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

Domain: chemistry
Published: 2026-04-07
Editorial outcome: major_revision

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.

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 therapy2-5. 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 cancers6,9. 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 proliferation10-12. Consequently, the therapeutic application of ponatinib has been investigated in multiple hematologic malignancies and solid tumors13-15. 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 cells16,17. 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.

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). Additionally, 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 subsets23,24 (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. Additionally, 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 Fig. 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 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).

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 Fig. 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 hours 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 3,589 differentially expressed genes (1,797 upregulated, 1,792 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 cells23-27.

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 formation23-26. 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 phenotype29,30, 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 immunotherapy23-25, 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/6J 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 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 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.

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).

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 longevity4,5. 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 peptide-pulsed antigen-presenting cells (APC) with or without ponatinib for 48 hours 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).

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.

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 cells23-26,35-37.

Since LCK is an essential member of 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 SCM 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 cells are characterized by long-term survival and robust antitumor activity, which contrast 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 cells4,36,37; 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.

Recent studies reported the influence of distinct CD4⁺ CAR-T cell subsets on therapeutic efficacy40,43,44. 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 therapy40,44, 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.

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.

We next investigated the in vivo antitumor efficacy of dasatinib using tumor-bearing mouse models. Wild-type C57BL/6J 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.

Methods
Sex as a biological variable
Female mice were used in this study. For the B16F10 tumor experiments in B6.Cg-Rag2tm1.1Cgn/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 cells EM (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 minutes. 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/6J 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 cells CM (CD8+CD44+CD62L+), and CD8+ T cells EM (CD8+CD44+CD62L-) were purified from the splenocytes of wild-type C57BL/6J mice with a BD FACSAria™ Fusion cell sorter (Supplementary Fig. 6b). The purity of each phenotype was confirmed to be >95% (Supplementary Fig. 4h). 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 hours, the cells were collected for Western blotting, and after 48 hours, 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 hours. 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/6J mice (#000664), BALB/c nude mice (#000651), B6.Cg-Rag2tm1.1Cgn/J (RAG2 knockout) mice (#008448), B6.129(Cg)-Foxp3tm3(Hbegf/GFP)Ayr/J (Foxp3-DTR) mice (#016958), C57BL/6-Tg(TcraTcrb)1100Mjb/J (OT-1) mice (#003831), and NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ (NSG) mice (#005557) were purchased from Jackson Laboratory (Bar Harbor, ME). CD45.1 C57BL/6J mice were purchased from Sankyo Labo Service Corporation (Tokyo, Japan). Female mice were used unless otherwise specified; for B16F10 tumor experiments in B6.Cg-Rag2tm1.1Cgn/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. 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/6J 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 peptide overnight (GenScript, #RP10611, Piscataway, NJ) (OVA peptide-pulsed APC). Subsequently, naive CD8+ T cells prepared from the splenocytes of OT-1 C57BL/6J donor mice and Mitomycin C (FUJIFILM Wako Pure Chemical, #139-18711)-treated OVA peptide-pulsed APCs were cocultured for 48 hours 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/6J 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 hours with OVA peptide–pulsed APCs (without ponatinib), and OT-1 CD8+ T cells (1 × 10⁶) were transferred into B16F10-SIINFEKL tumor-bearing wild-type C57BL/6J 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/6J mice were used as the source of transferred T cells, and CD45.1 C57BL/6J 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/6J mice by magnetic separation were stimulated with anti-CD3/CD28 mAbs in the presence of DMSO or ponatinib (10 nM) for 48 hours. 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/6J mice using magnetic separation were stimulated with anti-CD3/CD28 mAbs in the presence of DMSO or ponatinib (10 nM) for 2 hours. 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 hours. 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 hours. 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 hour. 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 a FMC63.BBz (anti-CD19) CAR coexpressing a transduction/selection marker, tEGFR (CD19CAR-tEGFR). The CD19 CAR-tEGFR structure consists of a CD8⍺ signal peptide, an anti-CD19 single-chain variable fragment (scFv), a CD8⍺ 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 hours after seeding. After 72 hours 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-Prkdcscid Il2rgtm1Wjl/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 after being approved by the Ethics Review Committee for Animal Experimentation of the National Cancer Center. 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 ponatinib46,71-73. 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 hours and maintained until 24 hours. For ponatinib treatment, the cells were initially exposed to 80 nM ponatinib, followed by stepwise decreases to 70 nM at 3 hours, 60 nM at 6 hours, and 50 nM at 9 hours, after which the concentration was maintained until 24 hours.

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 hours, 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-hour cycle, dasatinib decay was modeled as 50% reduction every 3 hours, which is within the clinically reported half-life range of approximately 3–5 hours73. 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.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Okuhiro and colleagues describe in their study that the tyrosine kinase inhibitor ponatinib affects TCR-dependent proliferation and differentiation of CD8+ T cells in mice and humans. In addition, they show that ponatinib-treatment potentiates CD8+ T cell activity in murine syngeneic models in combination with anti-PD1 treatment and in a murine CAR T cell model. Mechanistically, they show that inhibition of LCK and consequently the PIK3 kinase is downregulated in CD8+ T cells upon Ponatinib treatment and leads to the generation stem cell like T cells. This study overall sound and the experiments well-designed. However, not all conclusions are supported by experimental data.\n\nMajor remarks:\n1. Figure 1b: The authors conclude that there is no difference in cell division between DMSO and Ponatinib treated CD8+ T cells. However, the CFSE dilution plot presented suggests that there is a substantial proportion of T cells that did not divide after treatment (11.6 vs 4.7 %). In addition, it seems as more cells divided > 5 times after ponatinib treatment. The authors should show the frequency of cells per division step and adapt their conclusion accordingly. What is the phenotype of the non-dividing cells? TSCM?\n2. Figure 1: How are absolute Treg and CD8+ T cell numbers affected?\n3. Figure 3: What is the phenotype/differentiation of the cells that are injected? If more cells with a TSCM phenotype are injected, it is somehow not surprising that a higher frequency of TSCM cells are found at day 8 in the tumor. Authors should perform an in vivo treatment with ponatinib to proof that this finding has a clinical relevance.\n4. Figure 4: The authors should expose their T cells to a LCK or PI3K kinase activator in presence and absence of ponatinib to proof their conclusion presented in Figure 4m.\n5. Figures 6: The authors should treat the xenografted Nalm-6 mice with ponatinib in vivo to support their conclusion that CAR T cells can better persists in vivo. So far they authors only demonstrate that there is increased anti-tumoral activity if more CAR T cells with a TSCM phenotype are transferred in the xenografted mice.\n6. Figure 6: Re-challenge experiment. The authors should wait to re-challenge mice that were previously exposed to DMSO and Ponatinib in vivo until they have a similar leukemia load. Else different effector: target ratios are compared between groups. An alternative would be transfer CAR T cells from primary mice into secondary NALM-6 xenografted mice.\n7. Figure 7: The authors pooled CML patients and ALL patients in the Ponatinib group, while the control only contains CML patients. Please separate the Ponatinib group in CML and ALL patients and draw a conclusion comparing CML (Imatinib) with CML (Ponatinib).\n8. Discussion: Cardio toxicity is big issue in CAR T cell therapy. Ponatinib most cardio-toxic FDA-agent. Authors should at least discuss this topic as they propose to use Ponatinib in combination with CAR T cell therapy. Furthermore, CAR T cell patients frequently suffer from viral infections and complications.\nImatinib has been already shown to impair expansion of memory cytotoxic T cells without affecting the control of primary viral infections (Mumprecht et al. 2006 Blood). As CD8+ T cells are the effector cells that control most viral infections, the authors should discuss this limitation for the clinical use of Ponatinib\n9. Title: Please adapt the title. There is not proof that Ponatinib “rejuvenates” T cells. It just affects their differentiation state.\n\nMinor comments:\n1. Figure S4: The differences in survival are so marginal and not meaningful, yet the differences are highly significant. What statistical test was used? The authors should compare the mean overall survival between the groups.", "reviewer_2": "Dr Nishikawa and colleagues describe the role of Src family kinase inhibitor ponatinib on the phenotype, subset composition, and anti-tumor function of polyclonal and CAR-engineered T cells. The authors use both human and mouse models of cancer, including checkpoint blockade combined with in vivo ponatinib and expansion of CAR T cells ex vivo in the presence of ponatinib. The authors show ponatinib addition improves anti-tumor function of T cells and increases the frequency of \"Tscm\" cells thus potentially explaining the effect.\nOverall, many aspects of the study replicate prior observations with a similar drug, dasatinib, which also inhibits Src family kinases Lck and Fyn and reduces T-cell activation (Mestermann et al, STM ~2020) and exhaustion/differentiation (Weber et al., Science ~2021). Indeed, the observed results can be explained by inhibiting T-cell activation via a mechanism shared between dasatinib in the above studies and ponatinib in the current study, leading to an enrichment for less-differentiation naive-like T cells, downregulation of activation markers including CD25, CTLA-4 and FoxP3, differences in cell metabolism secondary to the lack of effector differentiation, and improved anti-tumor activity as observed in Weber/Mackall et al, Science paper (eg Figures 5 and 6 mirror findings in Figures 6 and 7 of the reference manuscript). The enrichment of T cells for Tscm is difficult to evaluate without measuring total expansion of T cells w/wo ponatinib and calculating absolute numbers of Tscm at the end of manufactuing in both conditions.\nThe observed differences in the activity of ponatinib vs imatinib are explained by a >1000-fold higher Lck-inhibiting activity of ponatinib/dasatinib compared to imatinib (Lee et al., Leukemia 2010). A similar effect could be achieved by lowering the concentration of anti-CD3/CD28 to reduce initial Lck/Fyn activity or, in some models including the OT-I model in Figure 3, removing T-cell activation altogether.\nTherefore, the novelty of this study is limited as both the mechanism and the effects of pharmacologic Lck/Fyn inhibition in CAR T cells have been previously described."}, "author_response": "We are grateful for the reviewer’s enthusiastic and supportive comments on our manuscript. In this revision, we addressed the reviewer’s concerns carefully to strengthen our conclusions. In particular, we have revised the Results and Discussion sections to provide clearer and more direct support for our conclusions using the data. Our detailed responses are described below. [Detailed point-by-point response follows]"}, {"round": 2, "reviewer_comments": {"reviewer_1": "Revised version of the manuscript:\nThe authors have adequately addressed most of my concerns through additional experiments, which have improved the overall quality of the manuscript. They have also revised the title, conclusions, and discussion accordingly. Therefore, I recommend accepting the manuscript for publication.", "reviewer_2": "The authors have responded to the critique by providing evidence of importance for the sustained Lck inhibition with ponatinib in vivo to achieve highest levels of Tscm and completely changing the title of the paper. While this is a welcome development, some concerns remain.\n1. The \"transient PK\" model in vitro does not accurately reflect plasma levels of dasatinib vs ponatinib as the authors completely withdraw dasatinib for 12h while maintaining ponatinib at intermediate concentrations. While the PK of dasatinib is indeed more short-lived, the differences can be offset by increasing the dose 2.5x (Yoshimura et al., Leukemia 2023) which results in the same AUC and functional Lck inhibition. It is wrong to assume dasatinib levels will be at 0 after 12h as its half-life is 5.6 hours.\n2. Concentrations are important. The authors use 3mg/kg of ponatinib and 50mg/kg dasatinib (16.7x higher). At high doses, dasatinib fully blocks Lck/Fyn activity thus precluding any TCR-mediated activity which may impact activation and survival of T cells and thus result in a minimal tumor control. As discussed in #1, dasatinib levels should be increased 2.5-3x to achieve the same Lck inhibition in mice and not over-cook it with a 16x increase.\n3. In the end, the main novelty of this study appears to be drug regimen and dosing but not a conceptual advancement that Bcr-Abl TKI also inhibit Lck and thus modulate T-cell differentiation.\n4. For #2, the authors discuss absolute cell counts but Figure 5i shows relative cell counts on the Y axis."}, "author_response": "We thank the reviewer for the thoughtful comments, acknowledgment of our efforts to provide additional evidence of the importance of stable and sustained Lck inhibition by ponatinib in vivo for inducing T cells, and recognition of our revised title. We have performed further experiments to address the concerns regarding the dosing and exposure conditions of ponatinib and dasatinib. Our detailed responses are described below. [Detailed point-by-point response follows]"}, {"round": 3, "reviewer_comments": {"reviewer_2": "The authors have put effort in addressing my comments. Unfortunately, some arguments are based on flawed assumptions or factually incorrect data and need to be re-visited.\n1. The authors provided an extensive analysis to answer query #1 citing PK data and clinical studies to justify dropping dasatinib concentration to 0 after 12h of each spike. Here's why those arguments aren't holding water:\ni) The authors claim the Yoshimura 2023 study used dasatinib doses (100-140mg QD) that \"far exceed approved dosage\" citing the Zhu 2023 paper. This is incorrect as dasatinib is indeed approved at 100-140 mg doses, and the Zhu paper is a study in pediatric patients where the doses are obviously lower than in adults.\nii) The authors argue the \"real half life\" of dasatinib is 3h not 5h. This is a meaningless discussion since there half life of the drug varies with plasma concentration and the early half life at high concentrations is shorter than the terminal half-life at lower levels, hence the 3-5h range. It is incorrect to assume dasatinib half life is stable at 3h.\niii) Regardless of the half life value, it is incorrect to postulate that dasatinib levels will go down to 0 after 12h. At a \"median\" half life of 4h, dasatinib levels will be at 12.5% of the initial concentration, which is approximately 3 times higher in dasatinib compared to ponatinib (~100-140mg dasatinib vs 45mg ponatinib).\niv) The \"PK\" model used in Fig 7 is visually incoherent since only dasatinib is ratcheted down to 0 after 12h (which is incorrect, as discussed above) but ponatinib stays at a plateau after 12h. Both drugs should be titrated down every 3-6h\nIn light of this, I feel the PK model needs to be re-done or omitted altogether.\nTo be clear, the effect of ponatinib demonstrated in these models is comprehensive and will be of interest to the clinical community. However, I do find the comparison with other TKI (imatinib and dasatinib) to be the weakest point of this study. It is well known that the strength of TCR stimulation affects T cell differentiation, with weaker TCR signaling favoring Tscm formation. This has been demonstrated with dasatinib (PMID: 34289897) which, together with ponatinib, is a strong inhibitor of Lck/Fyn - key kinases downstream of TCR/CD3 signaling. Imatinib, on the other hand, has a much weaker activity that's not meaningful at real-world plasma concentrations, so is not a \"fair comparison\", especially in Figure 4. It is well known that LCK inhibition by dasatinib suppresses TCR signaling including the PI3K-mTOR axis which is activated by PLCg signaling downstream of zeta chain that's directly inhibited by ponatinib or dasatinib.\nThe appropriate PK of ponatinib to achieve sustained TCR inhibition is a good point and is thus the main novelty of this paper but the fundamental mechanism is shared with dasatinib or other modulators of TCR signaling. Otherwise it feels like the authors are re-discovering the mechanism known for a while, but using a latest-generation drug."}, "author_response": "We would like to thank the reviewer for the comments and for giving us the opportunity to revise our manuscript. We have revised the manuscript to address the concerns regarding the dosing and exposure conditions of ponatinib and dasatinib. Our detailed responses are provided below. [Detailed point-by-point response follows]"}]

Decision letter