinib on the induction of T_SCM cells. Purified CD8⁺ T cells were stimulated with anti-CD3/CD28 mAbs in the presence or absence of ponatinib and treated with a Src family activator or transduced with constitutively active LCK to induce its overexpression (Supplementary Fig. 15a). Reactivation of LCK did not restore its catalytic activity. Since both methods activated LCK in an ATP-dependent manner³⁹, ponatinib completely abrogated LCK activation because of its strong ATP-competitive inhibitory function (Supplementary Fig. 15b–d). Therefore, we evaluated whether restoring AKT activation could counteract the effect of ponatinib on T_SCM cell induction (Supplementary Fig. 15a). Under the same stimulation conditions, CD8⁺ T cells were treated with an AKT activator or transduced with constitutively active AKT to induce its overexpression. Constitutive AKT activation resulted in increased phosphorylation of the downstream protein S6, confirming activation, whereas pharmacological AKT activation did not restore AKT phosphorylation or T_SCM cell induction (Supplementary Fig. 15e–h). Under these conditions, T_SCM cell induction was significantly inhibited, accompanied by reduced expression of TCF1 and FOXO1 (Supplementary Fig. 15i), indicating that the AKT pathway is a key mediator of ponatinib-driven T_SCM cell differentiation. Taken together, ponatinib inhibits LCK at an optimal intensity, attenuates the downstream PI3K pathway, including AKT, enhances the transcriptional activity of TCF7 and FOXO1, and consequently promotes the generation of T_SCM cells (summarized in Supplementary Fig. 16).

Ponatinib increases the generation of human CAR-T_SCM cells

We next asked whether ponatinib could also confer the T_SCM cell phenotype to CAR-T cells. T cells purified from human PBMCs were transduced with an anti-CD19 CAR-encoding lentiviral vector with a 4-1BB costimulatory domain and treated with DMSO (control-CAR-T cells) or ponatinib (ponatinib-CAR-T cells). Their phenotype and function were analyzed with scRNA-seq (Supplementary Data 1) and flow cytometry on day 8 (Fig. 7a and Supplementary Fig. 6c). CD8⁺ CAR-T (EGFR⁺CD8⁺) cells developed four clusters with notable differences between the ponatinib and control groups (Fig. 7b and Supplementary Fig. 17a). The ponatinib group comprised three clusters, Cytotoxic_1, Cytotoxic_2, and T_SCM, whereas the control group formed a single cluster: Memory. The expression of TCF7 and CCR7 was upregulated in the T_SCM cluster (Fig. 7c). Notably, the T_SCM cluster appeared exclusively in the ponatinib group (Fig. 7d and Supplementary Fig. 17b). While both the Memory and T_SCM clusters showed high expression of genes associated with a naive and memory phenotype, the T_SCM cluster presented a phenotype closer to that of naive T cells (Fig. 7e). By contrast, the Cytotoxic_1 and Cytotoxic_2 clusters were characterized by activation and cytotoxic markers, with distinct proliferation-related gene expression patterns (Supplementary Fig. 17c)⁴⁰. An analysis of a different gene set⁴¹ confirmed the phenotype of each cluster (Supplementary Fig. 17d, e). CAR-T_SCM cells are characterized by long-term survival and robust antitumor activity, which contrasts sharply with activated and cytotoxic subpopulations that provide limited efficacy in CAR-T cell therapy⁴². Compared with the Memory cluster in the control group, the T_SCM cluster exhibited reduced expression of genes associated with activation and exhaustion, but high expression of genes linked to stemness and a naive phenotype (Fig. 7f). Pathway enrichment analysis revealed upregulation of the WNT/β-catenin pathway, crucial for T_SCM cell development²⁶, in the T_SCM cluster. Conversely, pathways associated with apoptosis, TCR, CD28, and PI3K-mTORC1 signaling were downregulated (Fig. 7g and Supplementary Fig. 17f). Flow cytometry further confirmed the increased number and frequency of CAR-T_SCM cells; increased expression of TCF1 and FOXO1, which promote memory formation and inhibit exhaustion in CAR-T cells^4,36,37; and decreased expression of exhaustion markers on CD8⁺ CAR-T cells in the ponatinib group (Fig. 7h, i and Supplementary Fig. 18a, b). Thus, ponatinib optimally modulates TCR signaling, augmenting the transcriptional activity of FOXO1 and TCF7 and subsequently promoting the generation of CAR-T_SCM cells.

Fig. 7. Ponatinib promotes the development of CD8⁺ CAR-T_SCM cells in humans.

CD3⁺ T cells purified from the peripheral blood mononuclear cells (PBMC) of healthy individuals were activated with anti-CD3/CD28 monoclonal antibodies (mAb) and lentivirally transduced with the anti-CD19-BBz Chimeric antigen receptor (CAR) gene in the presence of dimethyl sulfoxide (DMSO; control) or ponatinib. The cells were analyzed by single-cell RNA sequence (scRNA-seq) and flow cytometry on Day 8 after transduction. For scRNA-seq, CD8⁺ CAR⁺ T cells (CD8⁺EGFR⁺) were enriched by fluorescence-assisted cell sorting. a Experimental scheme (top) and construct information for the anti-CD19-BBζ CAR gene (bottom). Created in BioRender. https://BioRender.com/cxg8twg (b) UMAP visualization of the scRNA-seq data for 28,724 cells. Each dot represents an individual cell. The cells are colored according to the group (top) and cluster (bottom) (n = 2 donors per group). c Expression of TCF7 and CCR7 in each cell (n = 2 donors per group). The colors represent the mean expression level. d Proportion of each cluster in the DMSO and ponatinib groups (n = 2 donors per group). e Dot plot showing the expression of genes related to the CD8⁺ T cell phenotype. The dot sizes represent the proportion of cells expressing each gene, and the dot colors indicate the mean expression levels of each gene. (n = 2 donors per group). f Volcano plot of differentially expressed genes between the T_SCM and Memory clusters. Molecules with significantly high and low expression in the T_SCM cluster compared with the Memory cluster are shown in red and blue, respectively (n = 2 donors per group). g Enriched MSigDB pathways in the T_SCM cluster identified by GSEApy (n = 2 donors per group). h Absolute numbers of CD8⁺ CAR-T_SCM cells generated from  2 × 10⁵ CD3⁺ T cells (n = 6 donors per group). i Representative histograms (left) and frequencies (right) of TCF1⁺ and FOXO1⁺ cells among CD8⁺ CAR-T cells in the DMSO (black) and ponatinib (red) groups (TCF1, n = 3 donors; FOXO1, n = 4 donors). scRNA-seq data (a–g) are provided in Supplementary Data 1. The FMO control is shown in gray, and the cutoff for positive cells is indicated by a dashed line (i). Data are presented as the mean ± SEM (h, i). Significance was assessed using two-sided Mann–Whitney U test (f) and two-sided paired t test (h, i).

Recent studies reported the influence of distinct CD4⁺ CAR-T cell subsets on therapeutic efficacy^40,43,44. Therefore, we also investigated the impact of ponatinib on CD4⁺ CAR-T cells. Ponatinib significantly reduced CAR-T cells with Treg cell phenotype (CAR-Treg cell), which reportedly attenuate the efficacy of CAR-T cell therapy⁴³, while increasing conventional CD4⁺ CAR-T cells with naive phenotype (CD4⁺Foxp3^⁻/lowCD45RA⁺) (Supplementary Fig. 18c, d). In addition, ponatinib-treated conventional CD4⁺ CAR-T cells exhibited increased cytokine production; Th2 cytokines, such as IL-4, which play a crucial role in CAR-T cell therapy^40,44, and the Th1 cytokine IFN-γ, were significantly upregulated (Supplementary Fig. 18e, f). Thus, ponatinib augments the therapeutic efficacy of CAR-T cells by promoting CAR-T_SCM cell generation, modulating CD4⁺ CAR-T cells, increasing IL-4 and IFN-γ production, and reducing CAR-Treg cells.

Ponatinib augments antitumor efficacy in humanized leukemia and solid tumor models

The enhanced generation of CAR-T_SCM cells prompted us to investigate the antitumor efficacy of ponatinib-CAR-T cells in a xenograft mouse model. Mice bearing Nalm-6 B-cell leukemia were treated with graded doses of ponatinib-CAR-T cells (Supplementary Fig. 19a). Ponatinib-CAR-T cells significantly inhibited tumor growth even after low-dose infusion (Supplementary Fig. 19b, c). Furthermore, when leukemia-bearing mice whose initial Nalm-6 tumors were completely eradicated by Day 28 after CAR-T cell transfer were rechallenged with a tenfold higher dose of Nalm-6 cells, ponatinib-CAR-T cells exhibited strong antitumor efficacy against the secondary challenge, indicating that ponatinib-CAR-T cells can continuously control the tumor upon rechallenge, a hallmark of memory T cells (Fig. 8a–c and Supplementary Fig. 19d). Consistently, ponatinib-CAR-T cells exhibited greater expansion than control-CAR-T cells; robust re-expansion was observed after rechallenge (Fig. 8d). In addition, peripheral ponatinib-CAR-T cells maintained a central memory phenotype (Fig. 8e, f and Supplementary Fig. 19e), suggesting that ponatinib-CAR-T cells maintain their T_SCM cell properties after in vivo infusion.

Fig. 8. Ponatinib augments human CAR-T cell efficacy in both leukemia and solid tumor models.

a–f Chimeric antigen receptor (CAR)-T cells generated in the presence of dimethyl sulfoxide (DMSO; control) or ponatinib were administered to Nalm-6-bearing NSG mice. Mice in which the initial tumors had been completely eliminated were rechallenged with a 10-fold higher dose of Nalm-6 cells on Day 28 after the first CAR-T cell administration. Peripheral blood was collected weekly from Day 7 until Day 42 to evaluate CAR-T cell engraftment, and a phenotypic analysis was performed on Day 14. a Experimental scheme. i.v., intravenous administration. Created in BioRender. https://BioRender.com/cxg8twg. b Bioluminescence imaging at the indicated time points (n = 5 mice per group). c Tumor burdens (total flux, photons/sec) are shown as a bar graph for the control and ponatinib groups on Day 28 (before rechallenge) and on Day 42 (after rechallenge) (DMSO, n = 11 mice; ponatinib n = 9 mice). d Absolute numbers of CAR-T cells (per 1 μL of blood) at the indicated time points (DMSO, n = 11 mice; ponatinib n = 9 mice). e Representative contour plots (CCR7 and CD45RA) of CD8⁺ CAR-T cells in the control (left, black) and ponatinib (left, red) groups, and frequencies of T central memory subset (T_CM: CCR7⁺CD45RA⁻) among CD8⁺ CAR-T cells (right) (n = 5 mice per group). f Representative histograms (left) and frequencies of TCF1⁺ cells among CD8⁺ CAR-T cells (right) in the DMSO (black) and ponatinib (red) groups (n = 5 mice per group). g–i The antitumor efficacy of ponatinib was evaluated in a solid tumor model. Human CD19-expressing A549 cells (hCD19-A549) were subcutaneously inoculated (s.c.) into NSG mice, and CAR-T cells were administered. Two experimental designs were tested: a preconditioning model, in which CAR-T cells were generated with DMSO or ponatinib, and a postinfusion treatment model, in which mice were administered CAR-T cells generated in the absence of ponatinib and subsequently received oral ponatinib or 1% 2-hydroxypropyl-β-cyclodextrin (Dex, control) starting on Day 3 after CAR-T cell infusion, following a 5-day-on, 2-day-off regimen for 4 weeks. For the postinfusion treatment model, tumor-infiltrating lymphocytes (TIL) were collected on Day 17 and analyzed by flow cytometry. g Experimental scheme. p.o., oral administration. Created in BioRender. https://BioRender.com/cxg8twg. h Tumor growth curves in the preconditioning model (n = 10 mice per group). i Tumor growth curve in the postinfusion treatment model (Dex with Mock, n = 3 mice; Ponatinib with Mock, n = 3 mice; Dex with CAR-T, n = 5 mice, Ponatinib with CAR-T, n = 5 mice). Dex with Mock, mice treated with the mock control and Dex; Ponatinib with Mock: mice treated with the mock control and ponatinib. Dex with CAR-T, mice treated with CAR-T cells and Dex; Ponatinib with CAR-T, mice treated with CAR-T cells and ponatinib. The FMO control is shown in gray, and the cutoff for positive cells is indicated by a dashed line (f). Data are presented as the mean ± SEM (c–f, h, i). Significance was assessed using two-sided Mann–Whitney U-test (c), two-way ANOVA with Tukey’s multiple comparison test (d, h, i) and two-sided unpaired t test (e, f). All experiments were performed independently at least two times with similar results.

We next assessed the impact of ponatinib on the efficacy of CAR-T cell therapy against solid tumors. Since Nalm-6 cells are highly sensitive to ponatinib, distinguishing the direct antitumor effects of ponatinib from the effects of CAR-T cells modulated by ponatinib is difficult. Conversely, A549 cells (a lung cancer cell line) are resistant to ponatinib, which allowed us to evaluate the effects of ponatinib on CAR-T cells in vivo without direct antitumor effects (Supplementary Fig. 20a). Therefore, we employed a humanized CD19-expressing A549 tumor model to assess the in vivo and ex vivo effects of ponatinib on CAR-T cells, using two experimental models: a preconditioning model, in which CAR-T cells were generated with or without ponatinib, and a postinfusion treatment model, in which CAR-T cells were generated without ponatinib, but the mice received ponatinib orally for 4 weeks (starting on Day 3 postinfusion and following a 5-day-on, 2-day-off schedule) (Fig. 8g). In the preconditioning model, ponatinib-CAR-T cells showed superior antitumor activity consistent with the results from leukemia models (Fig. 8h and Supplementary Fig. 20b, c). In the postinfusion treatment model, ponatinib treatment after CAR-T cell administration significantly augmented the antitumor effects of CAR-T cells (Fig. 8i and Supplementary Fig. 20d, e). TIL analysis (Day 17) revealed that ponatinib treatment increased CAR-T cell proliferation and reduced the expression of exhaustion markers and TOX by CD8⁺ CAR-T cells in the postinfusion treatment model (Supplementary Fig. 20f–h). Therefore, ponatinib augments the antitumor efficacy of CAR-T cells in both leukemia and solid tumor models by promoting the generation of CAR-T_SCM cells during their ex vivo generation. In addition, ponatinib treatment modulates the phenotype of CAR-T cells, supporting their durable persistence in vivo by reducing exhaustion.

Stable and sustained kinase inhibition promotes T_SCM cell induction

The prevention of T cell exhaustion by ponatinib is consistent with previous reports showing that dasatinib prevents CAR-T cell exhaustion by inhibiting Src family members and improving antitumor activity⁴⁵. Since both ponatinib and dasatinib are potent inhibitors of Src family kinases, we investigated whether dasatinib could also induce T_SCM cells, similar to ponatinib. CD8⁺ T cells were purified from PBMCs and stimulated with anti-CD3/CD28 mAbs in the presence of various concentrations of dasatinib. Dasatinib induced T_SCM cells at 20–30 nM, with efficacy similar to that observed with ponatinib (Supplementary Fig. 21a, b). However, considering the pharmacokinetic (PK) profile of dasatinib⁴⁶, exposure to this concentration range is limited in vivo, which may hinder its ability to induce T_SCM cells (Supplementary Fig. 21c). To address this issue, we conducted PK-mimicking experiments in which CD8⁺ T cells were stimulated with anti-CD3/CD28 mAbs at drug concentrations designed to recapitulate key features of the in vivo kinetics of ponatinib or dasatinib (Fig. 9a). Under these clinically relevant conditions, ponatinib consistently induced T_SCM cells and maintained stable and sustained inhibition of LCK and AKT phosphorylation. In contrast, dasatinib did not maintain LCK and AKT inhibition, resulting in less effective T_SCM cell induction (Fig. 9b, c). Furthermore, we assessed the effects of the PK dynamics of both agents on tumor cell cytotoxicity using K562 cells (Fig. 9a). Both ponatinib and dasatinib promoted apoptosis and reduced viability in K562 cells (Supplementary Fig. 21d). These findings highlight the fundamental difference in the response of tumor cells and T cells to kinase inhibitors: tumor cells, which rely heavily on driver kinases, can be eliminated through continuous or intermittent inhibition, whereas modulating T cell differentiation and promoting T_SCM cell induction are preferentially observed with stable and sustained suppression of TCR signaling through target kinase inhibition, which is more likely to be achieved in vivo by ponatinib.

Fig. 9. Stable and sustained kinase inhibition promotes T_SCM cell induction.

a–c Pharmacokinetic (PK)-mimicking in vitro experiments. Human CD8⁺ T cells purified from the peripheral blood mononuclear cells (PBMC) were stimulated with anti-CD3/CD28 monoclonal antibodies (mAb) and exposed to dimethyl sulfoxide (DMSO; control), ponatinib, or dasatinib at drug concentrations mimicking those observed on the basis of the in vivo PK of each drug, and the drug exposure was repeated daily for 5 days. K562 cells were treated under the same PK-mimicking conditions for 3 days. The PK-mimicking exposure schedules are in vitro approximations of clinically observed exposure patterns. a Experimental scheme. Created in BioRender. https://BioRender.com/cxg8twg. b Time-course analysis of phosphorylated LCK (p-LCK, Y394) and AKT (p-AKT) levels was performed every 12 h by flow cytometry (black, DMSO; red, ponatinib; green, dasatinib) (n = 3 donors per group). c Absolute numbers of T stem cell memory (T_SCM, CD8⁺CCR7⁺CD45RA⁺CD62L⁺CD95⁺) cells on Day 6 (n = 3 donors per group). d–g Wild-type C57BL/6 J mice were subcutaneously (s.c.) inoculated with MC38 cells and orally administered 1% 2-hydroxypropyl-β-cyclodextrin (Dex, control), ponatinib (3 mg/kg, once daily), continuous_dasatinib (50 mg/kg, once daily), or pulsed_dasatinib (50 mg/kg, twice daily, 3 days on/4 days off). The mice were euthanized on Day 8, and tumor-infiltrating lymphocytes (TIL) were collected for flow cytometry. d Experimental scheme. Created in BioRender. https://BioRender.com/cxg8twg. e Tumor growth curves (n = 9 mice per group). f Absolute numbers of T_SCM cells (CD8⁺CD44⁻CD62L⁺Sca-1⁺) (per 1 g of tumor) (n = 5 mice per group). g Ratios of T_SCM cells to regulatory T (Treg) cells (left) and CD8⁺ T cells to Treg cells (right) among TILs (n = 5 mice per group). h, i Wild-type C57BL/6 J mice were subcutaneously inoculated with MC38 cells and orally administered Dex, continuous_dasatinib, or pulsed_dasatinib, with or without an anti-PD-1 mAb. h Experimental scheme. Created in BioRender. https://BioRender.com/cxg8twg. i Tumor growth curves (Dex, n = 9; Continuous_Dasatinib, n = 9; Pulsed_Dasatinib, n = 10; Dex + anti-PD-1 mAb, n = 10; Continuous_Dasatinib + anti-PD-1 mAb, n = 10; Pulsed_Dasatinib + anti-PD-1 mAb, n = 10; all n-values represent the number of individual mice). Dex + anti-PD-1 mAb, mice treated with Dex and anti-PD-1 mAb; Continuous_Dasatinib + anti-PD-1 mAb, mice treated with continuous_dasatinib and anti-PD-1 mAb; Pulsed_Dasatinib + anti-PD-1 mAb, mice treated with pulsed_dasatinib and anti-PD-1 mAb. The average tumor size of the groups on a certain day is shown as a dot (e, i). Data are presented as the mean ± SEM (b, c, e–g, i). Significance was assessed using two-sided paired t test (b, c), two-way ANOVA with Tukey’s multiple comparison test (e, i), and two-sided Mann–Whitney U-test (f, g). All experiments were performed independently at least two times with similar results.

We next investigated the in vivo antitumor efficacy of dasatinib using tumor-bearing mouse models. Wild-type C57BL/6 J mice bearing MC38 or B16F10 tumors were orally administered ponatinib or dasatinib. Dasatinib was applied via two regimens: continuous daily dosing and a pulsed high-dose regimen (three days on and four days off), as previously described⁴⁵ (Fig. 9d). Ponatinib treatment significantly inhibited tumor growth in both tumor models, whereas dasatinib did not control tumor growth under either treatment regimen (Fig. 9e and Supplementary Fig. 22a–e). In the MC38 tumor model, mice were euthanized on Day 8, and TILs were collected and subjected to flow cytometry analysis (Fig. 9d). Unlike ponatinib, dasatinib did not induce T_SCM cells or increase the ratios of T_SCM cells to Treg cells and CD8⁺ T cells to Treg cells under either regimen (Fig. 9f, g). Moreover, we examined the antitumor efficacy of combining dasatinib with an anti-PD-1 mAb in the MC38 model (Fig. 9h). Dasatinib did not improve the therapeutic efficacy of PD-1 blockade under either regimen (Fig. 9i and Supplementary Fig. 22f, g). Consistent results were obtained even when dasatinib was administered at a lower dose (Supplementary Fig. 23a–g). Thus, even when the strength of TCR signaling was modulated by altering the treatment conditions, dasatinib consistently did not induce T_SCM cells in the TME or improve antitumor efficacy. Taken together, under these conditions, dasatinib does not achieve stable and sustained inhibition of Src family kinases in vivo and shows limited induction of T_SCM cells and effective antitumor immune responses, with no additional benefit in combination with PD-1 blockade. Thus, stable and sustained inhibition of Src family kinases is a critical feature of T_SCM cell induction in vivo, which is more likely to be achieved by ponatinib.

Ponatinib treatment increases T_SCM cells in patients with CML

Finally, PBMCs from patients with CML who were treated with imatinib, dasatinib, or ponatinib and achieved a deep molecular response were investigated (summarized in Supplementary Table 1). PBMCs were collected at trough drug concentrations and analyzed by flow cytometry (Fig. 10a). Treg cells significantly decreased, and T_SCM cells were significantly increased only in the ponatinib group compared with those in the imatinib group (Fig. 10b–d). Consequently, the ratio of T_SCM cells to Treg cells and CD8⁺ T cells to Treg cells were significantly higher specifically in the ponatinib group (Fig. 10e, f). No significant differences were observed between the dasatinib and imatinib groups. Therefore, ponatinib promotes T_SCM cells in patients with CML, highlighting the immunomodulatory effects of ponatinib.

Fig. 10. Ponatinib increases T_SCM cells in patients with CML.

Peripheral blood mononuclear cells (PBMC) from patients with chronic myeloid leukemia (CML) treated with either imatinib, dasatinib, or ponatinib were collected at trough levels and analyzed by flow cytometry (imatinib, n = 6 samples; dasatinib, n = 4 samples; ponatinib, n = 5 samples). a Experimental scheme. Created in BioRender. https://BioRender.com/cxg8twg. b Representative contour plots (left, CCR7 and CD45RA; right, CD95 and CD62L) of CD8⁺ T cells and naive CD8⁺ T cells (CD8⁺CD45RA⁺CCR7⁺), respectively, of the imatinib (blue), dasatinib (green), and ponatinib (red) groups. c Frequencies of T_SCM cells (CCR7⁺CD45RA⁺CD62L⁺CD95⁺) among CD3⁺ T cells. d Representative contour plots (CD45RA and FOXP3) of CD4⁺ T cells in the imatinib (blue), dasatinib (green), and ponatinib (red) groups (left) and frequencies of effector Treg cells (eTreg, CD4⁺CD45RA⁻FOXP3^hi) among CD3⁺ T cells (right). e Ratios of T_SCM cells to Treg cells. f Ratios of CD8⁺ T cells to Treg cells. Data are presented as the mean ± SEM (c–f). Significance was assessed using two-sided Mann–Whitney U-test (c–f).

Discussion

In this study, we show the immunomodulatory effects of ponatinib, which not only affects Treg cells like imatinib but also directly modulates CD8⁺ T cells. Compared with imatinib and dasatinib, ponatinib provided stable and sustained suppression of LCK, thereby directly controlling CD8⁺ T cell activation/differentiation to induce the T_SCM cell phenotype. The optimal inhibition of LCK by ponatinib downregulated the PI3K–AKT pathway and increased the expression of TCF7 and FOXO1, which are known to be involved in the development of T_SCM cells. Ponatinib promoted memory formation and mitigated T cell exhaustion. Consequently, ponatinib treatment skewed antitumor immune responses toward the T_SCM cell phenotype, resulting in augmented antitumor efficacy in preclinical models. Indeed, the combination of ponatinib with PD-1 blockade further increased antitumor efficacy and controlled tumor growth, including a PD-1 blockade-resistant tumor model⁴⁷.

Cancers with an inflamed TME (so-called immunologically hot tumors) consist of abundant effector T cells and often respond to ICIs, whereas cancers with a noninflamed TME (immunologically cold tumors) have limited immune cell infiltration and hardly respond to ICIs⁴⁷. In addition, Treg cells have been reported to be involved in the resistance to ICIs in various cancers²¹. PD-1 blockade exhibited a strong antitumor effect in combination with ponatinib treatment on B16F10 tumors, which are known to harbor a noninflamed TME⁴⁷. While combination therapy with imatinib or dasatinib and ipilimumab (an anti-CTLA-4 antibody) failed to show synergistic antitumor effects in patients with solid tumors in a phase I trial^48,49, this was likely due to inadequate, such as intermittent, control of LCK, which plays a key role in modulating T cell activation/differentiation. Given its potential to increase T_SCM cells and decrease Treg cells, ponatinib could become a therapeutic option.

While CAR-T cell therapy has shown substantial antitumor effects on hematologic malignancies, two key concerns have been raised: the duration of the antitumor effect and the suppression by Treg cells transduced with CAR-T constructs⁴³. For the former issue, accumulating evidence reveals that CAR-T_SCM cells are critical for durable antitumor effects. Notably, ponatinib has dual advantages in this regard: ponatinib facilitated the induction of CAR-T_SCM cells during CAR-T cell generation. In addition, ponatinib administration after CAR-T cell infusion increased the number of CAR-T cells in the TME and mitigated CAR-T cell exhaustion. For the latter issue, ponatinib inhibited Treg cell proliferation and induced Treg cell apoptosis. Therefore, ponatinib may provide multiple therapeutic benefits, including its use in the CAR-T cell generation process and in therapeutic processes in combination with conventional CAR-T cell therapy. In addition, it has been recently reported that Type II immunity is important for cancer therapy, particularly for durable antitumor effects⁵⁰. In CAR-T cell therapy, IL-4, a Th2 cytokine, reportedly plays an important role^40,44. In fact, during in vitro culture for CAR-T cell generation, CD4⁺ T cells were also transduced with the CAR-T cell construct. Since ponatinib controls the activation/differentiation of CD4⁺ T cells, CD4⁺ T cells with the potential to produce multiple cytokines, including IL-4, are induced, which may also contribute to the long-term functionality and memory formation of CD8⁺ T cells.

We demonstrated the potential of ponatinib as an immunomodulatory agent that can augment T_SCM cell induction and antitumor immunity. However, several safety issues need to be considered in clinical settings because ponatinib is among the most cardiotoxic FDA-approved agents⁵¹. In PBMCs from patients, T_SCM cell induction was confirmed even at an oral dosage of 15 mg/day, which has been reported to reduce the risk of cardiotoxicity⁵². However, the possibility of additive cardiotoxicity remains a critical concern when combined with CAR-T cell therapy, which is associated with cardiovascular risks⁵³. Furthermore, CAR-T cell recipients are highly susceptible to infections⁵⁴. While imatinib has been shown to impair memory CD8⁺ T cell expansion without affecting primary viral control⁵⁵, the effects of ponatinib on immune responses to infections remain unknown. Therefore, although ponatinib strongly increases T_SCM cell induction, cardiotoxicity and the risk of infection must be carefully monitored in clinical applications.

Stable and sustained inhibition of the target kinase is associated with more efficient T_SCM cell induction, whereas intermittent kinase inhibition achieves cytotoxic effects in tumor cells that are heavily dependent on driver kinases. Our findings suggest that pharmacokinetics is an important consideration in a clinical context for modulating effector T cell activation/differentiation—including efficient T_SCM cell induction—thereby augmenting antitumor immunity. Furthermore, our data reconcile the differences in the in vitro and in vivo effects of kinase inhibitors on T cell modulation and provide an important perspective for their clinical application as T cell modulators.

In conclusion, we demonstrate that ponatinib effectively induces T_SCM cells and augments antitumor immunity, thereby improving the efficacy of both ICI therapy and CAR-T cell therapy. In addition, our findings provide mechanistic insight into how kinase inhibition modulates T cell activation/differentiation, establishing a framework for employing the immunomodulatory effects of molecularly targeted reagents in clinical settings.

Methods

Sex as a biological variable

Female mice were used in this study. For the B16F10 tumor experiments in B6.Cg-Rag2^tm1.1Cgn/J mice, we used mice of both sexes (3 females and 3 males per group). While it has not been confirmed whether the results are relevant to male mice, both women and men were included in our study cohorts of healthy individuals (females, n = 4; males, n = 5) and patients with CML (females, n = 9; males, n = 6), suggesting that our results are applicable to both females and males.

Patients and samples

Peripheral blood was obtained from patients with CML treated at Akita University Hospital and Nagoya University Hospital from August 2023 to October 2025 (summarized in Supplementary Table 1). PBMCs were isolated by density gradient centrifugation with Ficoll‒Paque (GE Healthcare, Chicago, IL). All donors provided written informed consent before sampling, according to the Declaration of Helsinki. This study was approved by the Institutional Review Boards of Nagoya University (#32350), Akita University (#3178), and the National Cancer Center Japan (#2015-300).

Cell lines and reagents

The MC38 mouse colon cancer cell line was purchased from Kerafast (#ENH204; Boston, MA). MC38 cells stably expressing ovalbumin (MC38-OVA) were generated in-house. The B16F10 mouse melanoma cell line, the Nalm-6 human CD19⁺ B-cell leukemia cell line, the A549 human lung cancer cell line and the K562 human CML cell line were purchased from the American Type Culture Collection (ATCC) (#CRL-6475, #CRL-3273, #CCL-185, and #CCL-243, respectively; Manassas, VA). B16F10 cells stably expressing the OVA-derived SIINFEKL peptide (B16F10-SIINFEKL) were generated in-house. Nalm-6 and A549 cells were transduced with a lentiviral vector encoding firefly luciferase and green fluorescent protein (FF-GFP). MC38, MC38-OVA, B16F10, and B16F10-SIINFEKL cells were maintained in high-glucose Dulbecco’s modified Eagle’s medium (DMEM) (Fujifilm Wako Pure Chemical Corporation, #044-29765; Osaka, Japan). Nalm-6, A549, and K562 cells were maintained in Roswell Park Memorial Institute (RPMI)-1640 medium (Fujifilm Wako Pure Chemical Corporation, #189-02025). These media were supplemented with 10% fetal bovine serum (FBS) (Merck, Darmstadt, Germany).

The anti-mouse PD-1 mAb (clone RMPI-14) was purchased from BioLegend (#114101). The anti-mouse CD4 mAb (clone GK1.5, #BE0003-1) and anti-mouse CD8β mAb (clone 53–5.8, #BE0223) were purchased from BioXCell (Lebanon, NH). For in vitro and ex vivo assays, imatinib, dasatinib, ponatinib, the LCK inhibitor (RK 24466), and the AKT activator (SC79) were obtained from Selleck Chemicals (#S2475, #S1021, #S1490, #S0020, and #S7863; Houston, TX), and the Src family activator (YEEI phosphopeptide) was obtained from Santa Cruz Biotechnology (#sc-3052; Dallas, TX). For in vivo models, dasatinib and ponatinib were obtained from MedChemExpress (#HY-10181 and #HY-12047, Lexington, MA). Both drugs were suspended in water containing 1% 2-hydroxypropyl-beta-cyclodextrin (FUJIFILM Wako Pure Chemical, #324-84233) and freshly prepared before each oral gavage.

Culture of human PBMCs and T cells

Cells were cultured in the presence of DMSO (control), imatinib (2.5 μM), ponatinib, or dasatinib. In in vitro experiments using ponatinib and dasatinib, multiple concentrations were examined to evaluate dose-dependent effects. CD8⁺ T cells and Treg cells (CD4⁺CD25⁺) were purified from the PBMCs of healthy individuals using a CD8 MicroBeads (Miltenyi Biotec, #130-045-201, Bergisch Gladbach, Germany) and a CD4⁺ CD25⁺ Regulatory T Cell Isolation Kit (Miltenyi Biotec, #130-091-301), respectively, with an autoMACS separator (Miltenyi Biotec) according to the manufacturers’ instructions. Naive CD8⁺ T cells (CD8⁺CD45RA⁺CCR7⁺) and CD8⁺ T_EM cells (CD8⁺CD45RA⁻CCR7⁻) were purified from the PBMCs of healthy individuals with a FACSDiscover™ S8 cell sorter (BD Biosciences) (Supplementary Fig. 6a). The purity of each phenotype was confirmed to be > 95% (Supplementary Fig. 5d). Purified T cells were stimulated with Dynabeads™ Human T-Activator CD3/CD28 (Thermo Fisher Scientific, #11131D) at a 1:1 cell-to-bead ratio in RPMI-1640 supplemented with 10% FBS and 4 mM L-glutamine (Thermo Fisher Scientific, #25030-081). Recombinant human IL-2 (Roche, #10799068001, Basel, Switzerland) was added at 30 U/mL for PBMCs and CD8⁺ T cells, or 300 U/mL for Treg cells. For proliferation assays, purified CD8⁺ T cells and Treg cells were stained with 5 μM CellTrace CFSE (Thermo Fisher Scientific, #34554) in a 37 °C CO₂ incubator for 10 min. Subsequently, 2 × 10⁵ CFSE-labeled cells were stimulated with Dynabeads Human T-Activator CD3/CD28 at a 1:1 cell-to-bead ratio in the presence of the indicated concentrations of ponatinib. After 4 days, the CFSE dilution was analyzed to assess T cell proliferation using a FACSymphony A3 (BD Biosciences, San Jose, CA) and FlowJo v.10.6.1 (BD Biosciences).

Flow cytometry assays

For flow cytometry analysis, single-cell suspensions were prepared as follows. For TIL analysis, tumor tissues were minced and treated with Tumor & Tissue Dissociation Reagent (TTDR) (BD Biosciences, #661563)⁵⁶. Cells were washed with PBS (Nacalai, #14249-24, Kyoto, Japan) containing 2% FBS and stained with the indicated antibodies and fixable viability dye. After staining for cell surface markers, the cells were intracellularly stained with antibodies with a Foxp3/Transcription Factor Staining Buffer Set according to the manufacturer’s instructions (Thermo Fisher Scientific, #00-5523-00). Dextramer staining with MHC Dextramer H-2Kb/SIINFEKL/PE (ImmuDEX, #JDO2163, Virum, Denmark) was performed according to the manufacturer’s instructions.

Intracellular cytokine staining was performed with 50 ng/ml phorbol 12-myristate 13-acetate (PMA; Sigma–Aldrich; #P8139, Burlington, MA), 750 ng/ml ionomycin (Sigma–Aldrich; #I0634), and 1 × brefeldin A solution (BioLegend, #420601, San Diego, CA). A BD Cytofix/Cytoperm Fixation/Permeabilization Solution Kit (BD Biosciences, #554714) was used for the fixation and permeabilization of the cells. After washing, the cells were analyzed with an LSRFortessa or a FACSymphony A3 and FlowJo ver. 10.6.1 software. The antibodies used for cell staining are summarized in Supplementary Data 2. The dilution of the staining antibodies was performed according to the manufacturer’s instructions. CAR-T cells were counted with CountBright™ Absolute Counting Beads (Invitrogen, #C36950, Carlsbad, CA).

Culture of mouse T cells

An anti-mouse CD3ε antibody (10 µg/mL: BD Biosciences, #557306) was used to coat a flat-bottom 96-well plate at 4 °C overnight. CD3⁺ T cells, naive CD8⁺ T cells (CD8⁺CD44⁻CD62L⁺) and Treg cells (CD4⁺CD25⁺) were purified from the splenocytes of C57BL/6 J mice with an EasySep™ Mouse T Cell Isolation Kit (STEMCELL Technologies, #19851, Vancouver, BC), a MojoSort™ Mouse CD8 Naive T Cell Isolation Kit (BioLegend, #480044), or a CD4⁺CD25⁺ Regulatory T Cell Isolation Kit (Miltenyi Biotec, #130-091-041). CD8⁺ T naive cells (CD8⁺CD44⁻CD62L⁺), CD8⁺ T_CM cells (CD8⁺CD44⁺CD62L⁺), and CD8⁺ T_EM cells (CD8⁺CD44⁺CD62L⁻) were purified from the splenocytes of wild-type C57BL/6 J mice with a BD FACSAria™ Fusion cell sorter (Supplementary Fig. 6b). The purity of each phenotype was confirmed to be > 95% (Supplementary Fig. 8b). The purified cells were subsequently seeded into a flat bottom 96-well plate. An anti-mouse CD28 antibody (0.5 µg/mL, BD Biosciences, #553295) and recombinant mouse IL-2 (30 U/ml, R&D System, #402-ML, Minneapolis, MN) were added to the culture media. After 2 h, the cells were collected for Western blotting, and after 48 h, the cells were collected and subjected to flow cytometry, bulk RNA-seq, and Seahorse assays.

Bulk RNA-seq analysis

Murine naive CD8⁺ T cells purified from splenocytes were stimulated with anti-CD3/CD28 mAbs in the presence of DMSO or ponatinib for 48 h. RNA was then collected to analyze changes in immune-related molecules in response to ponatinib treatment. Total RNA was extracted using an RNeasy® Plus Micro Kit (Qiagen, #74034, Hilden, Germany) according to the manufacturer’s instructions. Paired-end RNA sequencing was performed with a NovaSeq 6000 instrument (Illumina, San Diego, CA), and the data were processed using the Galaxy web platform⁵⁷, R v.4.1.1 (R Foundation for Statistical Computing, Vienna, Austria) and Python v.3.10.12. Raw FASTQ files were processed using fastp v.0.19.5⁵⁸, and transcript abundance was quantified using Salmon v.1.5.1⁵⁹. Differentially expressed genes were determined using DESeq2 v.1.22.1⁶⁰. GSEA v.4.3.2⁶¹ was performed according to the instructions. The annotated GTF files (GENCODE vM27) were obtained from https://www.gencodegenes.org.

Animal models

C57BL/6 J mice (#000664), BALB/c nude mice (#000651), B6.Cg-Rag2^tm1.1Cgn/J (RAG2 knockout) mice (#008448), B6.129(Cg)-Foxp3^{tm3(Hbegf/GFP)Ayr}/J (Foxp3-DTR) mice (#016958), C57BL/6-^{Tg(TcraTcrb)1100Mjb}/J (OT-1) mice (#003831), and NOD.Cg-Prkdc^scid Il2rg^tm1Wjl/SzJ (NSG) mice (#005557) were purchased from Jackson Laboratory (Bar Harbor, ME). CD45.1 C57BL/6 J mice were purchased from Sankyo Labo Service Corporation (Tokyo, Japan). Female mice were used unless otherwise specified; for B16F10 tumor experiments in B6.Cg-Rag2^tm1.1Cgn/J mice, mice of both sexes were used (3 females and 3 males per group). All the mice were 6 to 10 weeks of age. Germ-free (GF) NSG and nude mice were purchased from The Jackson Laboratory and maintained in gnotobiotic facilities. Other mice were housed under specific pathogen-free (SPF) conditions. Both GF and SPF mice were provided with standard food, given free access to hypochlorous weak-acid water, and maintained under identical environmental conditions with a 12-12 h light-dark cycle (lights on at 8:00), a controlled temperature of 22 °C (20–26 °C), and humidity of 45% (40–60%). Experimental and control animals were housed in separate cages.

For in vivo experiments, the mice were subcutaneously inoculated with the indicated tumor cells (1 × 10⁶) in the right flank (Day 0). The mice were randomized to each group prior to the start of treatment. Ponatinib or dasatinib was administered orally (p.o.) starting on Day 3. Ponatinib was given at 3 mg/kg once daily for 14 consecutive days. Dasatinib was administered either at 50 mg/kg once daily for 14 consecutive days (continuous regimen) or at 50 mg/kg twice daily for 3 consecutive days followed by 4 days off (pulsed regimen)⁴⁵, and this cycle was repeated throughout the 14-day treatment period. A low dose of dasatinib (4 mg/kg) was also investigated using the same continuous regimen. The anti-PD-1 mAb was administered on Days 8, 11, and 14 to the MC38 model and on Days 5, 8, and 11 to the B16F10 model at 200 µg per mouse. Antibodies for depleting CD4⁺ T cells or CD8⁺ T cells were injected intraperitoneally on days -1, 1, 8, and 15 at 100 µg per mouse. For Treg cell depletion, diphtheria toxin solution (FUJIFILM Wako Pure Chemical, #048-34371) was injected intraperitoneally at a dose of 1 μg per mouse starting on Day 2 and administered twice a week for a total of four doses. The mice were monitored twice a week and euthanized upon exhibiting 1 of the following symptoms: >20% loss of initial body weight, pronounced lethargy, severe diarrhea, hunched posture, severe dermatitis, and a tumor exceeding 20 mm in maximum diameter. Euthanasia was performed by carbon dioxide inhalation or cervical dislocation under deep anesthesia, in accordance with institutional guidelines. The tumor volume was calculated as length × width²× 0.5. Animal care and experiments were conducted according to the guidelines of the Animal Committee of Nagoya University and the National Cancer Center after being approved by the Ethics Review Committee for Animal Experimentation of Nagoya University and the National Cancer Center Japan. All in vivo experiments were performed at least twice. The maximum tumor size permitted by institutional guidelines was defined by the humane endpoint criteria described above, and this limit was not exceeded in any experiment.

Adoptive cell transfer

APCs (Thy1.2⁻) were purified from the splenocytes of wild-type C57BL/6 J using PE anti-mouse CD90.2 (Thy1.2) Antibody (Biolegend, #105307) and Anti-PE MicroBeads (Miltenyi Biotec, #130-048-801) and pulsed with the OVA_257–264 peptide overnight (GenScript, #RP10611, Piscataway, NJ) (OVA_257–264 peptide-pulsed APC). Subsequently, naive CD8⁺ T cells prepared from the splenocytes of OT-1 C57BL/6 J donor mice and Mitomycin C (FUJIFILM Wako Pure Chemical, #139-18711)-treated OVA_257–264 peptide-pulsed APCs were cocultured for 48 h in the presence or absence of ponatinib (10 nM) before transfer. Then, CD8⁺ T cells were collected again using a MojoSort™ Mouse CD8⁺ T Cell Isolation Kit (BioLegend, #480008). Prior to transfer, the phenotype of CD8⁺ T cells was confirmed by flow cytometry. One day before transfer, C57BL/6 J recipient mice bearing SIINFEKL-expressing B16F10 tumors (2 × 10⁵ cells, s.c., Day 0) were pretreated with cyclophosphamide hydrate (100 mg/kg, i.p.; Shionogi & Co., Ltd., Osaka, Japan). A total of 1 × 10⁶ CD8⁺ T cells were then transferred intravenously on Day 9 after tumor inoculation. On Day 8 after ACT, the mice were euthanized, and the TILs were collected for the flow cytometry analysis.

For systemic ponatinib administration experiments, OT-1 CD8⁺ T cells were activated for 48 h with OVA_257–264 peptide–pulsed APCs (without ponatinib), and OT-1 CD8⁺ T cells (1 × 10⁶) were transferred into B16F10-SIINFEKL tumor-bearing wild-type C57BL/6 J mice on Day 9 after tumor inoculation. The mice were orally administered ponatinib (3 mg/kg) or 1% 2-hydroxypropyl-beta-cyclodextrin once daily beginning one day before cell transfer and continuing for 14 consecutive days thereafter. On Day 8 after ACT, the mice were euthanized, and the TILs were collected for the flow cytometry analysis.

In the tumor growth experiments, both the donor and recipient mice had a CD45.2 background. For the flow cytometry experiments evaluating the expansion of transferred cells, CD45.2 OT-1 C57BL/6 J mice were used as the source of transferred T cells, and CD45.1 C57BL/6 J mice were used as recipients.

Seahorse assay

Metabolic analyses were performed using a Seahorse Bioscience Analyzer XFe96⁶². Naive CD8⁺ T cells purified from the splenocytes of wild-type C57BL/6 J mice by magnetic separation were stimulated with anti-CD3/CD28 mAbs in the presence of DMSO or ponatinib (10 nM) for 48 h. The cells (0.2 × 10⁶) were subsequently resuspended in extracellular flux assay medium supplemented with 11 mM glucose, 2 mM glutamine, and 1 mM sodium pyruvate and plated on a Cell-Tak (Corning)-coated microplate, allowing the adhesion of mouse CD8⁺ T cells. Mitochondrial activity and glycolytic parameters were measured by determining the oxygen consumption rate (OCR) (pmol/min) and extracellular acidification rate (ECAR) (mpH/min), respectively, using real-time injections of oligomycin (1.5 µM), carbonyl cyanide trifluoromethoxy phenylhydrazone (FCCP; 0.5 µM), and rotenone and antimycin (both at 0.5 µM). Respiratory parameters were calculated according to the manufacturer’s instructions (Agilent, Santa Clara, CA). The reagents used are listed in Supplementary Table 2.

Western blotting

Naive CD8⁺ T cells purified from the splenocytes of wild-type C57BL/6 J mice using magnetic separation were stimulated with anti-CD3/CD28 mAbs in the presence of DMSO or ponatinib (10 nM) for 2 h. Human CD8⁺ T cells isolated from the PBMCs of healthy individuals using magnetic separation were stimulated with Dynabeads Human T-Activator CD3/CD28 and DMSO, imatinib (2.5 nM), or ponatinib (50 nM) for 24 h. After washing with ice-cold PBS (-), the cells were lysed in 2 × Laemmli Sample Buffer (Bio-Rad Laboratories, #1610737, Hercules, CA). The cell lysates were resolved by SDS–PAGE and transferred to polyvinylidene difluoride membranes. Proteins were detected by incubation with primary antibodies followed by an incubation with a horseradish peroxidase-conjugated secondary antibody. For the analysis of phosphorylated and total forms of the same protein, membranes were stripped after detection of phosphorylated proteins and reprobed with antibodies recognizing the corresponding total proteins. The antibodies used are summarized in Supplementary Table 3. Images were obtained using a LuminoGraph I (ATTO Corporation, #WSE-6100H, Tokyo, Japan). Quantification of band intensities was performed using ImageJ software (v.13.0.6; National Institutes of Health, Bethesda, MD) and its macro plugin⁶³. Multiple targets were analyzed from the same samples using stripping/reprobing and parallel gels/blots; therefore, β-actin was not available as a same-blot loading control for all proteins. Same-blot β-actin was used as a loading control when available; otherwise, β-actin run on a parallel gel/blot from the same samples was used as a sample processing control and is annotated in the figure legends.

Confocal microscopy

Human CD8⁺ T cells purified from the PBMCs of healthy individuals by magnetic separation were stimulated with Dynabeads Human T-Activator CD3/CD28 along with DMSO, imatinib (2.5 nM), or ponatinib (50 nM) for 24 h. Then the cells were collected and resuspended in 4% paraformaldehyde (FUJIFILM Wako Pure Chemical, #161-20141) for 30 min, permeabilized with 0.5% Triton X-100 (FUJIFILM Wako Pure Chemical, #169-21105) for 5 min and blocked with PBS supplemented with 10% BSA for 1 h. Antibody staining was performed using the antibody dilutions described in Supplementary Table 4. The conjugates were allowed to settle onto poly(D) lysine-coated slides. The colocalization of phosphorylated SRC family members (Tyr416) (clone D49G4) and CD28 (clone CD28.2) was examined. Three-color confocal and transmitted light images were acquired using a Zeiss LSM 880 microscope (Zeiss, Thornwood, NY). Multiple random fields of view were photographed, and the frequency of cells showing colocalization of pSrc and CD28 was determined. The antibodies used are listed in Supplementary Table 4.

Phospho-flow cytometry

CD8⁺ T cells were purified from the PBMCs of healthy individuals with CD8 microbeads (Miltenyi Biotec, #130-045-201) and stimulated with Dynabeads Human T-Activator CD3/CD28 with imatinib (2.5 nM), ponatinib (50 nM), or an LCK inhibitor, a Src family activator, or an AKT activator. CD8⁺ T cells were transduced with constitutively active (CA)_LCK or CA_AKT for overexpression studies. The cells were then placed on ice and fixed with BD Phosflow Fix Buffer Ⅰ (BD Biosciences, #557870). The fixed cells were permeabilized with BD Phosflow Perm Buffer Ⅰ (BD Biosciences, #557885), stained for phosphorylated proteins, and analyzed using a FACSymphony A3 (BD Biosciences) and FlowJo v.10.6.1 (BD Biosciences). The antibodies used for cell staining are summarized in Supplementary Table 5.

Lentiviral vector transduction to human CD8⁺ T cells

Human CD8⁺ T cells were purified from PBMCs using CD8 MicroBeads (Miltenyi Biotec, #130-045-201). Cells were stimulated with Dynabeads Human T-Activator CD3/CD28 at a 1:1 cell-to-bead ratio and lentivirally transduced with vectors encoding constitutively active LCK (LCK-Y394E)³⁹, constitutively active AKT1 (myr-AKT1-T308D/S473D)⁶⁴, or the corresponding empty vector, all of which encoded for truncated EGFR (tEGFR)⁶⁵. Transduction with the LCK and AKT vectors was performed separately. Cells were cultured in RPMI-1640 supplemented with 10% FBS and 4 mM L-glutamine with recombinant human IL-7 and IL-15 (10 ng/mL each; R&D Systems #207-IL and #247-ILB) and treated with ponatinib (100 nM) or DMSO as indicated. On Day 4, the beads were removed with a magnet, and the cells were analyzed by flow cytometry on Day 8.

Generation of Human CAR-T cells

For human CAR-T cell preparation, CD3⁺ T cells derived from healthy individuals were lentivirally transduced with an FMC63.BBz (anti-CD19) CAR coexpressing a transduction/selection marker, tEGFR (CD19CAR-tEGFR). The CD19 CAR-tEGFR structure consists of a CD8α signal peptide, an anti-CD19 single-chain variable fragment (scFv), a CD8α hinge, transmembrane domains, a 4-1BB costimulatory domain, and a CD3ζ signaling domain with a tEGFR that lacks the membrane distal EGF-binding domain and the cytoplasmic domain via self-cleavage of the T2A sequence (see Fig. 7a)^65,66. CD3⁺ T cells were purified from the PBMCs of healthy individuals with an EasySep™ Human T Cell Isolation Kit (STEMCELL Technologies, #17951) according to the manufacturers’ instructions. The cells were stimulated with Dynabeads Human T-Activator CD3/CD28 at a 1:1 cell-to-bead ratio, lentivirally transduced with the anti-CD19-BBz CAR gene, and treated with either ponatinib (100 nM) or DMSO in RPMI-1640 supplemented with 10% FBS and 4 mM L-glutamine (Thermo Fisher Scientific, #25030-081). Recombinant human IL-7 (R&D System, #207-IL) and IL-15 (R&D System, #247-ILB) were added at 10 ng/mL. On Day 4, the beads were removed with a magnet, and the cells were allowed to rest for another 4 days. For scRNA-seq analyses, cells were sorted into CD4⁺ CAR-T cells (CD3⁺CD4⁺EGFR⁺) and CD8⁺ CAR-T cells (CD3⁺CD8⁺EGFR⁺) using a FACSDiscover™ S8 Cell Sorter (BD Biosciences), mixed at a 1:9 ratio, and then processed for library preparation (Supplementary Fig. 6c). The CD4⁺ CAR-T cells were mechanically removed, and only the data from the CD8⁺ CAR-T cells were analyzed.

Analysis of scRNA-seq data

scRNA-seq libraries (n = 2 per group) were constructed with a Chromium Next GEM Single Cell 3′ Reagent Kit v3.1 (10x Genomics, #PN-1000268, Pleasanton, CA) according to the manufacturer’s instructions. Sample pooling prior to library preparation was not performed. The cell suspension of each sample was processed in a Chromium Controller (10x Genomics) to generate single-cell gel beads-in-emulsions (GEM) for sample and cell barcoding. Amplified cDNA and constructed libraries were assessed with an Agilent BioAnalyzer using a High Sensitivity DNA Kit (Agilent Technologies, #5067-4626). The libraries were then pooled and sequenced on a NovaSeq X plus instrument (Illumina). The raw base call files were demultiplexed into reads in FASTQ files using the Cellranger mkfastq pipeline v.7.2.0. The Cellranger count pipeline v.7.2.0 was used for read filtering and alignment to the hg38 reference genome to generate a gene‒barcode matrix for each library. Dimensional reduction using the UMAP method and cell clustering were performed with general Scanpy pipelines⁶⁷ v.1.10.3. Differentially expressed genes were determined by decoupler⁶⁸ v.1.8.0 and pyDESeq2⁶⁹ v.0.4.12. GSEA was performed using GSEApy⁷⁰ v.1.1.4.

Cell viability assay

Cell viability was assessed using a Cell Counting Kit-8 (CCK-8; Dojindo, #341-07761) assay according to the manufacturer’s instructions. Nalm-6 cells were seeded in 96-well plates and treated with ponatinib immediately after seeding. A549 cells were seeded in 96-well plates and treated with ponatinib 12 h after seeding. After 72 h of culture, 10 μL of CCK-8 reagent was added to each well, and the absorbance was measured at 450 nm using a microplate reader (Bio-Rad).

Mouse xenograft models

Female NOD. Cg-Prkdc^scid Il2rg^tm1Wjl/SzJ (NSG) mice (6 to 10 weeks of age) were intravenously inoculated with 1 × 10⁶ tumor cells via the tail vein (Day -7). For the Nalm-6-derived leukemia model, luciferase-expressing Nalm-6 cells were used. Nalm-6 cell engraftment, expansion, and clearance were monitored by bioluminescence imaging (BLI) following intraperitoneal injection of luciferin (Promega, #E1605, Madison, WI) using a Spectrum IVIS bioluminescence imager and quantified using Living Image software v.4.7.3 (Perkin Elmer, Waltham, MA) under isoflurane anesthesia. Mice were randomized to each group before CAR-T cell injection. The mice were administered intravenously with 8 × 10⁴ to 1 × 10⁶ ponatinib-CAR-T cells on Day 0. For rechallenge experiments, the mice were intravenously inoculated with 1 × 10⁶ tumor cells on Day – 7. On Day 28, mice whose tumor burden was not detectable by BLI were subjected to a second inoculation with 1 × 10⁷ tumor cells via the tail vein.

Peripheral blood was collected weekly starting on Day 7. After removal of red blood cells (RBCs) with RBC lysis buffer (Merck, # 11814389001), the cells were stained and subjected to flow cytometry. The number of CAR-T cells per μL was assessed with CountBright™ Absolute Counting Beads (Invitrogen, #C36950). The expression of several markers by CAR-T cells was also analyzed by flow cytometry on Day 14. For the A549-derived solid tumor model, A549 cells expressing truncated CD19 lacking its cytoplasmic domain (hCD19-A549) were used. The mice were subcutaneously inoculated with 2 × 10⁶ tumor cells in the right flank (Day -7 or Day -10). Some mice received oral ponatinib treatment at 3 mg/kg starting on Day 3 and following a 5-day-on, 2-day-off regimen for 4 weeks. In some mice, TILs were collected on Day 17 and subjected to a flow cytometry analysis to assess CD8⁺ T cell exhaustion following ponatinib treatment.

In A549-derived solid tumor models, the tumor size was monitored by measuring the tumors with calipers twice a week. The mice were euthanized if they exhibited any of the following symptoms: > 20% loss of initial body weight; a tumor greater than 20 mm in maximum diameter; Nalm-6 cell bioluminescence greater than 5 × 10¹¹ photons per second; or evidence of extensive disease (for example, inability to ambulate, groom, or eat; cachexia; excessive loss of fur; hunched posture; or other signs of disability). Euthanasia was performed by cervical dislocation under deep anesthesia, in accordance with institutional guidelines. The tumor volume was calculated as length × width²× 0.5. Animal care and experiments were conducted according to the guidelines of the Animal Committee of the National Cancer Center Japan after being approved by the Ethics Review Committee for Animal Experimentation of the National Cancer Center Japan. All in vivo experiments were performed at least twice. The maximum tumor size permitted by institutional guidelines was defined by the humane endpoint criteria described above, and this limit was not exceeded in any experiment.

PK-mimicking in vitro culture

Purified human CD8⁺ T cells were obtained from PBMCs and stimulated with Dynabeads Human T-Activator CD3/CD28 in the presence of either dasatinib or ponatinib. The drug concentrations and stepwise reduction scheme were designed to recapitulate clinically relevant exposure patterns, assuming once-daily oral administration of 100 mg of dasatinib or 30 mg of ponatinib^46,71–73. For dasatinib treatment, the cells were exposed to 120 nM dasatinib at the start of culture, and the drug concentration was sequentially reduced to 60, 30, 15, and 5 nM every 3 h and maintained until 24 h. For ponatinib treatment, the cells were initially exposed to 80 nM ponatinib, followed by stepwise decreases to 70 nM at 3 h, 60 nM at 6 h, and 50 nM at 9 h, after which the concentration was maintained until 24 h.

Drug concentrations were adjusted by stepwise dilution through the sequential direct addition of cytokine-containing, drug-free medium to the wells, thereby reducing the drug concentration without harvesting or washing the cells. After 24 h, the medium was completely replaced to restore the starting concentration. These cycles were repeated daily for 5 consecutive days. Phosphorylation of LCK and AKT was monitored at multiple time points, and CD8⁺ T cell phenotypes were assessed at the end of the 5-day culture period using flow cytometry. K562 cells were also treated on the same schedules, and cell viability, including apoptosis, was evaluated by flow cytometry after 3 days.

Due to experimental constraints requiring the completion of stepwise dilution procedures within a single 12 h cycle, dasatinib decay was modeled as 50% reduction every 3 h, which is within the clinically reported half-life range of approximately 3–5 h⁷³. The concentration was maintained at 5 nM, corresponding to the clinically recommended trough level to minimize adverse events⁷⁴. With this design, the calculated AUC was comparable to values reported in clinical pharmacokinetic studies⁷³. This model represents an in vitro approximation and is not intended to quantitatively reproduce in vivo pharmacokinetics.

Statistical analysis

Comparisons between groups were performed using the two-sided unpaired t test, the two-sided Mann–Whitney test, or the two-sided paired t test, as appropriate. Two-way ANOVA was used for analyses of tumor growth kinetics. Survival curves were generated by the Kaplan–Meier method and analyzed using the log-rank test. Statistical analyses were performed with GraphPad Prism 10 (GraphPad Software, San Diego, CA). P-values < 0.05 were considered statistically significant.

Reporting summary

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

Author contributions

Contributions: Y.O. and H.N. designed the study; Y.O., S.I., K.W., Y.Y., K.K., T.S., T.K., and H.N. performed the experiments and analyzed the data; Y.O. and Y.K. performed the bioinformatics analysis; T.S., Y.F., H.K., and N.T. provided clinical human samples; Y.O., K.W., Y.M., and H.N. wrote the paper; H.N. supervised the study.

Peer review

Peer review information

Nature Communications thanks Carsten Riether and the other anonymous reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

Data availability

The RNA-seq and single-cell RNA-seq data generated in this study have been deposited in the Gene Expression Omnibus (GEO) database under accession codes GSE295191 and GSE294652 [https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE295191, and https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE294652]. Raw and processed data are available at GEO. All data are included in the Supplementary Information or available from the authors, as are unique reagents used in this Article. The raw numbers for charts and graphs are available in the Source Data file whenever possible. Source data are provided in this paper.

Competing interests

H.N. received research funding and honoraria (lecture fee) from Ono Pharmaceutical, BMS, Chugai Pharmaceutical, BD Japan, Eisai and MSD; received honoraria (lecture fee) from Amgen; received research funding from Taiho Pharmaceutical, Daiichi-Sankyo, Kyowa Kirin, Zenyaku Kogyo, Oncolys BioPharma, Debiopharma, Rakuten Medical, Gliead-Kite Pharma, Moderna, Asahi-Kasei, Sysmex, Fujifilm, SRL, Hitachi, Astellas Pharmaceutical, Sumitomo Pharma, ARC Therapies, Meiji Seika Pharma, MiraBiologics, and RIKAKEN Holdings; and served as a founder and a scientific advisor of ARC Therapies and a scientific advisor of LTZ Therapeutics outside of this study. Y.M. and K.W. are founders and directors of ARC Therapies outside of this study. H.K. received research funding from FUJIFILM, Kyowa-Kirin, Bristol-Myers Squibb, Otsuka, Perseus Proteomics, Daiichi Sankyo, Abbvie, CURED, Astellas Pharma, Chugai, Zenyaku Kogyo, Nippon Shinyaku, Eisai, Takeda, Sumitomo Pharma, and Sanofi, and honoraria from Abbvie, Chugai, Astellas Pharma, and Novartis. N.T. received research funding from Asahi-Kasei Pharma Corporation, Otsuka Pharmaceutical, Novartis, and Pfizer. The remaining authors declare no competing financial interests.

Supplementary information

The online version contains supplementary material available at 10.1038/s41467-026-71375-2.