Dynamic feedback control of oncogenic tyrosine kinase signaling in acute leukemia

Jaewoong Lee; Ruifeng Sun; Kohei Kume; Mark E Robinson; Zhangliang Cheng; Kadriye Nehir Cosgun; Ning Ma; Christian Hurtz; Huimin Geng; Selina M Luger; Mark R Litzow; Elisabeth Paietta; Jianjun Chen; Nagarajan Vaidehi; Markus Müschen

doi:10.1126/scisignal.adw5054

. Author manuscript; available in PMC: 2026 Feb 22.

Published in final edited form as: Sci Signal. 2026 Feb 10;19(924):eadw5054. doi: 10.1126/scisignal.adw5054

Dynamic feedback control of oncogenic tyrosine kinase signaling in acute leukemia

Jaewoong Lee ^1,^2,^3,^4,^*, Ruifeng Sun ^1,^2,^*, Kohei Kume ¹, Mark E Robinson ¹, Zhangliang Cheng ^1,², Kadriye Nehir Cosgun ¹, Ning Ma ⁴, Christian Hurtz ⁵, Huimin Geng ⁶, Selina M Luger ⁷, Mark R Litzow ⁸, Elisabeth Paietta ⁹, Jianjun Chen ⁴, Nagarajan Vaidehi ⁴, Markus Müschen ^1,^2,^†

PMCID: PMC12924454 NIHMSID: NIHMS2147568 PMID: 41666265

Abstract

CD25 is a subunit of the interleukin-2 (IL-2) receptor on T cells and natural killer (NK) cells. Acute leukemias with oncogenic tyrosine kinases often include CD25⁺ leukemia subpopulations, which portend poor clinical outcomes for patients; however, acute leukemia cells do not respond to IL-2. Here, we identified CD25 and its phosphorylation by protein kinase Cδ (PKCδ) as central elements of a feedback loop that stabilized fluctuations in oncogenic tyrosine kinase signaling in acute lymphoblastic and myeloid leukemia. Genetic ablation of CD25 in murine and patient-derived xenograft (PDX) models of acute leukemias reduced clonal fitness, colony formation, and leukemia-initiation capacity in serial transplant recipients. Oncogenic tyrosine kinase signaling in leukemia cells stimulated NF-κB–mediated CD25 expression, whereas PKCδ-mediated phosphorylation of CD25 suppressed oncogenic tyrosine kinase signaling through inhibitory phosphatases, such as PTPN6. Interactome analyses and mass spectrometry–based global phosphoproteomic analyses showed that CD25 deletion abolished the phosphatase activity of PTPN6, resulting in enhanced activation of tyrosine kinases and NF-κB. Four injections of a CD25 antibody-drug conjugate induced complete remission in mice transplanted with PDX refractory leukemia. These findings highlight the dependency of tyrosine kinase–driven leukemias on robust feedback control and the role of PKCδ and CD25 in assembling its components.

INTRODUCTION

In addition to the oncogenic BCR-ABL1 tyrosine kinase¹, reflecting the Philadelphia chromosome (Ph) in Ph⁺ acute lymphoblastic leukemia (B-ALL), multiple recurrent lesions targeting Janus kinases (JAKs)^2–3 and the platelet-derived growth factor receptor (PDGFR) subfamily⁴ of receptor-type tyrosine kinases (RTKs) form a large subset of similar biological features with poor clinical outcomes, termed Ph-like B-ALL^5–7. Similarly to oncogenic tyrosine kinases in B-ALL, the FLT3^ITD tyrosine kinase⁸ is linked to unfavorable clinical outcomes in patients with acute myeloid leukemia (AML)⁹. In most acute leukemias, cells expressing CD25 are not detected. However, in some cases of B-ALL^10–11 and AML^12–14, leukemias include subpopulations of cells expressing CD25, which portends poor clinical outcome. CD25 was previously identified as a subunit of the interleukin-2 (IL-2) receptor (IL-2R) on lymphocytes, in particular T regulatory (T_reg) cells and natural killer (NK) cells^15–20. However, whether acute lymphoblastic and myeloid leukemia cells are responsive to IL-2 was not previously investigated; hence, the functional importance of CD25 surface expression in leukemic subpopulations and its association with the poor clinical outcomes of CD25⁺ B-ALL and AML cells was unclear.

The IL-2R β-chain (IL-2Rβ) and the common γ-chain (IL-2Rγ) form the IL-2R expressed on resting T cells^21–22, which can be complemented by CD25 to form the heterotrimeric, high-affinity IL-2R²³. The IL-2Rβ and IL-2Rγ-chains, but not CD25, contribute to IL-2 signal transduction^21–22. All three chains of the IL-2R are endocytosed together after IL-2 binding,; however, the IL-2Rβ and IL-2Rγ chains are targeted to late endosomes for degradation, whereas CD25 is segregated to early endosomes for recycling to the plasma membrane^24–25. Consistent with their different fates in IL-2–activated T cells, the half-life of CD25 (>40 hours) is much longer than that of IL-2Rβ and IL-2Rγ (~1 hour) chains^24–25. Its lack of contribution to IL-2 signaling, asynchronous half-life, and divergent intracellular trafficking suggest that CD25 may have signaling functions that are independent from those of the other two IL-2R chains. Previously, we and others showed that leukemia cells critically rely on the inhibitory phosphatases INPP5D and PTPN6 to curb oncogenic signaling^26–28. Here, we discovered a previously unrecognized CD25-dependent feedback loop for the recruitment and activation of INPP5D and PTPN6 to constrain oncogenic tyrosine kinase signaling in B-ALL and AML. These findings provide a mechanistic link between CD25 expression and poor clinical outcomes and identify CD25 as a promising target in the treatment of refractory tyrosine kinase-driven B-ALL and AML.

RESULTS

CD25 expression in B-ALL and AML subpopulations is linked to oncogenic tyrosine kinase signaling

In six clinical cohorts of leukemia patients, including pediatric and adult B-ALL (fig. S1a–d) and AML (fig. S2a–c), most leukemia cases lacked expression of CD25. However, leukemias with oncogenic tyrosine kinases including mutations in JAK1, JAK2, or JAK3, and BCR-ABL1, PDGFRB, CSF1R, and NTRK3 fusions in B-ALL (fig. S1e–h) and JAK2 and FLT3-ITD lesions in AML (fig. S2d–e) frequently contained CD25⁺ subpopulations, ranging from 5% to about 40% of leukemia cells (fig. S1g–h). Detection of CD25 mRNA abundance in B-ALL and AML samples, originating from CD25⁺ leukemia subpopulations, was associated with poor clinical outcomes in all six clinical cohorts (fig. S1a–d; fig. S2a–c).

In mouse models of tyrosine kinase–driven B-ALL and AML, we detected CD25⁺ leukemia subpopulations (Fig. 1a–c). Similarly, the expression of BCR-ABL1, PAX5-JAK2, EBF1-PDGFRB, and FLT3-ITD induced CD25 surface expression in a fraction of B cell and myeloid progenitors (fig. S2f). Tyrosine kinase inhibition in B-ALL (BCR-ABL1, JAK2) and AML (FLT3-ITD) PDX for 12 hours did not induce cell death in three patient-derived leukemia xenografts (PDX), but abolished CD25 surface expression (fig. S2g).

Leukemia subpopulations with active tyrosine kinase signaling are marked by CD25 expression

To study the relationship between oncogenic tyrosine kinase signaling and CD25⁺ leukemia subpopulations, we developed a dual reporter system for phosphorylation of the BCR-ABL1 substrate CRKL (at Tyr²⁰⁷; Y207)²⁹ and Ca²⁺ signaling (jRCaMP1a)³⁰. Doxycycline-inducible activation of BCR-ABL1 induced marked phosphorylation of CRKL-Y207 and subsequently reduced Ca²⁺ signaling (Fig. 1d). Gating flow cytometry analysis on CD25⁺ leukemia cells (Fig. 1e), revealed the emergence of a CD25⁺ leukemia subpopulation (21%) with marked phosphorylation of CRKL-Y207 and suppressed Ca²⁺ signaling. Upon doxycycline-mediated induction of BCR-ABL1 expression in all cells, increased tyrosine kinase activity and phosphorylation of CRKL-Y207 were confined to a leukemia subpopulation that was defined by CD25 expression and reduced Ca²⁺signaling (Fig. 1e). These findings suggest that CD25 surface expression marks leukemia subpopulations with oncogenic tyrosine kinase signaling and negative regulation of Ca²⁺signaling.

CD25 expression reflects negative feedback control of oncogenic tyrosine kinase signaling

The transcription factor Nr4a1 promotes negative feedback regulation of Ca²⁺ and NF-κB signaling^31–32 downstream of tyrosine kinase signaling in B cells and myeloid cells^33–34. To investigate the role of CD25 expression in the context of tyrosine kinase activity and negative regulation of Ca²⁺signaling in acute leukemias, we induced expression of oncogenic tyrosine kinases in murine B cell and myeloid progenitor cells that were isolated from Nr4a1-reporter knockin mice³³. Induction of BCR-ABL1 and FLT3-ITD induced CD25 surface expression in parallel with activation of the Ca²⁺and NF-κB feedback regulator Nr4a1 (Fig. 1a–c). Upon acute BCR-ABL1 activation, CD25 surface abundance was maximal after 12 hours. When BCR-ABL1 signaling continued for longer times, CD25 surface abundance returned to basal amount in most B-ALL cells, whereas CD25⁺ subpopulations remained detectable at steady state (Fig. 1c), similar to patient-derived BCR-ABL1 B-ALL cells (Fig. 1f).

CD25⁺ subpopulations are small, quiescent, and drug-resistant leukemia cells

CD25⁺ leukemia cells in three patient-derived BCR-ABL1 B-ALL samples were quiescent and mostly in the G0/G1 phase of the cell cycle. In contrast, matched CD25⁻ cells from the same B-ALL patient samples were actively proliferating (Fig. 1f–g). Compared to CD25⁻ cells, CD25⁺ B-ALL cells were much smaller in cell size (as measured by forward scatter) and quiescent (Fig. 1f–g). Consistent with previous work³⁵, the small cell size and quiescence of CD25⁺ leukemia cells was paralleled by markedly increased resistance to vincristine, which induces mitotic arrest in leukemia cells (Fig. 1h). These results suggest that CD25 is not only induced in the context of feedback control of oncogenic tyrosine signaling but also marks drug-resistant and quiescent subpopulations in patient-derived leukemia.

Absence of a clonal hierarchy between CD25⁺ and CD25⁻ leukemia cells

To determine the clonal relationship between the CD25⁺ and CD25⁻ leukemia subpopulations, we sorted single CD25⁺ and CD25⁻ BCR-ABL1 B-ALL cells from three patient-derived xenografts and confirmed their purity by flow cytometry (Fig. 1i). After two weeks of outgrowth, single cell–derived CD25⁺ and CD25⁻ B-ALL clones gave rise to predominantly CD25⁻ daughter cells and smaller CD25⁺ subpopulations (Fig. 1i), which resembled the proportions in patient-derived BCR-ABL1 B-ALL (Fig. 1f). The finding that single CD25⁺ and CD25⁻ BCR-ABL1 B-ALL cells were each able to reconstitute both populations suggests that there is no progenitor-progeny hierarchy between CD25⁺ and CD25⁻ clones and that individual clones can transition between CD25⁺ and CD25⁻ states. Confirming the lack of a clonal hierarchy, we found that CD25⁺ and CD25⁻ leukemia cells had similar capabilities of initiating leukemia in transplant recipients (Fig. 1j). At a low dose of 1,000 cells, CD25⁻ leukemia cells initiated fatal disease slightly more rapidly than did their CD25⁺ counterparts (Fig. 1j), likely reflecting the quiescent nature of CD25⁺ leukemia cells (Fig. 1g).

CD25 expression reflects NF-κB activation in leukemia subpopulations

To compare gene expression patterns between CD25⁺ and CD25⁻ cells within individual leukemia samples, we flow-sorted matched CD25⁺ and CD25⁻ populations from three patient-derived BCR-ABL1 B-ALL samples for RNA-sequencing (RNA-seq) analysis (fig. S3a). Gene set enrichment analyses (GSEA) identified NF-κB transcriptional targets, B cell receptor primary response genes, and NFATC2 target genes as the top-ranking gene sets in CD25⁺ subpopulations (Fig. 1k, fig. S3a). ChIP-seq analyses of BCR-ABL1 B-ALL cells revealed that multiple NF-κB subunits bound to intronic CD25 enhancer elements (Fig. 1l), suggesting that NF-κB transcriptionally activates CD25 expression. To test this hypothesis, we introduced IKK2^S177E and MYD88^L252P NF-κB activating oncogenes^36–37 into murine BCR-ABL1 B-ALL cells. Including the known NF-κB target gene IκBα as a reference, NF-κB activation increased CD25 mRNA abundance by 6- to 11-fold and CD25 surface abundance by ~15-fold (Fig. 1m–n). We also detected enrichment for an NF-κB gene expression signature in clinical B-ALL and AML cohorts, comparing leukemia cases that contained a CD25⁺ subpopulation (cut-off 1% CD25⁺) to other cases that only included CD25⁻ leukemia cells (fig. S3b–d). For the B-ALL cohort (ECOG 2993; n = 165 patients), we identified a gene expression signature that correctly classified B-ALL cases containing a CD25⁺ subpopulation at 94% accuracy (fig. S3b). This gene set was strongly enriched for NF-κB and NF-κB target genes in B-ALL cases that contained a CD25⁺ subpopulation (fig. S3c). Similar observations were made for four clinical cohorts of patients with AML. Comparing gene expression patterns in AML cases with and without CD25⁺ subpopulations from the TCGA (n=140), MDACC (n=191), MLL Munich Leukemia Laboratory (n=325), and the Dresden, Munich, Ulm group (n=251) consistently showed that AML cases with CD25⁺ subpopulations were enriched for NF-κB transcriptional programs (fig. S3d).

B cell precursors express CD25 in response to oncogenic tyrosine kinase signaling but not IL-2 signaling

In T cells and NK-cells, CD25 has a central role as one of three chains of the IL-2R^15–17. Murine thymocytes responded to IL-2 with marked phosphorylation of STAT5; however, pre-B cells were not responsive to IL-2 (fig. S4a–b). Similarly, T cells and NK cells further increased their surface CD25 abundance upon IL-2 exposure^15–17; however, pre-B cells did not express CD25 upon addition of IL-2. Consistent with their lack of IL-2 responsiveness, pre-B cells, B-ALL cells, and AML cells lacked expression of the IL-2Rβ chain, which is required for the transduction of IL-2 signals^40–41. We found that doxycycline-induced BCR-ABL1 expression and initiation of oncogenic tyrosine kinase signaling markedly increased CD25 surface abundance (fig. S4a). Together, these results suggest that unlike for IL-2 signaling in T cells and NK cells, CD25 expression on B-lineage cells is increased by oncogenic tyrosine kinases and not by IL-2⁴⁰. Compared to tyrosine kinase–driven leukemia, tamoxifen-inducible deletion of CD25 in normal B cell precursors only transiently reduced competitive fitness in vitro (fig. S4c). Because CD25 is transiently expressed at the pre-B cell stage³⁸, we assessed the consequences of CD25 ablation on B cell development in vivo, in a genetic mouse model of B cell–specific deletion of CD25 (Cd25^fl/fl × Mb1-Cre). Studying B cell populations by flow cytometry ex vivo revealed minor changes, including a reduction in the number of pre-BCR⁺ Fraction C’ pre-B cells and a modest increase in the number of immature B-cells (Fraction F; fig. S4d–f). Consistent with a previous study⁴², these results suggest that CD25 is largely dispensable for normal B cell development.

CD25 is essential for competitive fitness and the initiation of tyrosine kinase–driven leukemia

CD25 was selectively expressed on the surface of B-ALL and AML cells that harbored oncogenic tyrosine kinases (fig. S1e–h, S2d) and was increased by oncogenic tyrosine kinase signaling (Fig. 1a–c, fig. S2f–g). To determine the functional role of CD25 expression in tyrosine kinase–driven B-ALL (BCR-ABL1, PAX5-JAK2, EBF1-PDGFRB) and AML (FLT3^ITD), we developed genetic models for 4-hydroxy-tamoxifen (4-OHT)-inducible deletion of Cd25 (Cd25^fl/fl). B cell and myeloid progenitor cells from the bone marrow of Cd25^fl/fl mice were transduced with retroviruses expressing BCR-ABL1, PAX5-JAK2 and EBF1-PDGFRB to model tyrosine kinase–driven B-ALL and MLL-AF9 and FLT3^ITD as an AML model. Growth factor–independent Cd25^fl/fl B-ALL and AML cells with tyrosine kinase oncogenes were transduced with GFP-tagged 4-OHT inducible Cre-ER^T2 or ER^T2 empty vector controls. In competitive cell culture assays, ablation of CD25 rapidly reduced the fraction of Cre (GFP⁺) cells, revealing the loss of competitive fitness of tyrosine kinase–driven B-ALL and AML cells upon Cd25 deletion (Fig. 2a–d). After the ablation of CD25, BCR-ABL1 B-ALL and MLL-AF9 FLT3^ITD AML cells underwent cell cycle arrest in the G₀/G₁ phase and lost colony formation capacity (Fig. 2e–f). Consistently, ablation of CD25 also affected the ability to initiate leukemia in serial transplant recipients (Fig. 2g). Deletion of Cd25 significantly prolonged the survival of primary transplant recipients. In serial transplant experiments, CD25 ablation abolished leukemia initiation, and secondary recipients survived for indefinite periods of time (Fig. 2g).

Fig. 2. — (A to D) Murine IL-7–dependent B-cell precursors from *Cd25*^fl/fl bone marrow were retrovirally transformed to express BCR-ABL1 (A), EBF1-PDGFRB (B) and PAX5-JAK2 (C) to develop growth factor-independent B-ALL. B-ALL cells were subsequently transduced with 4-hydroxy-tamoxifen (4-OHT)-inducible Cre-ER^T2-GFP or ER^T2-GFP empty vector control. For in vitro studies, percentages of GFP⁺ cells were measured by flow cytometry at the indicated times (A to C). Effects of inducible deletion of CD25 in *Cd25*^fl/fl B-ALL cells were monitored by flow cytometry after 4-OHT treatment. Flow plots are representative of three biological replicates. (D) Murine IL-3–, IL-6–, and SCF-dependent progenitor cells from *Cd25*^fl/fl bone marrow were retrovirally transformed to express MLL-AF9 and FLT3^ITD to develop growth factor–independent AML. FLT3^ITD AML cells were subsequently transduced with 4-OHT–inducible Cre-ER^T2-GFP or ER^T2-GFP empty vector control. Percentages of GFP⁺ cells were measured by flow cytometry at the indicated times. Flow plots are representative of three biological replicates. (E and F) Murine *Cd25*^fl/fl BCR-ABL1-driven B-ALL cells (E) and FLT3^ITD-driven AML cells (F) were transduced with retroviruses to express 4-OHT-inducible Cre-ER^T2 or ER^T2 empty vector control. Cell cycle analyses were performed by measuring BrdU incorporation in combination with 7AAD staining after 4-OHT treatment for 2 days, and the percentages of cells in S-phase are indicated (top). Cells were plated in semi-solid methylcellulose in the presence of 4-OHT for colony-forming assay for 7 days, and colony-forming capacity was assessed (bottom). Plots and microscopic images are representative of three biological replicates. Scale bar, 2.5 mm. (G) For in vivo studies, BCR-ABL1 *Cd25*^fl/fl B-ALL cells were transduced with retroviruses to express Cre-ER^T2-GFP or ER^T2-GFP empty vector controls. 50,000 GFP⁺ B-ALL cells were injected for each condition into sublethally irradiated (2 Gy) NSG mice (i.v. tail vein). Seven mice per group were used in each experiment. Kaplan-Meier analysis comparing overall survival of recipient mice was performed, and statistical significance was assessed by Mantel–Cox log-rank test. (H) Patient-derived Ph⁺ (PDX2) xenograft B-ALL cells were electroporated to introduce Cas9 RNPs containing guide RNAs for CRISPR-mediated deletion of CD25 (g-CD25) or nontargeting controls (g-NT). PDX2 B-ALL cells were subjected to colony-forming assay, and total colony numbers were counted 14 days after plating in semi-solid methylcellulose. Microscopic images of colony formation assay are representative of three biological replicates. Scale bar, 7 mm (left), 2.5 mm (middle), and 1 mm (right). (I) Human B-ALL cells (MUTZ5) with *JAK2*^R683G and *IGH@CRLF2* were transduced with lentiviruses to express doxycycline-inducible Cas9 and RFP-labeled guide RNAs for CRISPR-deletion of CD25 (sg-CD25) or non-targeting controls (sg-NT). Percentages of RFP⁺ cells were measured by flow cytometry at the indicated times. Flow plots are representative of three biological replicates.

To corroborate these results in patient-derived human BCR-ABL1 B-ALL cells, we developed a CRISPR-based approach for genetic deletion of CD25 in B-ALL PDX. CRISPR-based deletion of CD25 compromised the colony formation capacity of BCR-ABL1 B-ALL PDX cells, reducing colony numbers by ~50-fold (Fig. 2h). In a complementary approach, we transduced a human B-ALL cell line (MUTZ5) with oncogenic JAK2^R683G and IGH@CRLF2 with 4-OHT-inducible iCas9 and RFP-labeled guide RNAs (gRNAs) for CRISPR-mediated deletion of CD25 or with non-targeting controls. Similar to mouse models of tyrosine kinase–driven B-ALL and AML (Fig. 2a–d), inducible deletion of CD25 in human MUTZ5 B-ALL cells resulted in rapid depletion in competitive cell culture assays (Fig. 2i). The Dependency Map (DepMap) project includes CD25 CRISPR-deletion data for 12 B-ALL and 35 AML cell lines. These cell lines did not exhibit significant CRISPR-scores for CD25 deletion; however, none of the 12 B-ALL and only 3 of 35 AML cell lines carried known oncogenic tyrosine kinases⁴³. Moreover, CRISPR-scores for ABL1, JAK2, JAK3, and FLT3 were negative in all 12 B-ALL and were positive in only 1 of 35 (ABL1, FLT3, JAK3) and 3 of 35 (JAK2) AML cell lines⁴³, suggesting that tyrosine kinase–related dependencies in B-ALL and AML cells are not reliably detected in DepMap datasets.

CD25 functions as negative feedback regulator of NF-κB signaling

CD25⁺ subpopulations in acute leukemia showed increased NF-κB activity, and NF-κB activation induced transcriptional activation and surface expression of CD25 (Fig. 1i–l; fig. S3b–d). Studying transcriptional changes upon deletion of CD25 in BCR-ABL1 B-ALL and MLL-AF9 FLT3^ITD AML cells revealed strong activation of NF-κB feedback inhibitors, including Nr4a1, Nfkbia, Nfkbie, and Nfkbiz (Fig. 3a–b), consistent with CD25-mediated feedback control of NF-κB signaling in tyrosine kinase–driven B-ALL and AML. Inducible deletion of CD25 in BCR-ABL1 B-ALL and MLL-AF9 FLT3^ITD AML cells increased NF-κB pathway activation, including phosphorylation of PKCβ, RelA (p65), and IκBα (Fig. 3c, fig. S9a). Together, these findings suggest a negative feedback loop in which NF-κB signaling induces transcriptional activation of CD25 (Fig. 1i–l), which in turn is required to negatively regulate NF-κB activity (Fig. 3a–c). Closer inspection of RNA-seq data provided direct evidence supporting a role of CD25 as an essential negative feedback regulator of NF-κB. Consistent with loxP sites flanking exons 2 and 3 of Il2ra (which encodes CD25), RNA-seq analysis revealed a non-productive CD25 transcript lacking exons 2 to 3 (Fig. 3d). The non-productive CD25 transcript was among the mRNAs that was most prominently increased in abundance upon CD25 deletion in BCR-ABL1 B-ALL cells (Fig. 3a). Consistent with a role for CD25 in a negative feedback loop, loss of CD25 function resulted in excessive NF-κB activation (Fig. 3a–c), which in turn induced increased RNA-seq counts of the non-functional CD25 transcript (Fig. 3d). NF-κB functions as a tumor suppressor during early B cell development and prevents the development of tyrosine kinase–driven B-ALL⁴⁴. To assess whether increased NF-κB activation mechanistically contributed to the death of BCR-ABL1 B-ALL cells upon CD25 deletion, we leveraged the MALT1 paracaspase small-molecule inhibitor MI-2 to attenuate NF-κB signaling⁴⁵. Indeed, treatment with MI-2 largely mitigated the toxicity of CD25 deletion in BCR-ABL1 B-ALL cells (Fig. 3e).

Fig. 3. — (A and B) RNA sequencing was performed with BCR-ABL1–transformed *Cd25*^fl/fl B-ALL cells (n=3) expressing 4-OHT-inducible Cre-ER^T2 or ER^T2 after treatment with 4-OHT for 2 days. Relative rlog-normalized gene expression values for differentially expressed genes (P<0.05 and log₂-transformed fold change >1; Wald test with Benjamini–Hochberg correction) were plotted as a heat map with row scaling. Increased abundance of a truncated *Il2ra* (*Cd25*) transcript is highlighted in red (A). Gene set enrichment analysis (NF-κB-induced, NF-κB-repressed, NFATC1 targets, NFATC2 targets) for genes ranked by ratio of 4-OHT to vehicle as log₂-transformed fold change. Red lines indicate running enrichment score (right axis); gray bars indicate log₂ fold change (left axis). Statistical significance was determined by two-tailed Kolmogorov-Smirnov test (B). (C) *Cd25*^fl/fl BCR-ABL1 B-ALL cells and *Cd25*^fl/fl MLL-AF9 and FLT3^ITD AML cells expressing 4-OHT-inducible Cre-ER^T2 or ER^T2 were treated with 4-OHT for 2 days for inducible deletion of CD25. Cell lysates from B-ALL and AML cells with and without CD25 deletion were analyzed by Western blotting for the abundance of NF-κB signaling–related proteins, NF-κB p65-pSer⁵³⁶, NF-κB p65, IκBα-pSer^32/36, and IκBα. β-actin was used as loading control. Blots represent three biological replicates for each cell type and condition. Densitometric analysis of bands of interest are shown in fig. S9. (D) RNA-seq reads aligned to the *Cd25* locus; loxP sites flanking exons 2 and 3 are highlighted in *Cd25*^fl/fl *BCR-ABL1*-transformed B-ALL cells expressing 4-OHT-inducible Cre-ER^T2. (E) *Cd25*^fl/fl BCR-ABL1 B-ALL cells were transduced with retroviruses to express 4-OHT-inducible Cre-ER^T2-GFP or ER^T2-GFP empty vector control. Percentages of GFP⁺ cells were measured by flow cytometry at different times after 4-OHT treatment in the presence or absence of the MALT1 paracaspase small-molecule inhibitor MI-2 (500 nM) to attenuate NF-κB signaling³⁸ or with vehicle as a control. Data are representative of three biological replicates. (F) Autonomous Ca²⁺ flux was assessed by green fluorescent Ca²⁺ indicator (GCaMP6s) upon inducible deletion of *CD25* with 4-OHT treatment in *Cd25*^fl/fl BCR-ABL1 B-ALL cells. Top: Representative time-course plots from a single cell for 10 min with a 6-s time interval (x-axis). Bottom: Fold-changes in GCaMP6s intensity from 50 single cells are shown as a heat-map. (G) NFATC2 localization was analyzed by immunofluorescence with NFATC2 and DAPI staining upon 4-OHT-induced deletion of CD25 in BCR-ABL1-transformed *Cd25*^fl/fl B-ALL cells. Scale bars, 22 μm (top), and 10 μm (bottom).

CD25 inhibits Ca²⁺ oscillations and NFAT signaling in tyrosine kinase–driven B-ALL

In addition to affecting NF-κB, deletion of CD25 also resulted in selective activation of NFAT (NFATC1, NFATC2) target genes, including Ccl3, Egr2, and Egr3, as well as NFAT transcriptional programs (Fig. 3a–b). Because NFAT activation is typically the result of increased Ca²⁺ signaling⁴⁶, we engineered CD25^fl/fl B-ALL cells with the GCaMP6s Ca²⁺ biosensor³⁰ and Cre-ER^T2 for inducible deletion of CD25. Genetic deletion of CD25 in B-ALL cells resulted in autonomous, high-frequency (~20 mHz) Ca²⁺ oscillations (Fig. 3f). Immunofluorescence experiments showed that deletion of CD25 and high-frequency Ca²⁺ oscillations indeed increased nuclear NFATC2 abundance (Fig. 3g). These findings are consistent with our previous observation that the induction of BCR-ABL1 expression results in oncogenic tyrosine kinase activity that is concentrated in a CD25⁺ leukemia subpopulation with suppressed Ca²⁺ signaling (Fig. 1d–e). Together, these results suggest that CD25 serves a previously unrecognized role in negative feedback control of Ca²⁺, NF-κB, and NFAT signaling downstream of oncogenic tyrosine kinases in acute leukemias.

CD25 is required for the activation of inhibitory phosphatases to curb oncogenic tyrosine kinase signaling

To elucidate how CD25 interacted with oncogenic tyrosine kinase signaling, we performed global phosphoproteomic and mass spectrometry experiments to identify changes in tyrosine and serine/threonine phosphorylation upon CD25 deletion in BCR-ABL1 B-ALL cells. The most prominent increases in the extent of tyrosine phosphorylation were observed among known proximal substrates of BCR-ABL1, including the ABL1, FYN, JAK1, JAK2, JAK3, and SYK tyrosine kinases, as well as STAT5A and STAT5B. Among the greatest increases in the extent of serine/threonine phosphorylation were multiple sites of PKCδ, the only inhibitory member of the PKC family^47–51. In addition, the inhibitory phosphatase (INPP5D) scaffolds CIN85⁵² and SASH3⁵³, NFKB1 and the NF-κB activator RLTPR (CARMIL2)⁵⁴ were heavily phosphorylated upon deletion of CD25 (Fig. 4a–b). Moreover, phosphorylation of residues that are indicative of phosphatase activation was most prominently lost in response to CD25 deletion. Loss of phosphorylation events included the ITIMs of the inhibitory receptors CD22 and BTLA, which serve as binding partners of SH2 domains of the inhibitory phosphatases INPP5D and PTPN6 to initiate their activation^55–56. In addition, the PKCδ adaptor RACK1, the inhibitory INPP5D adaptors DOK1, DOK3, as well as SH3BP5, the inhibitor of the tyrosine kinase BTK were among the most prominent proteins that lost tyrosine phosphorylation upon deletion of CD25 (Fig. 4a–b). These results suggest that Cre-mediated deletion of CD25 caused major imbalances between acute increases of activators of tyrosine kinase and NF-κB-signaling and loss of inhibitory INPP5D and PTPN6 signaling. Previously, we and others showed that B-ALL cells critically rely on the inhibitory phosphatases INPP5D and PTPN6 to curb oncogenic signaling^26–28. These results suggest a previously unrecognized CD25-dependent feedback loop for the recruitment and activation of INPP5D and PTPN6 to balance the signaling strength of oncogenic tyrosine kinases. To validate the phosphoproteomics results, we examined the functional consequences of CD25 deletion on phosphatase activation (INPP5D, PTPN6), PKCδ, and tyrosine kinase signaling in BCR-ABL1 B-ALL by Western blotting analysis. Induced deletion of CD25 substantially decreased the extent of phosphorylation of INPP5D and PTPN6, consistent with defective phosphatase activation. In contrast, the tyrosine kinases SYK and BTK, their linker BLNK, and PKCδ were markedly increased in phosphorylation (Fig. 4c, fig. S9b).

Fig. 4. — (A and B) *Cd25*^fl/fl *Mb1*-Cre-ER^T2 B-ALL cells transformed with BCR-ABL1 were treated with 4-OHT or vehicle for 2 days (n=4). The relative amounts of phosphorylated proteins, as determined by mass spectrometry, with phospho-tyrosine residues (A) or phospho-serine/threonine residues (B) were plotted by log₂-transformed fold change (moderated t-statistic). (C) *Cd25*^fl/fl pre-B cells and BCR-ABL1 B-ALL cells expressing 4-OHT-inducible Cre-ER^T2 or ER^T2 were treated with 4-OHT for 2 days for inducible deletion of CD25. Cell lysates from pre-B and B-ALL cells with and without CD25 deletion were analyzed by Western blotting for phosphorylation of inhibitory phosphatases (Inpp5d-pY¹⁰²⁰, Inpp5d, Ptpn6-pY⁵⁶⁴, Ptpn6), as well as Inpp5d- and Ptpn6-downstream substrates Blnk-pTyr⁹⁶, Blnk, Btk-pTyr²²³, and Btk. β-actin was used as a loading control. Blots are representative of three biological replicates. (D) Patient-derived B-ALL xenografts, PDX2 harboring BCR-ABL1, were electroporated to introduce Cas9 RNPs containing guide RNAs for CRISPR-mediated deletion of CD25 (gCD25) or nontargeting control (gNT). Abundances of ERK1/2-pThr²⁰²/Tyr²⁰⁴, ERK1/2, IκBα-pSer^32/36 and IκBα were assessed by Western blotting analysis, using β-actin as a loading control. Blots represent three biological replicates. (E) Ca²⁺ mobilization was assessed by GCaMP6s upon CRISPR-Cas9-mediated deletion of *CD25* (gCD25) in patient-derived B-ALL (PDX2) cells. gNT was used as guide RNA control. Top: Representative time-course plots from a single cell for 30 min with a 20-s time interval (x-axis). Bottom: Fold-changes in GCaMP6s intensity from 50 single cells are shown as a heat-map. (F) For Bio-ID interactome analyses, the BirA biotin ligase was fused to the C terminus of CD25. CD25-BirA and BirA control were overexpressed in patient-derived Ph⁺ B-ALL (PDX2) cells. Twenty-four hours after CRISPR-Cas9-mediated deletion of endogenous CD25, PDX2 B-ALL cells were incubated with 50 μM biotin in culture medium for 10 min, and biotin-labeled CD25-interactomes were analyzed by mass spectrometry. Log₂-fold enrichment (x-axis) and significance (y-axis) of CD25-BirA vs BirA control in PDX2 cells were plotted (n=4; moderated t-statistic). (G) Co-immunoprecipitation was performed in patient-derived Ph⁺ B-ALL (PDX2) cells using anti-CD25 antibodies covalently cross-linked with protein A/G magnetic beads. CD25-interacting proteins were assessed by Western blotting analysis. Whole cell lysates (WCLs) were used as positive controls for immunoblots. The isotype control for CD25 antibody (IgG) was used as a negative control. AKT and p38 were used as specificity controls. Densitometric analysis of bands of interest in the representative Western blots are shown in fig. S9.

Whereas deletion of CD25 in FLT3^ITD MLL-AF9 AML cells resulted in loss of colony formation and competitive fitness, this was not the case in AML cells that were transformed by MLL-AF9 alone (fig. S5a–d). Similarly, CD25 deletion introduced signaling imbalances in FLT3^ITD MLL-AF9 AML cells but not in AML cells transformed by MLL-AF9 alone (fig. S5e–f, fig. S9g–h). These results suggest that CD25 is not only selectively expressed in tyrosine kinase–driven AML, but also selectively required for feedback control of oncogenic tyrosine kinase signaling. To determine whether CD25 had a similar inhibitory function in the feedback control of oncogenic tyrosine kinase signaling in patient-derived leukemia cells, we introduced CRISPR-mediated deletion of CD25 in BCR-ABL1 B-ALL cells from patient-derived xenografts (PDX; Fig. 4d–g). Consistent with experiments with murine BCR-ABL1 B-ALL cells (Fig. 3c, Fig. 4a–c), CRISPR-mediated deletion of CD25 in human BCR-ABL1 B-ALL PDX resulted in the enhanced activation of ERK and NF-κB signaling, which was reflected by increased phosphorylation and degradation of IκBα (Fig. 4d, fig. S9c). Corroborating results in murine models of BCR-ABL1 B-ALL (Fig. 3f), deletion of CD25 in human B-ALL PDX also resulted in autonomous Ca²⁺ oscillations (Fig. 4e).

CD25 interacts with regulators of tyrosine kinase signaling in BCR-ABL1 B-ALL

To determine how CD25 enabled the recruitment and activation of INPP5D and PTPN6 to curb tyrosine kinase-signaling, we performed Bio-ID proximity labeling experiments in engineered human BCR-ABL1 B-ALL (PDX2) that did or did not express CD25. To prevent residual WT cells from outcompeting them, CD25^−/− PDX2 cells were sorted and then reconstituted with fusions between the CD25 cytoplasmic tail and the biotin ligase BirA for labeling, streptavidin-pulldown, and mass spectrometry identification of CD25-interacting proteins. The CD25 interactomes included the tyrosine kinases JAK1, LYN, and BTK, the phosphatases INPP5D, PTPN6, the phosphatase scaffold SASH3⁵², and CD22, which contains ITIMs that are required for INPP5D and PTPN6activation^54–55, as well as the B cell signaling molecules CD19, RFTN1, and BLNK (Fig. 4f). Interactions between CD25 and the Ca²⁺ flux regulators LRMP (IRAG2)⁵⁷ and STIM1⁵⁸ were of interest given that deletion of CD25 resulted in autonomous Ca²⁺ oscillations (Fig. 3f, Fig. 4e). Interactions between CD25 and PKCδ, RACK1, LYN and JAK1, INPP5D, and STIM1 were confirmed by co-immunoprecipitation and Western blotting analysis (Fig. 4g).

A motif in the cytoplasmic tail of PKCδ is critical for stable CD25 surface expression

To elucidate the mechanism by which CD25 exerted feedback control of tyrosine kinase signaling in tyrosine kinase–driven B-ALL and AML cells, we examined the role of its 13–amino acid residue cytoplasmic tail (Fig. 5a). The CD25 tail includes two highly conserved S/T phosphorylation sites (Ser²⁶⁸ and Thr²⁷¹) that were previously identified as targets for phosphorylation by PKC⁵⁹; however, the importance of CD25-Ser²⁶⁸/Thr²⁷¹ phosphorylation is unclear. To identify upstream kinases that phosphorylated CD25-Ser²⁶⁸/Thr²⁷¹, we performed in vitro kinase assays with 60 serine/threonine kinases and a 13-amino acid residue peptide corresponding to the cytoplasmic tail of CD25 including Ser²⁶⁸ and Thr²⁷¹ (CD25-WT), as well as an S268A mutant peptide (CD25-S268A). We identified multiple PKC family members and PKCδ as the most selective for CD25-WT over CD25-S268A (Fig. 5b–c). Proteomic (reverse-phase protein array, RPPA) datasets for 208 B-ALL samples in the MDACC ALL360 cohort⁶⁰ showed significant correlations with CD25 flow cytometry and PKCδ phosphorylation (PKCδ-pSer⁶⁶⁴ and pSer⁶⁴⁵; r=0.38, P=5.8E-04; Fig. 5d; fig. S6a). Whereas most B-ALL cases lacked both CD25 expression and PKCδ phosphorylation, the correlation was mainly based on a small group of B-ALL cases with BCR-ABL1 and JAK2 mutations (Fig. 5d).

Fig. 5. — (A) Alignment of PKC substrate residues with the cytoplasmic tail of CD25 (amino acid residues 261 to 272). (B and C) In vitro kinase assays of 62 known serine/threonine kinases against the recombinant cytoplasmic tail of wild-type (WT) CD25 (WQRRQRKS²⁶⁸RRTI) or the S268A mutant (WQRRQRKA²⁶⁸RRTI) were performed. The percentages of kinase efficiency were quantitated as the percentage of remaining ³³P ATP (n=2). The relative kinase activity of each individual kinase is represented as a bar chart comparing WT and S268A-mutant (B) and as a volcano plot (C) for fold-differences in kinase activity against the WT and S268A mutant. The gray bars are stacked on top of the red bars. (D) Reverse-phase protein assay (RPPA) dataset of PKCδ phosphorylation at Ser⁶⁶⁴ (S664) and Ser⁶⁴⁵ (S645) were analyzed on a cohort of adult B-ALL patient samples from MDACC ALL360 (n=208). B-ALL samples with BCR-ABL1 and JAK2 lesions are indicated as red circles, whereas other B-ALL are indicated with gray circles. Correlation between PKCδ-pSer^664/645 and CD25 abundance was determined by flow cytometric analysis and shown as dot plots. Linear regression is indicated as a red line. (E) Ph⁺ B-ALL PDX cells (PDX2) were treated with 5 nM I3A (a PKCδ-selective agonist) or vehicle (DMSO) for 6 and 12 hours. CD25 surface and intracellular staining were performed in parallel for each sample. Flow cytometry plots and CD25 MFI values are representative of three biological replicates. (F) Patient-derived B-ALL with BCR-ABL1 (PDX2) and CRLF2-rearranged, JAK2^R683S (NCL1) or *FLT3*^ITD AML (PDX21) xenografts were treated with 1 and 5 nM I3A or vehicle control for an hour. Western blotting analysis was performed to detect the relative amounts of PKCδ-pThr⁵⁰⁵, PKCδ, CD25-pSer²⁶⁸, CD25, ERK1/2-pThr²⁰²/Tyr²⁰⁴, and ERK1/2. β-actin was used as a loading control. (G to I) PKCδ was deleted in BCR-ABL1 B-ALL (PDX2) and CRLF2-rearranged, JAK2^R683S (NCL1) xenografts by CRISPR-Cas9 (gPRKCD), and a non-targeting guide (gNT) was used as control. PDX2 cells were subsequently treated with 5 nM I3A or DMSO for 6 hours. CD25 surface and intracellular staining were performed in parallel for each sample. Representative flow cytometry plots (G) and fold changes in CD25 MFI values (H) are shown. Solid lines in (H) indicate CD25 staining for *BCR-ABL1*⁺ PDX2 cells, whereas dashed lines indicate CD25 staining for *JAK2*^R683S NCL1 cells. (I) Western blotting analysis of the relative amounts of PKCδ-pThr⁵⁰⁵, PKCδ, CD25-pSer²⁶⁸, and CD25 in PDX2 cells upon treatment with the indicated concentrations of I3A or vehicle (DMSO) for 1 hour. β-actin was used as a loading control. (J) Family pedigree with two different *IL2RA* (CD25) mutations, Trp¹⁷⁷* (blue, resulting in a stop codon at Trp¹⁷⁷) and K267Rfs* (red, resulting in a frameshift mutation preceding Ser²⁶⁸). Three compound heterozygous siblings are depicted with clinical phenotypes (T1D, Type 1 Diabetes; ICA, Islet Cell Antibodies; ITP, immune thrombocytopenic purpura). CD25 expression in primary T cells from healthy (green) and compound heterozygous (red) donors after CD3/CD28 stimulation was compared by flow cytometry. CD25 expression was analyzed two days after deletion (g-CD25) of K267Rfs* allele (black), and a nontargeting guide RNA was used as control (g-NT; red; middle). CD25 expression was assessed after HDRT-mediated correction of K267Rfs* frameshift mutation (blue; right). CD25 MFIs (× 10³) in CD4⁺ populations are shown. (K) Murine *Prkcd*^+/+ and *Prkcd*^−/− BCR-ABL1 B-ALL cells were plated in semi-solid methylcellulose for colony-forming assay. Microscopic images and colony numbers are representative of three biological replicates. Scale bar, 7 mm (top) and 2.5 mm (bottom). (L) *Prkcd*^+/+ and *Prkcd*^−/− pre-B cells were transduced with retroviruses to express GFP-tagged inducible BCR-ABL1 or empty vector (EV-GFP). Changes in the numbers of GFP⁺ cells upon acute activation of BCR-ABL1 were monitored by flow cytometry, and fold-changes in GFP⁺ populations were plotted. Data are represents of three biological replicates. Statistical significance was assessed by two-tailed t-test. Densitometric analysis of bands of interest in the Western blots are shown in fig. S9.

Because PKCδ is the only PKC family member that negatively regulates immune cell signaling^46–50, we examined whether PKCδ contributed to CD25-mediated negative feedback control of oncogenic tyrosine kinase signaling in leukemia cells. Treatment of patient-derived B-ALL (BCR-ABL1, JAK2^R683S, FLT3^D835Y) and AML (FLT3^ITD) xenografts with the PKCδ-selective agonist ingenol 3-angelate (I3A)⁶¹ induced PKCδ activation, CD25-tail phosphorylation (Ser²⁶⁸), and the accumulation of CD25 at the cell surface (Fig. 5f–g, fig. S6b, fig. S9i), suggesting that PKCδ was sufficient to induce phosphorylation and cell surface expression of CD25. CRISPR-mediated deletion of PKCδ (PRKCD) in two B-ALL PDX (BCR-ABL1, JAK2^R683S) substantially reduced both CD25 protein abundance under basal conditions and CD25-tail phosphorylation (Ser²⁶⁸; Fig. 5g–I, fig. S9e). This result supports the phosphorylation of the CD25-tail being mediated by PKCδ rather than another PKC family members and that PKCδ is required for the stable expression of CD25.

PKCδ-mediated stabilization of CD25 is required for leukemogenesis

Previous studies showed that individuals with inherited PKCδ (PRKCD) germline point mutations suffer from systemic autoimmunity and B cell hyperactivation^47–48, as do patients with inherited CD25 mutations^62–63. In a family with inherited biallelic CD25 germline mutations⁶⁴, a frameshift mutation (K267Rfs*) immediately preceding the PKCδ-phosphorylation site (Ser²⁶⁸, Thr²⁷¹) was associated with loss of CD25 surface expression, whereas CRISPR-mediated correction of the PKCδ-motif restored the cell surface expression of CD25 (Fig. 5j). The critical function of PKCδ in negatively regulating B cell signaling was previously recognized^49–51; however the mechanistic basis of PKCδ-mediated signaling inhibition was not clear. These results suggest that PKCδ exerts negative regulation of immune cell and oncogenic tyrosine kinase signaling through the phosphorylation and stabilization of CD25. We showed earlier that CD25 function is essential for competitive fitness, colony formation, and leukemia initiation of tyrosine kinase–driven B-ALL (BCR-ABL1, PAX5-JAK2, and EBF1-PDGFRB) and AML (FLT3^ITD; Fig. 2). Consistent with a critical role of PKCδ in the phosphorylation and stabilization of CD25, Prkcd^−/− progenitor cells were resistant to BCR-ABL1–mediated transformation and failed to form colonies or initiate the development of cytokine-independent leukemia (Fig. 5k–l).

Structural basis of the CD25-dependent recruitment and activation of inhibitory phosphatases

Based on these findings, we tested the hypothesis that CD25 function and its ability to activate INPP5D and PTPN6 depended on PKCδ-mediated phosphorylation. This scenario would be consistent with previous studies of mast cells and platelets, suggesting that PKCδ plays an important role in activating inhibitory phosphatases^65–66. We examined whether PKCδ-mediated phosphorylation of CD25 at Ser²⁶⁸/Thr²⁷¹ enabled the recruitment of INPP5D and PTPN6 or other components of a CD25-dependent inhibitory complex. To investigate Ser²⁶⁸/Thr²⁷¹-dependent interactions of CD25, we performed Bio-ID proximity labeling experiments in CD25^−/− PDX2 (BCR-ABL1 B-ALL) cells. After deletion of CD25, PDX2 cells were reconstituted with fusions between intact CD25 (CD25-ST) or S268A/T271V mutant CD25 (CD25-AV) and the biotin ligase BirA for labeling, streptavidin-pulldown, and mass spectrometry–based identification of interacting proteins. We used the PKCδ-agonist I3A⁶¹ to induce PKCδ-mediated phosphorylation of CD25 at Ser²⁶⁸/Thr²⁷¹ (Fig. 5f, fig. S9d). Comparing Bio-ID interactomes for CD25-ST and CD25-AV, we identified INPP5D, PTPN6, and the PKCδ-scaffold protein RACK1 among the CD25-interacting proteins that were most selective for CD25-ST as compared to the CD25-AV mutant (Fig. 6a, fig. S7a). Treatment with the PKCδ-agonist I3A increased the CD25-ST–selective interactions between the cytoplasmic tail of CD25 and the inhibitory phosphatases PTPN6, INPP6D, and PTPN1, the inhibitory receptor CD22, and the PKCδ-scaffold RACK1 (Fig. 6a, fig. S7b).

Fig. 6. — (A) For Bio-ID interactome analyses, the BirA biotin ligase was fused to the C terminus of WT (CD25-ST) and mutant (CD25-AV) CD25. CD25^WT-BirA, CD25^S268A/T271V-BirA, and BirA-EV control were reconstituted in *CD25*-deficient B-ALL PDX (PDX2) cells. Cells were incubated with 50 μM exogenous biotin in culture medium for 10 min upon treatment with I3A. Log₂-fold enrichment (x-axis) and significance (y-axis) of CD25^WT-BirA vs CD25^S268A/T271V-BirA were plotted (n=4; moderated t-statistic). (B) CD25^−/− PDX2 B-ALL cells were reconstituted with CD25-ST or CD25-AV and subjected to co-immunoprecipitation with anti-CD25 antibody followed by Western blotting analysis to validate the CD25-interacting proteins identified from the Bio-ID interactome analyses. WCL was used as a positive control. β-actin was used as a specificity control. (C and D) The proximity of CD25 to INPP5D (C) or PTPN6 (D) was assessed by in situ PLA in CD25^−/− PDX2 B-ALL cells reconstituted with CD25-ST or CD25-AV. Cells were treated with 5 nM I3A or vehicle for 10 min. The PLA signal (red), DAPI (blue), and wheat germ agglutinin (green) are shown. Representative microscopic images are shown. Scale bar, 5 μm. (E to G) Chimeric CD19:CD25 chains were constructed by fusing the extracellular domain of mouse CD19 (amino acid residues 1 to 287) to the transmembrane domain and cytoplasmic tail of mouse CD25 (ST or AV) with a GFP tag. Empty vector (EV) or mouse CD19 (CD19) were used as controls. (E) Murine *Cd25*^−/− *BCR-ABL1* B-ALL cells reconstituted with wild-type CD25, chimeric CD19:CD25 chains, or empty vector (EV) were plated in semi-solid methylcellulose for colony-forming assays. Microscopic images and colony numbers are representative of three biological replicates. Scale bar, 7 mm (top), 2.5 mm (bottom). Statistical significance was assessed by two-tailed t-test (right). (F) Murine *Cd25*^−/− *BCR-ABL1* B-ALL cells were reconstituted with GFP-tagged WT CD25, chimeric CD19:CD25 chains, or empty vector (EV). Enrichment of GFP⁺ cells (means ± SD) was monitored by flow cytometry. Data are representative of three biological replicates. (G) The relative abundances of Inpp5d-pTyr¹⁰²⁰, Inpp5d, Ptpn6-pTyr⁵⁶⁴, Ptpn6, and Rack1 were assessed by Western blotting analysis of *Cd25*^−/− *BCR-ABL1* B-ALL cells reconstituted with the indicated constructs. β-actin was used as loading control. Blots represent three biological replicates. Densitometric analysis of bands of interest in the Western blots are shown in fig. S9.

To corroborate that these interactions depended on an intact CD25 tail and the PKCδ-phosphorylation site (Ser²⁶⁸/Thr²⁷¹; CD25-ST), we performed co-immunoprecipitation experiments. To this end, CD25-deficient PDX2 B-ALL cells were reconstituted with CD25-ST or CD25-AV. Western blotting analyses with anti-CD25 antibodies confirmed that interactions between CD25 and INPP5D, PTPN6, PKCδ, and RACK1 were dependent on Ser²⁶⁸/Thr²⁷¹ (CD25-ST) and were abolished by the CD25-AV mutation (Fig. 6b). These results were further confirmed in proximity ligation assays (PLAs) for INPP5D and PTPN6 in PDX2 B-ALL cells expressing CD25-ST- or CD25-AV. Low basal interactions between CD25-ST and INPP5D and PTPN6 were increased by treatment with I3A (Fig. 6c–d). However, no substantial interactions between the CD25-AV mutant and INPP5D or PTPN6 were detected (Fig. 6c–d).

The cytoplasmic tail of CD25 is required for the recruitment and activation of inhibitory phosphatases

To study the mechanistic role of the cytoplasmic tail of CD25, we generated chimeric constructs containing the transmembrane domain (residues 239 to 259) and the cytoplasmic tail (residues 260 to 272) of CD25 fused to the extracellular domain (ECD) of CD19, a surface receptor that is constitutively expressed on B-ALL cells (Fig. 6e–h). Thus, we reconstituted CD25-deficient murine B-ALL cells with constructs for full-length CD25, full-length CD19, and chimeric constructs between the CD19-ECD and the intact (CD25-ST) or S268A/T271V mutant (CD25-AV) CD25 tail. The expression of full-length CD19 had no marked effects; however, fusion of the cytoplasmic tail CD25-ST to CD19-ECD was sufficient to rescue colony-forming capacity and competitive fitness in CD25^−/− B-ALL cells (Fig. 6e–f). In contrast, the fusion between CD19-ECD and mutant CD25-tail (CD25-AV) failed to confer a survival advantage or to restore colony formation capacity (Fig. 6e–f). Similarly, CD25-ST, but not the CD25-AV mutant, restored the phosphorylation and activation of INPP5D and PTPN6 and stable expression of RACK1 (Fig. 6g, fig. S9f). These results suggest that the cytoplasmic tail of CD25 is sufficient (albeit when the Ser²⁶⁸/Thr²⁷¹ motif is intact), to restore survival, colony formation, activation of INPP5D and PTPN6, and stable expression of RACK1 in Cd25^−/− murine B-ALL cells.

Because inducible deletion of CD25 in human tyrosine kinase–driven B-ALL cells resulted in loss of competitive fitness (Fig. 2i), we examined whether the PKCδ-phosphorylatable Ser²⁶⁸/Thr²⁷¹ motif was required to restore functional defects in CD25-deficient B-ALL cells. To this end, we deleted CD25 in human B-ALL cells carrying the BCR-ABL1 tyrosine kinase (SUP-B15 cell line and MXP5 PDX) by electroporation with CRISPR-Cas9 RNPs and CD25-targeting guide RNAs. CD25^−/− B-ALL cells were then transduced to express GFP-tagged, doxycycline-inducible Tet-On CD25-ST (wild-type), CD25-AV (S268A/T271V mutant), or GFP empty vector control. As expected, inducible reconstitution of the cells with wild-type CD25 restored competitive fitness (Fig. 7a–b) and colony formation capacity in human B-ALL cells (Fig. 7c). Inducible expression of the CD25-AV mutant in CD25^−/− B-ALL cells, however, did not confer a substantial competitive advantage and failed to increase colony formation (Fig. 7a–c). Hence, these results with human B-ALL cells confirm findings from a murine B-ALL model (Fig. 6e–g) and highlight the central importance of the PKCδ target RRWRKS²⁶⁸RRT²⁷¹ motif in maintaining competitive fitness and colony formation capacity in B-ALL cells. Similar to results from experiments with tyrosine kinase–driven B-lineage ALL, reconstitution of myeloid-lineage CD25^−/− MLL-AF9 FLT3^ITD AML cells with inducible wildtype CD25-ST, but not the CD25-AV mutant, restored colony formation (fig. S7c).

Fig. 7. — (A and B) SUP-B15 cells (A) and MXP5 (B) PDX BCR-ABL1 B-ALL cells were electroporated with RNPs of Cas9 and CD25-targeting guide RNAs. CD25^−/− B-ALL cells were enriched by flow sorting and then transduced with lentiviruses to express GFP-tagged, doxycycline-inducible (Tet-On) CD25-ST, CD25-AV, or GFP empty vector control. Expression of the three constructs was induced by doxycycline, and the percentages of GFP⁺ cells were monitored at the indicated times by flow cytometry. Data from three biological replicates are shown. (C) CD25^−/− SUP-B15 cells were transduced with lentiviruses to express GFP-tagged, doxycycline-inducible (Tet-On) CD25-ST, CD25-AV, or GFP empty vector control. Cells were flow sorted after doxycycline treatment to select GFP⁺ cells. Then, 20,000 sorted GFP⁺ cells were plated in semi-solid methylcellulose medium, and colonies were counted after 10 days. Microscopic images are representative of three biological replicates. Scale bar, 7 mm. Statistical differences were assessed by two-tailed t-test.

Activation of the PKCδ scaffold protein RACK1 can rescue the loss of CD25 function

Our Bio-ID interactome and co-immunoprecipitation studies confirmed that CD25 can interact with RACK1 and that these interactions depend on the PKCδ phosphorylation site in the tail of CD25 (Fig. 6a–b; fig. S7a–b). To corroborate the CD25-RACK1 interaction and its dependency on an intact CD25-ST tail, we performed PLAs. Under basal conditions, RACK1 interacted with the intact CD25-ST but not with the CD25-AV mutant. Treatment with the PKCδ agonist I3A significantly increased the PLA signals (Fig. 8a). Previous work suggested that upon PKCδ-mediated phosphorylation, RACK1 can recruit inhibitory phosphatases and induce their activation^67–69. As a result of these interactions, RACK1 functions as an inhibitory scaffold to negatively regulate Src and other tyrosine kinases^70–72. Given the ability of RACK1 to negatively regulate tyrosine kinases, we assessed whether the assembly of RACK1 complexes at the plasma membrane could rescue defective feedback control of oncogenic tyrosine kinase signaling in CD25-deficient leukemia cells. For this reason, we engineered membrane-tethered (myristoylated) RACK1 in CD25^−/− BCR-ABL1-driven B-ALL cells. Membrane-tethered RACK1 restored competitive fitness and the colony formation capability of CD25^−/− B-ALL cells to a similar extent as did reconstitution of the cells with CD25 (Fig. 8b–c). These results suggest that CD25 orchestrates negative feedback regulation and recruitment of inhibitory phosphatases in BCR-ABL1 B-ALL cells, in part, by facilitating interactions between PKCδ and its inhibitory scaffold RACK1.

Fig. 8. — (A) The proximity of CD25 to RACK1 in CD25^−/− PDX2 B-ALL cells reconstituted with CD25-ST or CD25-AV was assessed by in situ PLA after treatment with 5 nM I3A or vehicle for 10 min. The PLA signal (red), DAPI (blue), and wheat germ agglutinin (green) are shown. Representative microscopic images are shown. Scale bar, 5 μm. Quantitative analyses are shown as PLA signals per cell of each sample with the mean values shown as a red line. Statistical significance was assessed by two-tailed t-test. (B and C) Murine *Cd25*^−/− *BCR-ABL1* B-ALL cells were reconstituted with constitutively active (myristoylated) RACK1-GFP or empty vector (EV). (B) The abundance of the PKCδ scaffold RACK1 was validated by intracellular flow cytometry (left). Enrichment of GFP⁺ cells was monitored by flow cytometry (right). (C) Murine *Cd25*^−/− *BCR-ABL1* B-ALL cells reconstituted with wildtype CD25, EV, or constitutively active (myristoylated) RACK1 were plated in semi-solid methylcellulose for colony-forming assays. Microscopic images and colony numbers are representative of three biological replicates. Scale bar, 7 mm (top), 2.5 mm (bottom). Statistical significance was assessed by two-tailed t-test. (D) Representative structures are shown as surface representations for the ternary complexes between CD25 (ST or AV, pink), PKCδ (blue), and RACK1 (gray). (E) The ratios of population densities of stable (Assembled) versus less stable conformational states (Disassembled) in MD simulations of CD25-ST- or CD25-AV-PKCδ-RACK1 ternary complexes were plotted. (F and G) Detailed PKCδ-RACK1 interaction interface for stable states (left) and the initial step of weakening of the interface interactions transitioning to less stable states (right) are shown. (G) Dashed lines label all the contacts which were persistent in >60% of the simulation time. (H) Combined interaction energy between PKCδ-CD25, CD25-RACK1, and PKCδ-RACK1 in CD25-ST- or CD25-AV-PKCδ-RACK1 ternary complexes is shown as a violin plot. Statistical significance was assessed by two-tailed t-test.

Structural modeling of the PKCδ:CD25:RACK1 ternary complex

Previous work showed that the activation and membrane-anchoring of PKCδ depends on its scaffold RACK1, which stabilizes PKCδ and its substrates^73–74. Consistent with a role for RACK1 in stabilizing PKCδ and its substrate CD25, our co-immunoprecipitation experiments showed that CD25 interacted with both PKCδ and RACK1 (Fig. 6b) and that stable CD25 expression depends on PKCδ (Fig. 5g–i). Conversely, the cytoplasmic tail of CD25 is required for the stable expression of RACK1 (Fig. 6g, fig. S9f), suggesting that PKCδ:CD25:RACK1 may form a stable, ternary complex. For this reason, we constructed a three-dimensional (3D) structural model of the ternary PKCδ:CD25:RACK1 complex based on known features of the interaction between PKC isoforms and blades 3 and 6 of RACK1^75–76. We first modeled the catalytic domain of PKCδ with SWISS-model⁷⁷, using rat PKCδ in complex with its selective agonist ingenol-3-angelate⁶⁰ (PDB ID: 7KO6) as a template⁷⁸, and then the substrate CD25-tail bound to PKCδ based on our previously validated structural model of substrate-bound PKCs⁷⁹. The structural model of RACK1 was generated by homology modeling with RACK1 structure as the template (PDB ID: 4AOW)⁸⁰. The initial model of the PKCδ:CD25:RACK1 complex was energy-minimized with Rosetta⁸¹, which was followed by five runs of all-atom molecular dynamics (MD) simulations⁸² (see Materials and Methods section). MD simulations of CD25-ST and CD25-AV with ATP and Mg²⁺ revealed two possible states of the complex that were favored by CD25-ST and CD25-AV, respectively. The complex containing CD25-ST favored a tight conformation. In contrast, PKCδ:CD25:RACK1 containing CD25-AV transitioned to a less stable complex with RACK1 departing from the PKCδ:CD25-AV interface (Fig. 8d–e). CD25-AV retained some interactions with PKCδ; however, contacts between RACK1 and CD25-AV and PKCδ (van der Waals, hydrogen bond, and salt-bridge) were almost entirely disrupted (Fig. 8f–g). Calculation of conformational population densities of stable (assembled) and unstable (disassembled) states in MD simulations revealed that complexes with CD25-ST favored stable states, whereas complexes with CD25-AV favored less stable states (Fig. 8e). The interaction energies for RACK1:CD25, RACK1:PKCδ, and PKCδ:CD25 were calculated with the GROMACS energy module⁸³, and the sum of these three energies was used to evaluate the stability of the complex during MD simulation. The PKCδ:CD25:RACK1 complex containing CD25-ST explored one major conformation in which the complex was stable with an interaction energy average of −2174 kJ/mol, whereas CD25-AV formed a destabilized complex with an average interaction energy of −1360 kJ/mol (Fig. 8h), suggesting that the cytoplasmic tail of CD25 and PKCδ-mediated phosphorylation of Ser²⁶⁸/Thr²⁷¹ are required to form a stable complex. Together, these results support a structural model in which PKCδ-mediated phosphorylation of CD25 stabilizes a ternary complex with RACK1 for subsequent recruitment and activation of inhibitory phosphatases (INPP5D and PTPN6) to restrain signaling by oncogenic tyrosine kinases in leukemia cells.

CD25 surface expression as a therapeutic target to eradicate relapsed tyrosine kinase–driven leukemia

Previous studies linked CD25 expression in a subset of B-ALL¹⁰ and AML¹² cells to poor clinical outcomes. Here, we showed that CD25 marks leukemia subpopulations with active feedback control of oncogenic tyrosine kinase signaling. The presence of CD25⁺ subpopulations portends unfavorable outcomes for B-ALL and AML patients (fig. S1, S2). For this reason, we examined the therapeutic targeting of CD25 surface expression in two PDX models from patients with relapsed tyrosine kinase–driven B-ALL (BCR-ABL1, MXP5; CRLF2-rearranged and JAK2^R683S, NCL1). To target CD25 surface expression, we studied a pyrrolobenzodiazepine (PBD) dimer–containing antibody-drug conjugate (ADC; camidanlumab tesirine)⁸⁴. After sublethal irradiation (2 Gy) and 7 days after engraftment of BCR-ABL1 and JAK2-mutant B-ALL, NSG recipient mice were treated four times with weekly injections of 0.6 mg/kg CD25-ADC or B12-ADC control (human non-binding IgG1; fig. S8a–c). Mice treated with B12-ADC developed progressive leukemia and died within 25 to 65 days after engraftment. Mice in the CD25-ADC treatment group developed progressive disease until after the second injection (fig. S8a). After the second (day 21) and subsequent injections (days 35 and 49), all mice gradually achieved remission (fig. S8a), recovered, and survived for indefinite periods of time (fig. S8b–c). Because only a subpopulation of leukemia cells expresses CD25 on the surface at any given time (Fig. 1a–d, fig. S1g, S2g), the lasting effect of CD25-ADC treatment was unexpected. However, reflecting dynamic feedback control of oncogenic tyrosine kinase signaling, we found that CD25-negative B-ALL cells give rise to new CD25⁺ subpopulations within 14 days (Fig. 1g). With a half-life of 10.4 days in patients⁸⁵, it is conceivable that multiple injections over the course of 50 days could result in eradication of the entire leukemia population (fig. S8a–c), including cells that were initially CD25-negative and later transitioned to a CD25⁺ state (Fig. 1g).

In contrast, one single injection of CD25-ADC significantly prolonged overall survival but was not sufficient to eradicate relapsed BCR-ABL1 B-ALL, and all mice eventually died after 200 days (fig. S8d–e). Whereas this experiment suggests that multiple injections of CD25-ADC over a longer period of time would be necessary to eliminate leukemia cells that were initially CD25-negative, we tested whether increased CD25 surface expression enabled the eradication of leukemia with a single injection of CD25-ADC. Our previous experiments showed that the PKCδ agonist I3A can increase CD25 cell surface abundance by 20- to 30-fold (Fig. 5e, 5g). For this reason, we repeated the treatment of NSG mice engrafted with BCR-ABL1 B-ALL cells (MXP5) with only one injection of CD25-ADC or control B12-ADC on day 7 in combination with vehicle or 40 mg/kg of the PKCδ agonist I3A. Although I3A combined with control B12-ADC did not extend survival times, injection of I3A in combination with CD25-ADC significantly prolonged survival and achieved indefinite survival for about half of the treated mice (fig. S8d–e). We conclude that repeated injections with CD25-ADC achieved leukemia eradication and long-term survival in PDX models from patients who relapsed with tyrosine kinase–driven B-ALL, without significant weight loss and other signs of toxicity (fig. S9a–c). In addition, partial responses to subtherapeutic doses of CD25-ADC were enhanced by enforced CD25 surface expression by the PKCδ agonist I3A (fig. S8d–e).

DISCUSSION

Whereas CD25 has been extensively studied as IL-2R chain on T cells and NK cells, our findings reveal a previously unrecognized role for CD25 expressed in acute leukemia subpopulations, which portend poor clinical outcomes for patients. Combining genetic mouse and PDX models with interactome and phosphoproteomic studies, we identified CD25 and its phosphorylation by PKCδ as central elements of a previously uncharacterized feedback loop to stabilize fluctuations of oncogenic tyrosine kinase signaling in acute lymphoblastic and myeloid leukemia. To leverage the dependency of tyrosine kinase–driven acute leukemias on CD25-mediated feedback control, we assessed a CD25-ADC in preclinical models based on refractory leukemia PDX. These experiments highlighted the distinct dependency of tyrosine kinase–driven leukemias on robust feedback control and the unexpected function of PKCδ and CD25 in assembling its components. Targeting CD25 also results in the depletion of CD25⁺ T_reg cells, which may enhance antitumor immunity but also result in systemic autoimmunity⁸⁶. A phase 1 clinical trial showed both the safety and efficacy of CD25-ADC treatment in 34 AML and one B-ALL patients⁸⁷. However, whether outcomes correlated with the presence of oncogenic tyrosine kinases in the patients was not studied. Moreover, the trial was terminated during dose escalation, because a subsequently started clinical trial for patients with B and T cell lymphoma, in particular classical Hodgkin’s lymphoma had a clearer efficacy signal⁸⁸. In this context, our results support a rationale for future clinical studies to explore the benefit of CD25-ADC for patients with tyrosine kinase–driven refractory B-ALL and AML.

MATERIALS AND METHODS

Patient-derived B-ALL and AML samples

Deidentified patient samples (listed in table S1) were obtained from patients who gave informed consent and were in compliance with a protocol approved by the Yale Institutional Review Board (IRB) and IRBs at collaborating institutions (table S1). Patient-derived xenografts were cultured on OP9 stroma in Alpha Minimum Essential Medium (MEMα; Life Technologies) containing GlutaMAX, 20% fetal bovine serum (FBS), 1 mM sodium pyruvate, 100 IU/ml penicillin and 100 μg/ml streptomycin at 37°C in a humidified incubator with 5% CO₂. All human primary samples were tested negative for mycoplasma by detection kit (MycoAlert PLUS, LONZA).

Murine primary and leukemia cells

Bone marrow cells from 6- to 10-week old mice were harvested by flushing the cavities of femurs and tibia with chilled PBS, followed by filtering through a 40-μm strainer to yield a single-cell suspension. Spleen cells were directly extracted by forcing tissues through a 40-μm strainer into chilled PBS. Filtered cells were further incubated with lysis buffer (RBC Lysis Buffer, BioLegend) to lyse erythrocytes. After washing with chilled PBS, cells were subjected to further experiments. For IL-7–dependent pre-B cell culture, bone marrow cells were harvested and cultured in Iscove’s modified Dulbecco’s medium (IMDM; GIBCO) with GlutaMAX containing 20% FBS, 50 μM 2-mercaptoethanol, 100 IU/ml penicillin, 100 μg/ml streptomycin in the presence of 10 ng/ml recombinant mouse IL-7 (Peprotech). For the BCR–ABL1 leukemia model, pre-B cells were retrovirally transformed by BCR-ABL1 and then IL-7 was withdrawn to select the transduced cells. For myeloid progenitor cells, murine bone marrow cells were cultured in IMDM containing 10 ng/ml recombinant mouse IL-3, 25 ng/ml recombinant mouse IL-6, and 50 ng/ml recombinant mouse stem cell factor (SCF) (PeproTech). Myeloid progenitor cells were retrovirally transformed by FLT3^ITD and then cytokines were withdrawn to select the transduced cells.

Animals

C57BL/6, NOD.Cg-Prkdc^scid Il2rg^tm1Wjl/SzJ (NSG), B6.C(Cg)-Cd79a^tm1(cre)Reth/EhobJ (Mb1-Cre), and B6.C- Cd79a^{tm3(Cre/ERT2)Reth}/EhobJ (Mb1-CreER^T2) mice were purchased from Jackson Laboratory. Cd25^fl/fl mice were obtained from Dr. Fontenot and Dr. Rudensky. Cd25^fl/fl mice were crossed with Mb1-Cre, Mb1-Cre-ER^T2 to generate a B lineage–specific knockout model. B6;129X1-Prkcd^tm1Msg/J (Prkcd^−/−) mice were obtained from Dr. Tarakhovsky. Tg(Nr4a1-EGFP)GY139Gsat/Mmcd (Nr4a1-GFP) mice were obtained from Dr. Hogquist. All animal procedures and protocols were approved by the Institutional Animal Care and Use Committee of Yale University (IACUC). We have complied with all relevant ethical regulations. All mouse models used in the study are listed in table S3. Mice were genotyped by Transnetyx (Cordova, TN). The sequences of the genotyping primers used in this study are listed in table S4.

Retroviral and lentiviral transduction

Constructs used for retroviral or lentiviral transduction are listed in table S5. Retroviral supernatants were generated by co-transfection of HEK 293FT cells with retroviral constructs together with pHIT60 (gag-pol) and pHIT123 (for mouse) or pAMPHO (for human ecotropic envelope) using Lipofectamine 2000 (Invitrogen) and culture in high glucose Dulbecco’s modified Eagle’s medium (DMEM, GIBCO) with GlutaMAX containing 10% FBS, 100 IU/ml penicillin, 100 μg/ml streptomycin, 25 mmol l⁻¹ HEPES, 1 mmol l⁻¹ sodium pyruvate and 0.1 mmol l⁻¹ non-essential amino acids for 16 hours. After induction with 10 mM sodium butyrate for 8 hours, the virus-containing supernatant was collected and filtered through a 0.45-μm filter. The virus-containing supernatants were loaded by centrifugation at 2000g for 90 min at 32°C on non-tissue culture 6-well plates coated with 50 μg/ml Retronectin (Takara). For viral transduction, 3 to 5 × 10⁶ cells were transduced per well by centrifugation at 600g for 30 min in the appropriate culture medium and maintained at 37°C at 5% CO₂ for 48 hours. Lentiviral supernatants were generated by co-transfection of HEK 293FT cells with lentiviral constructs together with pCD/NL-BH and VSVG with Lipofectamine 2000 (Invitrogen). The lentiviral supernatant was replaced with fresh medium on the next day.

Gene deletion by non-viral genome editing

Ribonucleoprotein (RNP) formulation, preparation of single-strand DNA templates, and electroporation were performed as previously described⁸⁸. Briefly, chemically synthesized crRNAs (160 μM) and tracrRNAs (160 μM) reconstituted in Duplex Buffer (IDT technologies) were mixed 1:1 by volume and annealed by incubation at 37°C for 30 min. The ssODNenh (electroporation enhancer) was synthesized from IDT and resuspended to 100 μM in water and mixed 1:1 v/v with preformed gRNAs. Recombinant Cas9-NLS (40 μM) was then mixed 1:1 by volume with gRNA to produce RNA RNP complexes. RNP complexes were freshly prepared before electroporation. Electroporation was performed by using pulse code EH-115 on a Lonza 4D 96-well electroporation system. Predesigned Alt-R CRISPR-Cas9 guide RNAs were purchased from IDT and are listed in table S6. Non-targeting control guide RNAs were purchased from IDT.

Quantitative RT-PCR

Quantitative real-time PCR was performed with SYBRGreen mix from Invitrogen according to standard PCR conditions with a ViiA 7 Real-Time PCR System (Applied Biosystems) and was analyzed by QuantStudio Real-Time PCR software (Thermo Fisher Scientific).

Western blotting

Cells were washed with chilled PBS and then lysed in CelLytic buffer (Sigma-Aldrich) supplemented with 1% protease inhibitor cocktail (Roche Diagnostics), 1% phosphatase inhibitor cocktail (EMD Millipore) and 1 mM PMSF on ice. A total of 10 μg of cell lysates were separated on mini precast gels (Bio-Rad), which was followed by transfer to nitrocellulose membranes (Bio-Rad). Nitrocellulose membranes incubated with the primary antibodies were blotted with alkaline phosphatase–conjugated secondary antibodies (Invitrogen) and chemiluminescent substrate (Invitrogen). Images were collected with the ChemiDoc MP Imaging System (BioRad). Antibodies used in this study (table S7) were diluted 1:750 to 1:1000 in blocking buffer for use.

Flow cytometry

PBS-washed cells were blocked with TruStain FcX (Biolegend) for 10 min on ice and then stained with the appropriate antibodies (listed in table S7) or isotype control for 25 min on ice. Cells were then washed and resuspended in chilled PBS containing 0.75 μg/ml DAPI to exclude dead cells. Acquisition was performed with a LSRFortessa flow cytometer (BD Biosciences). Fluorescence-based cell sorting was performed with a FACSAria II (BD Biosciences). FACS data were analyzed with FlowJo software (FlowJo, LLC). For apoptosis analyses, annexin V and 7-AAD (BD Biosciences) were used. For cell cycle analysis, the BrdU flow cytometry kit (BD Biosciences) was used according to the manufacturer’s instructions. For intracellular staining of cytoplasmic proteins, cells were first stained for cell surface antigens and subsequently fixed in BD Cytofix/Cytoperm buffer containing the fixative paraformaldehyde and the detergent saponin. Cells were then washed and resuspended in BD Perm/Wash buffer and stained with the appropriate antibodies. For statistical quantification, data were plotted with GraphPad Prism 10 or SigmaPlot. FACS antibodies were added to 0.5 to 1 × 10⁶ cells/50 μl in PBS for a final dilution of 1:50 for human cells or 1:200 for mouse cells.

Kinase assays

To predict putative kinases targeting CD25, GPS 5.0 (Group-based Prediction System: http://gps.biocuckoo.cn/) was used against the cytoplasmic tail of CD25 (WQRRQRKSRRTI)⁸⁹. In vitro profiling of the 62 kinases identified by GPS 5.0 was performed at Reaction Biology Corporation (www.reactionbiology.com; Malvern, PA) using the “HotSpot” assay platform as previously described⁹⁰. Briefly, specific kinase and substrate pairs together with the required cofactors were prepared in reaction buffer: 20 mM Hepes (pH 7.5), 10 mM MgCl₂, 1 mM EGTA, 0.02% Brij35, 0.02 mg/ml BSA, 0.1 mM Na₃VO₄, 2 mM DTT, 1% DMSO). Compounds were added to the reaction, followed by the addition of ³³P ATP to a final concentration of 10 μM. After the reaction mixes were spotted onto P81 ion exchange filter paper, unbound phosphate was removed by extensive washing of the filters in 0.75% phosphoric acid. After subtraction of the background derived from control reactions containing inactive enzyme, kinase activity was quantitated as the percentage remaining ³³P ATP in the test samples.

Proximity ligation assays

For PLAs, cells were treated with 5 nM I3A or vehicle control for 10 min at 37°C, 5% CO₂. Cells were immediately washed with chilled PBS and subsequently fixed in fixation buffer (Biolegend) containing 4% paraformaldehyde (PFA) for 25 min on ice and then washed with chilled PBS. For cellular membrane staining, cells were labelled for 5 min at room temperature with 5 μg/ml WGA conjugated to Alexa Fluor 488 (Thermo Fisher Scientific). Cells were immediately washed and permeabilized in Perm/Wash Buffer (BD Biosciences), followed by blocking in Duolink Blocking buffer for 30 min at room temperature. Cells were incubated with the appropriate primary antibodies (table S7) at a 1:150 dilution overnight at 4°C. Cells were washed and settled on Cell-Tak–coated Shandon Single Cytoslides (Corning) by cytospin at 400g for 5 min. PLA reactions were performed according to the manufacturer’s protocol (Duolink, Sigma). Briefly, primary antibodies were incubated with Duolink in situ PLA probe Plus or Minus (Sigma-Aldrich) and then visualized with Duolink Detection Reagent Red (Sigma-Aldrich). Cells were mounted with Duolink in situ Mounting Medium with DAPI (Sigma-Aldrich). Microscopic images were acquired with an Olympus IX3–55 and analyzed with CellSens imaging software (Olympus) and ImageJ. For quantification of PLA signals, one dot was defined as a pixel size of 5 × 5 by BlobFinder software. Statistical significance was analyzed with the unpaired Student’s t-test and plotted with GraphPad Prism 10.

RNA-seq analysis

Normal IL-7–dependent pre-B cells and BCR-ABL1-transduced pre-B cells from Cd25^fl/fl mice were transduced with 4-OHT–inducible Cre-ER^T2 or ER^T2 and treated with 4-OHT or vehicle for 48 hours. Total mRNA was extracted with a RNeasy Kit (Qiagen) according to the manufacturer’s instructions. Sequencing was performed on an Illumina Hiseq 2500 (Illumina) as previously described⁹¹. Transcripts were quantified against the Gencode GRCm38 reference transcriptome vM25 using either Salmon v1.4.0⁹² for differential expression analysis or STAR v2.7.6a⁹³ for alignment and visualization. Differential gene expression was analyzed in R with the DESeq2 package v1.36.0, and shrunken log₂ fold change values were used for GSEA using fgsea v1.22.0. Gene sets were obtained from MSigDB or internal data as indicated.

Bio-ID proteomics and data analysis

Bio-ID proteomics was performed as previously described⁹¹. Briefly, CD25-deficient patient-derived PDX2 B-ALL cells with CRISPR/Cas9-mediated gene deletion were reconstituted with wild-type CD25 (ST) or PKCδ-motif mutant CD25 (AV) carrying S268A and T271V with C-terminal BirA (engineered biotin ligase) and selected by culture in 1 μg/ml puromycin for 3 days. A BirA-expressing construct was used as a negative control. To induce the biotinylation of proteins proximal to CD25, the cells were treated with 50 μmol/l biotin for 10 min in the presence or absence of I3A. Cells were washed three times with chilled PBS and lysed in IP/WASH buffer (Pierce) in the presence of 1 × HALT protease inhibitor (Thermo Fisher). The lysates were incubated with Streptavidin C1 MyOne Dynabeads (Invitrogen) for 16 hours at 4°C. Unbound proteins were washed three times with 2% SDS-PBS, three times with PBS, and three times with pure water. The eluted proteins were gel-purified, subjected to in-gel digestion, and then analyzed by mass spectrometry. For LC-MS/MS analysis, peptides were analyzed with a Dionex UltiMate 3000 Rapid Separation LC (RSLC) systems and an Orbitrap mass spectrometer (ThermoFisher Scientific). Peptide samples (6 μl) were loaded onto the trap column, which was 150 μm × 3 cm in-house packed with 3-μm C18 beads. The analytical column was a 75 μm × 10.5 cm PicoChip column packed with 3-μm C18 beads (New Objectives). The flow rate was kept at 300 nl/min. Solvent A was 0.1% formic acid (FA) in water and Solvent B was 0.1% FA in ACN. The peptide was separated on a 120-min analytical gradient from 5% acetonitrile (ACN)/0.1% FA to 40% ACN/0.1% FA. The mass spectrometer was operated in data-dependent mode. The source voltage was 2.40 kV and the capillary temperature was 275⁰C. MS1 scans were acquired from 400 to 2000m/z at 60,000 resolving power and the automatic gain control (AGC) was set to 1 × 10⁶. The fifteen most abundant precursor ions in each MS1 scan were selected for fragmentation. Precursors were selected with an isolation width of 1 Da and fragmented by collision-induced dissociation (CID) at 35% normalized collision energy in the ion trap, whereas previously selected ions were dynamically excluded from re-selection for 60 s. The MS2 AGC was set to 3 × 10⁵. For data analysis, proteins were identified from the MS raw files with the Mascot search engine (Matrix science). MS/MS spectra were searched against the SwissProt human database. All searches included carbamidomethyl cysteine as a fixed modification and oxidized methionine, deamidated asparagine and glutamine, and acetylated N terminus as variable modifications. Three missed tryptic cleavages were allowed. The MS1 precursor mass tolerance was set to 10 ppm and the MS2 tolerance was set to 0.6 Da. A 1% false discovery rate (FDR) cutoff was applied at the peptide level. Only proteins with a minimum of two peptides above the cutoff were considered for further study. For any peptide not detected in all samples of a given condition, background peptide abundances were imputed from a Gaussian distribution centered around the minimal observed abundance using the MinProb method; otherwise, missing at random (MAR) abundances⁹⁴ were imputed with the MLE method using the MSnbase package in R. Differential abundance testing was performed using linear modeling with empirical Bayes statistics with the DEP and limma packages.

Phosphoproteomic analysis

Cd25^fl/fl Mb1-Cre-ER^T2 BCR–ABL1 pre-B ALL cells were treated with 4-OHT for 48 hours and subjected to proteomics analysis as previously described⁹¹. For global phosphoproteomics, cellular extracts were prepared in urea lysis buffer, sonicated, centrifuged, reduced with DTT, and alkylated with iodoacetamide. Total protein (15 mg) for each sample was digested with trypsin, and 500 mg of total protein for each sample was digested with Lys/trypsin for IMAC analysis. Samples were purified over C18 columns and dried in a lyophilizer. Dried samples were resuspended and enriched with Fe-IMAC beads and purified over C18 STAGE tips (Rappsilber). For pY phosphoproteomics, dried samples were resuspended and enriched with the Phosphotyrosine pY-1000 Motif Antibody (CST #8803) and were purified over C18 STAGE tips (Rappsilber). Replicate injections of each sample were run non-sequentially on the instrument. Peptides were eluted with a 150-min (IMAC) linear gradient of acetonitrile in 0.125% formic acid delivered at 280 nl/min. Tandem mass spectra were collected in a data-dependent manner with a Thermo Orbitrap Fusion Lumos Tribrid mass spectrometer using a top-twenty MS/MS method, a dynamic repeat count of one, and a repeat duration of 30 s. Real-time recalibration of mass error was performed using lock mass (Olsen) with a singly charged polysiloxane ion m/z = 371.101237. MS/MS spectra were evaluated with SEQUEST and the Core platform from Harvard University (Eng, Huttlin, Villen). Files were searched against the SwissProt Homo sapiens FASTA database. A mass accuracy of ± 5 ppm was used for precursor ions and 0.02 Da for the product ions. Enzyme specificity was limited to trypsin, with at least one tryptic (lysine- or arginine-containing) terminus required per peptide and up to four mis-cleavages allowed. Cysteine carboxamidomethylation was specified as a static modification, and oxidation of methionine and phosphorylation on serine, threonine, and tyrosine residues were allowed as variable modifications. Reverse decoy databases were included for all searches to estimate FDRs and filtered with a 1% FDR in the Linear Discriminant module of Core. Peptides were also manually filtered with a ± 5 ppm mass error range and the presence of a phosphorylated residue. All quantitative results were generated with Skyline (MacLean) to extract the integrated peak area of the corresponding peptide assignments. The accuracy of the quantitative data was ensured by manual review in Skyline or in the ion chromatogram files.

Nfatc2 immunofluorescence and quantification

Cd25^fl/fl BCR–ABL1 pre-B ALL cells carrying 4-OHT-inducible Cre-ER^T2 or ER^T2 were treated with 4-OHT for 2 days. The cells were washed and settled on Superfrost Plus Microscope Slides (Fisherbrand) by cytospin at 450 rpm for 5 min. Cells were then fixed with 4% PFA (CytoFIx, BD Biosciences) for 20 min at room temperature and then washed three times with PBS. Cells were then permeabilized with methanol (Perm Buffer III, BD Biosciences), washed three times with PBS, and blocked with Blocking buffer (Sigma) for 30 min at room temperature. Slides were incubated overnight at 4°C with Nfatc2 antibody (CST, 1:150). The following day, the cells were washed three times with PBS and incubated for 1 hour at room temperature with Alexa Fluor 647–conjugated anti-rabbit IgG as secondary antibody (Invitrogen). Slides were washed three times with PBS, dried, and mounted in Prolong Gold anti-fade mounting media with DAPI (ThermoFisher). Images were acquired with a Cytation 5 Cell Imaging Reader (Biotek) and analyzed with Gen5 version 3.11 imaging software (Biotek). Quantitative image analysis was conducted with QuPath 0.2.2.

Single-cell Ca²⁺ measurements

Ultrasensitive protein calcium sensor (GCaMP6s) was subcloned from pGP-CMV-GCaMP6s (Addgene #40753) into the MSCV-IRES-Puro vector. Cells transduced to express GCaMP6s were selected in the presence of puromycin, then plated at 1 × 10⁶ cells/ml on RetroNectin-coated non-tissue plates. Fluorescent images were captured with a 300-ms exposure time and 2-s interval time with Q-Capture pro 7 (Version 7.0.5, Q-imaging) software connected with an Olympus IX71 microscope and a QImaging Exi Aqua camera. Fluorescence images were captured at the exposure times and intervals indicated in the figures with ZEN software (Version 2.3, Zeiss) connected with a Zeiss Axio Observer 7 microscope and an Axiocam 702 mono camera. For image quantification, ImageJ with the Time Series Analyzer V3 plugin (http://imagej.nih.gov/ij/plugins/time-series.html) was used to calculate the background-subtracted GCaMP6s intensity of single cells in each frame. Curve fitting of the fold-changes in GCaMP6s intensity were plotted with the fast Fourier transform built into Matlab (Uhlén, 2004, Sci. STKE). The single-cell Ca²⁺ amounts were visualized by a heat-map with JMP version 9.0 (SAS Institute).

Colony-forming assay

Ten thousand mouse BCR-ABL1 ALL cells or patient-derived B-ALL cells were resuspended in MethoCult medium (M3231 for mouse cells and H4230 for human cells, StemCell Technologies) and plated on 3-cm diameter dishes with an extra dish filled with water to prevent evaporation. After 7 to 14 days, colony numbers were counted with a GelCount analyzer (Oxford Optronix). Ten thousand AML cells were resuspended in MethoCult medium (M3231). After 21 days, colonies were counted with a GelCount analyzer.

Co-immunoprecipitation and mass spectrometry

Co-immunoprecipitation was performed with the Pierce Crosslink Magnetic IP/Co-IP kit (Thermo Scientific) according to the manufacturer’s instructions. Briefly, patient-derived Ph⁺ B-ALL cells were transduced with MSCV-CD25-ST-IRES-Puro or MSCV-CD25-AV-IRES-Puro. Transduced cells were selected with 1 μg/ml puromycin, harvested, and washed with PBS before undergoing lysis with IP lysis/Wash buffer. Each 5 μg of anti-CD25 (clone, M-A251; Biolegend) per sample was coupled to protein A/G magnetic beads and covalently cross-linked with 20 μM disuccinimidyl suberate (DSS). The antibody-crosslinked beads were incubated with cell lysate, washed to remove non-bound material, and eluted in a low-pH elution buffer that dissociates bound antigen from the antibody-crosslinked beads. The enriched antigen in low pH was immediately neutralized and subjected to Western blotting analysis.

In vivo leukemia initiation assay

For in vivo leukemia initiation assays, the numbers of Cd25^fl/fl BCR–ABL1 pre-B ALL cells indicated in the figure legends were injected into sublethally irradiated (200 cGy) 8- to 10-week old female NSG mice (Jackson Laboratories, ME) through the tail vain. Mice were euthanized when they showed signs of leukemia burden, such as a hunched back, weight loss, or an inability to move, and then the bone marrow, spleen, or both were collected to assess leukemia infiltration by flow cytometry. Kaplan-Meier survival analysis was performed with GraphPad Prism 10 (GraphPad Software Inc.) to compare overall survival (OS) rates. The Mantel-Cox log-rank test was used for statistical analysis with GraphPad Prism 10.

Assessment of CD25-ADC efficacy

For in vivo experiments, patient-derived B-ALL cells were transduced with retrovirus expressing firefly luciferase and then selected by blasticidin. Cells (numbers indicated in the figure legends) were washed and then injected through the tail vein into sublethally irradiated (200 cGy) 8- to 10-week old female NSG recipient mice.. The in vivo expansion and leukemic burden were monitored by luciferase bioimaging with an IVIS 100 bioluminescence/optical imaging system (Xenogen) at the times indicated in the figure legends. Briefly, D-luciferin (Promega) dissolved in PBS was injected intraperitoneally at a dose of 2.5 mg per mouse 15 min before luminescence was measured. All mice were initially anesthetized with 5% isoflurane and then were maintained during the detection of light emission with 2% isoflurane introduced through a nose cone. CD25-ADC and the control ADC dissolved in PBS were administered through the tail vein at dose of 600 μg/kg body weight at the times indicated in the figure legends. For I3A administration, PEG dissolved in PBS as a 20% (v/v) working solution was used as a vehicle for either I3A at a dose of 40 μg/kg body weight or DMSO. A total volume of 600 μl of PEG/PBS-I3A or PEG/PBS-DMSO control was administered through intraperitoneal (i.p.) injection. Mice were euthanized when they showed signs of leukemic burden, such as a hunched back, weight loss, or an inability to move. Kaplan-Meier survival analysis was performed using GraphPad Prism 10 to compare OS rates. The Mantel-Cox log-rank test was used for statistical analysis with GraphPad Prism 10.

Modeling of the RACK1-PKCδ–CD25 tail complex

The initial model of the RACK1-PKCδ–CD25 tail complex was constructed based on multiple pieces of evidence. McCahill et al.⁹⁵ proposed an interaction model between PKCβ and blades 3 and 6 of RACK1 based on previous work^96–98. They highlighted the sequence similarity between the Ca²⁺-binding site on the C2 domain of PKCβ and blade 6 of RACK1⁹⁵, which suggested that RACK1 interacts with the catalytic domain (CD) of PKCβ similarly to the way that the PKCβ C2 domain interacts with the CD. We assumed that PKCδ and PKCβ might interact with RACK1 using the same motif, because PKCδ also has a C2 domain. Using the model of the interaction between the C2 and CD domains of PKCβ proposed by Jones et al.⁹⁹, we proposed a model for the interaction between the C2 and CD domains of PKCδ when pseudo substrate (PS) was present. We combined the RACK1-PKCβ model from McCahil et al. and the PKCδ C2-CD-PS model based from Jones et al. to construct the initial model of the RACK1-PKCδ–CD25 tail complex. Because there is no available crystal structure for full length PKCδ, we first modeled the CD of PKCδ with the SWISS model¹⁰⁰, using the structure of PKCθ¹⁰¹ (PDB ID: 5F9E) as a template. We then modeled CD25 tail–bound PKCδ based on our previously validated structural model of substrate-bound PKCa¹⁰². The amino acid sequence of the CD25 tail used in this study is: WQRRQRKSRRTI. Next, we constructed the PKCδ CD–CD25 tail–RACK1 complex by overlaying the PKCδ CD–CD25-C2 complex based on the evidence from Jones et al.⁹⁹, and then replacing the PKCδ C2 domain with RACK1 by aligning RACK1 blade 6 with the Ca²⁺-binding motif of the C2 domain. The structure of RACK1 was acquired by homology modeling using the WT RACK1 sequence based on the RACK1 structure (PDB ID: 4AOW)¹⁰³. All homology modeling steps were performed with SWISS-model¹⁰⁰ and with the protein sequence acquired from UniProt. The initial model of the RACK1-PKCδ–CD25 tail complex was energy-minimized with Rosetta¹⁰⁴, followed by 5 runs of all-atom MD simulations (100-ns × 5) totaling to 500 ns to stabilize a solution structure of the complex. We clustered the conformations in the trajectories based on the root mean square deviations (RMSD) cutoff of 1.5 Å, and the centroid of the top occupied conformation cluster was chosen as the most stable conformation for further simulations. Two sets of simulations were performed based on the most stable confirmation: (i) The PKCδ–CD25 tail–RACK1 complex consisting of WT CD25 with ATP and Mg²⁺, and (ii) the complex with the S268A mutation in CD25 with ATP and Mg²⁺. We performed 5 simulations (each for 200 ns) for a total of 1000 ns for each system. The trajectories from 100 ns to 200 ns for each velocity run were combined into a 500-ns trajectory (100 ns × 5) and used for all of the analysis.

MD simulation details

All simulation systems were generated using CHARMM-GUI¹⁰⁵ with a CHARMM36m force field¹⁰⁶. The force field parameter for ATP was taken from the CHARMM36 General Force Field. The PKCδ CD–CD25 tail–RACK1 complex with ATP and Mg²⁺ was immersed in a 104-Å³ cubic TIP3 water box and then the solvated system was neutralized with 0.15 M KCl. The S268A mutation was performed with CHARMM-GUI and prepared using the same procedure as that for the complex containing WT CD25. We performed all simulations with the GROMACS 2016 package¹⁰⁷. The LINCS algorithm was applied on all bonds and angles of water molecules with a 2-fs time step used for integration. A cutoff of 12 Å was applied for non-bond interactions, and the particle mesh Ewald method¹⁰⁸ was used to treat long-range van der Waals interactions. Each system generated from the CHARMM-GUI first went through energy minimization, with the steepest descent method until the maximum force was less than 1000 kJ/mol/nm. The system was then slowly heated from 0 to 310 K in an NVT ensemble during a 1-ns heating process. The temperature was maintained with the Nosé-Hoover thermostat¹⁰⁹. A harmonic position restraint of 5 kcal/mol/Å² was applied on all heavy atoms during the heating process. We further equilibrated the system in the NPT ensemble for 30 ns to gradually release the restraint from 5 kcal/mol-Å² to 0 kcal/mol/Å² with a 1 kcal/mol/Å² per 5-ns window. Constant pressure was maintained with the Parrinello-Rahman method¹¹⁰ when coupled to a 1-bar bath. The last frame from the equilibrium process was used as the initial conformation for production simulations. We used five random seeds to generate different initial velocities for five production runs lasting for either 100 or 200 ns.

Population density of two conformations

MD simulations of the CD25-ST and CD25-AV mutant revealed two possible states of the RACK1-PKCδ–CD25 tail complex. The complex was stable in the majority of CD25-ST and some of CD25-AV mutant simulation. However, two of the five MD simulation runs of CD25-AV transitioned to a less stable complex, featuring RACK1 departing from the PKCδ–CD25 tail complex. To evaluate the population of the less-stable conformations, the RMSD of the RACK1-PKCδ–CD25 tail complex was calculated by selecting all of the Cα atoms for the CD25-ST and CD25-AV trajectories. A cutoff of 6 Å was chosen to separate a stable complex from the less stable complex conformation. The population densities for CD25-ST and CD25-AV sampling in both these conformational states were calculated.

Complex interaction energy analysis

The interaction energy of the complex was calculated as the sum of the short-range columbic interaction energy and the Lennard-Jones potential interaction energy with the GROMACS energy module. The interaction energies between RACK1 and the CD25 tail, RACK1 and PKCδ, and PKCδ and the CD25 tail were calculated, and the sum of these three energies was used to evaluate the stability of the complex during MD simulations. The results showed that the RACK1-PKCδ–CD25-ST tail complex explored one major conformation in which the complex was stable with an interaction energy average at −2174 kJ/mol. However, the RACK1-PKCδ–CD25-AV tail complex showed two different conformations. One conformation occurred when RACK1-PKCδ–CD25 tail formed a stable complex with an average interaction energy of −2245 kJ/mol. The other conformation with destabilized RACK1 showed an average interaction energy of −1360 kJ/mol. The interaction energy analysis indicated that the complex containing WT CD25 is far more stable than the complex containing the CD25-AV mutant.

BCR-ABL1 kinase reporter

The BCR-ABL1 kinase reporter pFX-Pickles 2.34NES was obtained from Addgene (#100028) and subcloned into the MSCV-IRES-Puro vector. The packaged retrovirus was used to transduce both BCR-ABL1–transformed pre-B ALL cells and normal pre-B cells, and normal pre-B cells served as a negative control. After transduction, Pickles 2.34NES⁺ cells were FACS sorted and expanded in IMDM with GlutaMAX containing 20% FBS, 50 μM 2-mercaptoethanol, 100 IU/ml penicillin, 100 μg/ml streptomycin with or without 10 ng/ml murine IL-7. Cells were plated onto a Cell-Tak–coated 24-well glass-bottom plate (Eppendorf) at a density of 1 × 10⁶ cells/ml with 500 μl of FluoroBrite DMEM (Thermo Fisher Scientific) supplemented with GlutaMAX, 10% FBS, 1 mM sodium pyruvate, 25 mM HEPES, and 0.1 mM non-essential amino acids. For fluorescence imaging, images were captured with ZEN Black software version 2.3 (Carl Zeiss), which controlled a Zeiss LSM 880 microscope equipped with an AiryScan detector. To quantify BCR-ABL1 activity, FRET to ECFP ratio images were generated with the Image Calculator function in ZEN Blue software version 2.3 (Carl Zeiss). The average values of the ratio in defined cell regions were calculated with Fiji ImageJ version 1.52e. The quantification results were visualized with GraphPad Prism version 8.0.0 (GraphPad Software, Inc.).

Quantification and statistical analysis

Data are shown as means ± SD, unless stated otherwise in the figure legends. Statistical analysis was performed with GraphPad Prism 10 using an unpaired two-tailed t test or a log-rank test, as indicated in the figure legends. Statistical significance was considered at P < 0.05. For in vivo transplantation experiments, the minimal number of mice in each group was calculated through use of the ‘cpower’ function in the R/Hmisc package. Kaplan-Meier survival analysis was used to estimate OS with GraphPad Prism 10. A Mantel-Cox log-rank test was used to compare the differences between two groups. No animals were excluded. For patient OS analysis, patients in each dataset were divided into two groups based on whether their expression was above or below the median of CD25 abundance, and Kaplan–Meier survival analysis was used to estimate OS. Patient-outcome data for B-ALL were obtained from the National Cancer Institute TARGET DATA Matrix of the Children’s Oncology Group (COG) Clinical Trial P9906 (GSE11877)^111,112, the Eastern Cooperative Oncology Group (ECOG) Clinical Trial E2993 (GSE5314)¹¹³, and Children’s Oncology Group study AALL0232 (GSE68790)¹¹⁴. Patient-outcome data for AML were obtained from the genomics study of acute myeloid leukemia program at Washington University (GSE10358)^115,116, gene expression profiling study of 524 cases of de novo AML (GSE14468)^117–119, and TCGA LAML (https://portal.gdc.cancer.gov/projects/TCGA-LAML). ChIP-seq data of the NF-κB subunits RelA, RelB, cRel, and p50 at the CD25 locus in human lymphoblastoid B cells were obtained from GSE55105¹²⁰. A log-rank test was used to compare survival differences between patient groups. The R package ‘survival’ Version 2.35–8 was used for the survival analysis, and the Cox proportional hazards regression model in R was used for the multivariate analysis (https://www.r-project.org/). The investigators were not blinded to allocation during experiments and outcome assessment. Experiments were repeated to ensure the reproducibility of the observations.

Supplementary Material

NIHMS2147568-supplement-Supplementary_Material.pdf^{(8.8MB, pdf)}

Figs. S1 to S9.

Tables S1 to S7.

Acknowledgments:

We thank L. Klemm and current and former members of the Müschen laboratory for their support and helpful discussions, J. Fontenot (Biogen) for providing CD25^fl/fl mice, M. Carroll (University of Pennsylvania) for the PAX5-JAK2 and EBF1-PDGFRB constructs, A. Marson, V. Vykunta and B. Shy (University of California San Francisco) for sharing flow cytometry data from CD25-deficient patients, F. Zammarchi and P. H. van Berkel (ADCT) for helpful discussions of the CD25-ADC treatment studies.

Funding:

Research in the Müschen laboratory is funded by the NIH through an NCI Outstanding Investigator Award R35CA197628, R01CA282877, R01CA213138, R01AI164692, R01AI192914 and P01CA233412 (to M.M.), R01CA271497 (to J.C.), the Howard Hughes Medical Institute HHMI-55108547 (to M.M.), the Arthur H. and Isabel Bunker Chair in Hematology (to M.M.), Blood Cancer United through Translational Research Program Grant 6709-26 and SCOR grant 7026-21, the V Foundation for Cancer Research through Translational Research Grant T2018-003B and All Stars Award AST2025-015 (to M.M.). M.M. is a Howard Hughes Medical Institute (HHMI) Faculty Scholar. C.H. is supported by the Rally Foundation for Childhood Cancer, The Leukemia Research Foundation, the V Foundation, NIH/NCI K22 CA251649, and Alex’s Lemonade Stand Foundation. J.L. was supported by the National Research Foundation of Korea (NRF) grant (RS-2024-00339966) of the government of the Republic of Korea and by Samsung Research Funding & Incubation Center of Samsung Electronics under Project Number SRFC-MA2402-11.

Footnotes

Competing interests: M.M. has received research support from ADC Therapeutics. The other authors declare that they have no competing interests.

Data and materials availability:

No original code or materials were generated or used in this study. RNA-seq data for Cd25^fl/fl pre-B and BCR-ABL1 cells are available at GSE210872. Gene expression profiling in CD25⁺ versus CD25⁻ patient-derived Ph⁺ ALL is available at GSE115819. Proteomics data have been deposited to the ProteomeXchange Consortium through the PRIDE partner repository with the following accession numbers: tyrosine-phosphoproteome, PXD036128; global-phosphoproteome, PXD038996; CD25 interactomes, PXD035989. All other data needed to evaluate the conclusions in the paper are included in the paper or the Supplementary Materials.

References and Notes

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Data Availability Statement

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PERMALINK

Dynamic feedback control of oncogenic tyrosine kinase signaling in acute leukemia

Jaewoong Lee

Ruifeng Sun

Kohei Kume

Mark E Robinson

Zhangliang Cheng

Kadriye Nehir Cosgun

Ning Ma

Christian Hurtz

Huimin Geng

Selina M Luger

Mark R Litzow

Elisabeth Paietta

Jianjun Chen

Nagarajan Vaidehi

Markus Müschen

Abstract

INTRODUCTION

RESULTS

CD25 expression in B-ALL and AML subpopulations is linked to oncogenic tyrosine kinase signaling

Fig. 1. CD25 expression in acute leukemia subpopulations marks a quiescent cell state.

Leukemia subpopulations with active tyrosine kinase signaling are marked by CD25 expression

CD25 expression reflects negative feedback control of oncogenic tyrosine kinase signaling

CD25+ subpopulations are small, quiescent, and drug-resistant leukemia cells

Absence of a clonal hierarchy between CD25+ and CD25− leukemia cells

CD25 expression reflects NF-κB activation in leukemia subpopulations

B cell precursors express CD25 in response to oncogenic tyrosine kinase signaling but not IL-2 signaling

CD25 is essential for competitive fitness and the initiation of tyrosine kinase–driven leukemia

Fig. 2. CD25 is essential for tyrosine kinase–driven acute leukemia.

CD25 functions as negative feedback regulator of NF-κB signaling

Fig. 3. CD25 functions as a feedback regulator of NFAT and NF-κB signaling.

CD25 inhibits Ca2+ oscillations and NFAT signaling in tyrosine kinase–driven B-ALL

CD25 is required for the activation of inhibitory phosphatases to curb oncogenic tyrosine kinase signaling

Fig. 4. CD25 inhibits oncogenic tyrosine kinase signaling in acute leukemia.

CD25 interacts with regulators of tyrosine kinase signaling in BCR-ABL1 B-ALL

A motif in the cytoplasmic tail of PKCδ is critical for stable CD25 surface expression

Fig. 5. PKCδ-dependent phosphorylation of the cytoplasmic tail of CD25 is required for its cell surface expression.

PKCδ-mediated stabilization of CD25 is required for leukemogenesis

Structural basis of the CD25-dependent recruitment and activation of inhibitory phosphatases

Fig. 6. CD25-dependent recruitment and activation of the inhibitory phosphatases PTPN6 and INPP5D.

The cytoplasmic tail of CD25 is required for the recruitment and activation of inhibitory phosphatases

Fig. 7. Rescue of leukemia cell survival and colony formation by reconstitution with CD25 depends on an intact cytoplasmic tail motif.

Activation of the PKCδ scaffold protein RACK1 can rescue the loss of CD25 function

Fig. 8. Structural basis of the CD25-dependent formation of PKCδ-containing scaffolds.

Structural modeling of the PKCδ:CD25:RACK1 ternary complex

CD25 surface expression as a therapeutic target to eradicate relapsed tyrosine kinase–driven leukemia

DISCUSSION

MATERIALS AND METHODS

Patient-derived B-ALL and AML samples

Murine primary and leukemia cells

Animals

Retroviral and lentiviral transduction

Gene deletion by non-viral genome editing

Quantitative RT-PCR

Western blotting

Flow cytometry

Kinase assays

Proximity ligation assays

RNA-seq analysis

Bio-ID proteomics and data analysis

Phosphoproteomic analysis

Nfatc2 immunofluorescence and quantification

Single-cell Ca2+ measurements

Colony-forming assay

Co-immunoprecipitation and mass spectrometry

In vivo leukemia initiation assay

Assessment of CD25-ADC efficacy

Modeling of the RACK1-PKCδ–CD25 tail complex

MD simulation details

Population density of two conformations

Complex interaction energy analysis

BCR-ABL1 kinase reporter

Quantification and statistical analysis

Supplementary Material

Acknowledgments:

Funding:

Footnotes

Data and materials availability:

References and Notes

CD25⁺ subpopulations are small, quiescent, and drug-resistant leukemia cells

Absence of a clonal hierarchy between CD25⁺ and CD25⁻ leukemia cells

CD25 inhibits Ca²⁺ oscillations and NFAT signaling in tyrosine kinase–driven B-ALL

Single-cell Ca²⁺ measurements