Requirements for survivin in terminal differentiation of erythroid cells and maintenance of hematopoietic stem and progenitor cells

Because survivin deficiency results in periimplantation lethality (14), we used a conditional knockout strategy to study the requirement for survivin in in vivo hematopoiesis. Conditional gene-targeted mice, with all four exons of the survivin gene flanked by loxP sites (31), were bred to mice expressing Cre recombinase under the control of the endogenous Mx1 locus, which can be transiently activated in response to type-I IFN (37). Subsequently, mice harboring the Mx1-Cre transgene and either heterozygous or homozygous floxed alleles of survivin were treated with polyinosinic-polycytidylic acid (pI-pC), which is a synthetic double-stranded RNA, to induce deletion of survivin in the hematopoietic compartment. pI-pC treatment of Mx1-Cre/sur^fl/+ and Mx1-Cre/sur^fl/fl adult mice resulted in robust excision of the survivin floxed allele in BM cells as determined by PCR analysis (Fig. 1 A). To our surprise, administration of pI-pC to Mx1-Cre/sur^fl/fl mice caused rapid death (Fig. 1 B). Indeed, all of the animals became moribund within 10 d of pI-pC treatment. In comparison, the control littermates showed no ill effect after pI-pC injection, whereas the heterozygous-deleted mice showed 20% mortality. Analysis of peripheral blood, BM, and spleen (a prominent site of hematopoiesis in young mice) of survivin-deficient animals revealed a wide spectrum of hematopoietic defects. As clearly evident in Fig. 1 D, there was almost a complete ablation of the BM of Mx1-Cre/sur^fl/fl mice. Consistent with this gross BM depletion, peripheral blood smears from survivin-deleted animals exhibited marked pancytopenia, including reticulocytopenia (Fig. 1 E). In addition, a plethora of erythrocyte abnormalities, such as red cell fragmentation, anisopoikilocytosis, and hypochromia, were visible. Furthermore, Mx1-Cre/sur^fl/fl mice had smaller and extremely hypocellular spleens (Fig. 1, C and F) caused by a reduced white and red pulp. The red pulp, which is a site of normal extramedullary hematopoiesis in young mice, was paucicellular. Most dramatic was the complete absence of nucleated erythroid progenitors that stained positive for the erythroid-specific marker Ter119 in the spleens of Mx1-Cre/sur^fl/fl mice, as shown by immunohistochemical analysis for Ter119 (Fig. 1 G). Control mice that were treated with pI-pC did not display any of these phenotypes (Fig. 1, B–G). Interestingly, however, we observed an intermediate phenotype in survivin heterozygous-deleted mice (as discussed in Haploinsufficiency of survivin leads to defects in erythropoiesis).

To better understand why the animals die within 2 wk of survivin depletion, we repeated the pI-pC injections on another group of mice (8 wk of age) and analyzed their complete blood counts (CBCs) on days 7 and 11 after treatment. CBC analysis at day 7 revealed that survivin-depleted mice had lower hemoglobin and white cell and platelet counts compared with the control animals (unpublished data). By day 11, the survivin-depleted mice developed definitive anemia. Thrombocytopenia and leukopenia were reflected in the automated CBC, as well as on the peripheral blood smears shown in Fig. 1 E; homozygous-deleted animals displayed pancytopenia, with multiple defects in circulating erythrocytes. Therefore, we believe that the development of anemia in the periphery is mostly caused by underproduction, which may or may not be exacerbated by some peripheral destruction. Based on the presence of fragmented red cells in the periphery, we hypothesize that a thrombotic microangiopathic process might contribute to the peripheral destruction of red cells in some of the mice. Furthermore, to investigate if the rapid morbidity and mortality of the survivin-depleted mice was caused by injury to nonhematopoietic tissues, gross and microscopic examination was performed on the major organs from control and survivin-depleted mice. There was no evidence of organ damage in the kidneys, lungs, livers, and hearts of the survivin-depleted mice (unpublished data). Under the circumstances, we speculate that the morbidity and mortality observed in the survivin-depleted animals is secondary to BM failure. It is possible that anemia, infections caused by neutropenia, or hemorrhage caused by thrombocytopenia contribute to early morbidity and mortality in these animals.

Because the BM histology indicated that survivin deletion affected multiple hematopoietic lineages, we used flow cytometry to examine specific cell populations in either the BM or spleen of control, Mx1-Cre/sur^fl/+, and Mx1-Cre/sur^fl/fl mice after pI-pC injection. In contrast to littermate controls, Gr-1⁺Mac-1⁺ myeloid cells and CD41⁺ megakaryocytes were both reduced in the BM of homozygous-deleted mice as early as 36 h after pI-pC induction (Fig. 2, A and B). C-kit⁺ progenitors were also dramatically diminished in the spleen of homozygous-deleted mice by 10 d of pI-pC administration (Fig. 2 C). Consistent with these results, total BM cells, isolated from homozygous-deleted mice 1 d after pI-pC injection, failed to give rise to any myeloid, erythroid, or megakaryocytic colonies in colony-forming assays (unpublished data). The rapid time frame in which hematopoietic progenitors disappear after survivin excision suggests survivin is critical for the maintenance of hematopoietic cells.

We next focused on analyzing the effect of survivin deletion in the erythroid lineage. During their maturation, erythroid cells progressively become smaller and exhibit chromatin condensation, hemoglobinization, cell cycle exit, and enucleation. As erythroblasts undergo terminal differentiation, they also down-regulate expression of the transferrin receptor CD71 and up-regulate the erythroid-specific marker Ter119. Erythroid cells can be segregated into four subsets by their CD71 and Ter119 expression level (R1 proerythroblasts, Ter119^lowCD71^high; R2 basophilic erythroblasts, Ter119^high CD71^high; R3 polychromatophilic erythroblasts, Ter119^high CD71^med; and R4 orthochromatophilic erythroblasts and reticulocytes, Ter119^highCD71^low) (38). To examine the consequences of survivin deletion specifically on erythropoiesis, we analyzed the distribution of erythroid subsets in the spleen of 1-mo-old Mx1-Cre/sur^fl/fl mice 10 d after pI-pC administration. Compared with their littermate controls, homozygous-deleted mice exhibited a marked reduction in the percentages of R1 and R2 subsets and an increase in the percentage of the R4 subset (Fig. 2 D). To investigate the erythroid perturbation more carefully, we isolated R2 and R4 erythroid cells from pI-pC–treated control and Mx1-Cre/sur^fl/fl spleens by FACS for morphological evaluation. Because survivin deletion results in rapid BM ablation and early death in Mx1-Cre/sur^fl/fl mice, the experiment was performed at an earlier time point, 4 d after pI-pC injection. Survivin excision in total spleen cells was confirmed by PCR analysis (Fig. 3 A). At this early time point, the spleens from survivin homozygous-deleted mice were comparable in size to those of control littermates and harbored similar absolute numbers of R4 cells, but strikingly fewer R2 cells (Fig. 3 B). Morphological evaluation of the R4 population by both May-Grünwald Giemsa and benzidine staining revealed that the majority of cells in survivin-deleted animals were mature, hemoglobinized, and enucleated erythrocytes that were indistinguishable from those of controls (Fig. 3 C). These data support an essential role for survivin in proliferating proerythroblasts and basophilic erythroblasts. In contrast, R4 cells, which had already exited the cell cycle and extruded their nuclei, were not affected by survivin loss, and the total number of cells within the R4 population was not significantly altered, resulting in an apparent relative increase in the percentage of cells in the R4 gate.

To determine whether aberrant proliferation or increased cell death was responsible for the erythroid defect, we next analyzed the cell cycle profile and DNA content of survivin-depleted erythroid precursors. BM cells from control and Mx1-Cre/sur^fl/fl mice were harvested 48 h after pI-pC treatment and stained with DAPI and antibodies against CD71 and Ter119. Analysis of survivin-deficient erythroblasts showed increased G₁ and decreased S phase cells, indicating that these cells were either cell cycle-arrested or had exited the cell cycle faster (Fig. 4 A). In addition, survivin-deficient erythroblasts exhibited an increase in cells with ≥4 N DNA content, as well as a moderate increase in cells with sub-G₁ DNA (Fig. 4 B). The increase in the number of cells with sub-G₁ DNA suggests that some of the cells have undergone cell death, either as a consequence of aberrant cell cycle progression or initiation of apoptosis.

We next investigated whether the death seen in these cells was a consequence of activation of the intrinsic mitochondrial apoptotic pathway. If survivin depletion resulted in initiation of this pathway, then ectopic Bcl-2 expression in these cells might rescue the phenotype caused by survivin loss. To address this question, we used vav-Bcl-2 transgenic mice, in which constitutive expression of the human Bcl-2 cDNA is restricted to nucleated hematopoietic cells by the vav promoter (39). To assess the ability of vav-Bcl-2 to promote the survival of erythroid progenitors, lineage-negative BM cells from vav-Bcl-2 transgenic mice or their wild-type littermates were subjected to varying doses of gamma irradiation (0, 200, or 300 rads) and plated in erythroid colony-forming assays with IL3, SCF, and EPO. As demonstrated in Fig. 4 C, vav-Bcl-2, indeed, promoted the survival of erythroid progenitors in response to gamma irradiation. In fact, after treatment with 200 or 300 rads of gamma irradiation, the vav-Bcl-2 transgene led to a twofold increase in erythroid colony formation in vitro, compared with wild-type controls. Furthermore, vav-Bcl-2 also prolonged the survival of erythroid progenitors after 3 d of EPO deprivation (unpublished data). To determine whether overexpression of Bcl-2 can rescue the defects caused by survivin deficiency, we crossed vav-Bcl-2 transgenic mice to Mx1-Cre/sur^fl/fl mice to generate Mx1-Cre/sur^fl/fl; Bcl-2 animals. Bcl-2 and survivin expression were detected by Western blot analysis of protein extracts prepared from the spleens of pI-pC–treated control, Bcl-2 transgenic, or Mx1-Cre/sur^fl/fl; Bcl-2 mice (Fig. 4 D). Although ectopic expression of Bcl-2 under the vav promoter protected erythroid cells on a wild-type survivin background, it failed to rescue the smaller spleen, loss of erythroid progenitors, and eventual demise of Mx1-Cre/sur^fl/fl; Bcl-2 mice after survivin deletion (Fig. 4 E).

We previously reported that heterozygous loss of survivin caused a nearly 50% reduction in erythroid colony formation, but a much less dramatic change in the number of megakaryocytic colonies (35). These findings suggested that the erythroid lineage might be uniquely sensitive to the dosage of survivin. In this current study, we observed defects in a proportion of the heterozygous-deleted mice that are consistent with a dose-dependence for survivin in erythroid cells. First, 2 out of the 10 Mx1-Cre/sur^fll+ animals, but none of the control animals, died within 10 d of pI-pC treatment (Fig. 1 B). Second, four out of the five BM sections from heterozygous-deleted mice were identified as harboring fewer erythroid precursors in a blinded analysis by a pathologist (S. Gurbuxani; Fig. 1 D and not depicted). Additionally, peripheral blood smears from these mice displayed an increase in hypochromic erythrocytes (Fig. 1 E, red arrows), which were not observed in any pI-pC–treated control animals.

Furthermore, analysis of the CD71/Ter119 erythroid population by flow cytometry revealed that 20% of the heterozygous-deleted animals harbored an altered proportion of erythroid cells, with affected animals displaying an increased proportion of R2 cells and a concomitant decrease in the R4 population (Fig. 2 D and Fig. 5 A). In the data depicted in Fig. 5 A, which are representative of the affected group of animals, nearly half as many R4 cells and a greater number of R2 cells were isolated from the heterozygous-deleted spleen in comparison to the control spleen. To further explore the nature of this defect, R2 and R4 cells from pI-pC–treated control, and Mx1-Cre/sur^fll+ mice were collected by FACS. Excision of the floxed allele was confirmed by PCR analysis (Fig. 5 B). May-Grünwald Giemsa staining of purified cells revealed that although the R2 progenitors appeared similar, the R4 erythroid population was distinctly different in the two groups (Fig. 5, C and D). Heterozygous survivin-deficient R4 cells exhibited a striking increase in the proportion of nucleated erythroblasts, with a concomitant decrease in enucleated erythrocytes. Interestingly, the R4 population of survivin heterozygous-deleted erythroblasts appeared immature and possessed less condensed, larger nuclei, which is indicative of a block in terminal differentiation. Although this phenotype was not fully penetrant, the megaloblastic features observed in a significant proportion of heterozygous-deleted animals suggest that survivin plays a novel role in erythroid terminal differentiation.

Whereas the preceding data suggest that survivin deficiency results in impaired proliferation and subsequent loss of hematopoietic progenitors, Mx1-Cre excision is not absolutely limited to hematopoietic cells (37). Therefore, to determine whether the requirement for survivin is cell autonomous, we transplanted total CD45.2⁺-nucleated BM cells from control, Mx1-Cre/sur^fl/+, or Mx1-Cre/sur^fl/fl mice, along with wild-type CD45.1⁺/CD45.2⁺ support cells, into lethally irradiated CD45.1⁺ congenic recipients (Fig. 6 A). Engraftment of donor cells was first verified by staining for CD45 surface marker expression in peripheral blood 5 wk after transplantation. The engraftment of CD45.2⁺ cells ranged from 7 to 69% (Fig. 6 B). Recipients were then treated with pI-pC, and the contribution of CD45.2⁺ cells to the hematopoietic compartment was assessed. Survivin excision led to pronounced depletion of CD45.2⁺ cells in the peripheral blood, supporting a role for survivin in hematopoietic stem and progenitor cells (Fig. 6 C). Administration of pI-pC to Mx1-Cre/sur^fl/fl transplanted mice also led to near complete disappearance of the BM CD45.2⁺ hematopoietic cells within 2 wk of pI-pC treatment in all five mice analyzed (Fig. 6, D and E). No significant changes were observed in either control or Mx1-Cre/sur^fll+ transplanted mice. These data show that there is a cell autonomous requirement for survivin in hematopoietic progenitors. Next, to determine whether HSCs are also susceptible to survivin deletion, we elucidated the origin of the HSC populations in control and Mx1-Cre/sur^fl/fl transplanted animals 10 wk after pI-pC administration. Flow cytometric analyses revealed that the lin⁻c-kit⁺Sca-1⁺ HSC populations in Mx1-Cre/sur^fl/fl transplanted mice were exclusively derived from wild-type CD45.1⁺/CD45.2⁺ support cells, whereas the HSC populations in control animals expressed either CD45.2⁺ alone or CD45.1⁺/CD45.2⁺ cell surface markers (Fig. 7).

We show that survivin is an essential gene in the adult and is indispensable for the proliferation and maintenance of both hematopoietic stem and progenitor cells. Similar to inducible deletion of the antiapoptotic Bcl-2 family member myeloid leukemia-1 (MCL-1) in mice, homozygous deletion of survivin resulted in rapid death, ablation of the BM, and loss of early BM progenitor populations, including HSCs (40). Consistent with our findings, T lineage-specific survivin knockout mice revealed critical requirements for survivin in the development and homeostasis of T cells (30, 31). In response to mitogen-induced proliferation, survivin-deficient T cells exhibited p53-mediated cell cycle arrest, a spindle formation defect, and spontaneous cell death. In addition, p53 inactivation or introduction of Bcl-2 failed to rescue the thymocyte developmental defects caused by survivin deficiency (30). In light of these published works, our study suggests survivin may function as both a chromosomal passenger protein and an inhibitor of apoptosis in the maintenance of hematopoietic stem and progenitor populations.

In addition, we demonstrate that haploinsufficiency of survivin affects erythroid terminal maturation. Erythroid progenitors undergo a series of three to four rapid “differentiation divisions” to give rise to mature, enucleated, and hemoglobinized erythrocytes (41). During this process, the G1 phase of the cell cycle is reduced from 11 h to 5 h, and the cell size is reduced from a 12-μm proerythroblast to a 4-μm erythrocyte (mouse). This altered cell cycle progression is accompanied by extensive changes in gene expression, including a rapid down-regulation of D-type cyclins and their partner, cyclin-dependent kinase type 4 (cdk4). In contrast, expression levels of S- and G₂–M-associated cell cycle regulators, such as cyclin A and cdk1/cdc2, are transiently up-regulated. In this study, we show a dose-dependent requirement for survivin in erythropoiesis. We previously reported that survivin levels increase as erythroid cells mature, with both mRNA and protein present in cells throughout even the late orthochromatic stage. In fact, erythroid cells express approximately fourfold higher levels of survivin mRNA and protein than the related megakaryocytic lineage during their terminal differentiation (35). Therefore, reduced levels of key mitotic regulators, including survivin, may compromise the efficiency of the cell cycle machinery and subsequently, lead to defects in erythroid maturation. Consistent with this model, mice heterozygous for BubR1, which is a spindle assembly checkpoint protein, exhibited decreased erythropoiesis and a higher frequency of anemia (42). Interestingly, stable association of BubR1 to kinetochores and maintenance of the spindle assembly checkpoint is critically dependent on survivin (12, 24). In the presence of reduced survivin levels, Aurora B kinase, INCENP, and Borealin may less efficiently localize to different components of the mitotic apparatus, including the inner centromeres, where the CPC plays an important auxiliary role in spindle checkpoint surveillance. Because Aurora B, INCENP, and survivin form a complex that is required for multiple aspects of mitosis and cytokinesis, a reduction in survivin levels likely disrupts proper targeting of Aurora B kinase to its substrates. It remains to be determined whether survivin has a novel role in erythroid terminal differentiation independent of its mitotic function.

Several preclinical studies have shown that disrupting survivin expression or function in cancer cells decreases their proliferation and enhances apoptosis (43). The absence of significant side effects on hematopoietic cells reported in these studies may be caused by local intratumor injection and minimal systemic dissemination of anti-survivin therapies. More recently, shepherdin, which is a small peptide that destabilizes survivin by blocking its interaction with the molecular chaperone heat shock protein 90 (Hsp90), has been shown to affect human CD34⁺ hematopoietic progenitor colony formation in vitro (44). In that study, treatment of CD34⁺ cells with shepherdin inhibited precursor erythroid burst-forming unit colony formation and, at slightly higher peptide concentrations, granulocyte–macrophage colony-forming units and granulocyte erythrocyte macrophage megakaryocyte colony-forming units. Our study confirms that the hematopoietic system is extremely sensitive to survivin disruption and emphasizes the need for innovative and selective anti-survivin cancer therapies.

Conditionally gene-targeted mice, with all four exons of the survivin gene flanked by loxP sites, were obtained from A. Winoto (University of California, Berkeley, Berkeley, CA) (31). Mx1-Cre transgenic mice harboring Cre recombinase under the control of the interferon-responsive Mx1 promoter in a C57BL/6 background were purchased from The Jackson Laboratory. Vav-Bcl-2 transgenic mice, in which constitutive expression of a human Bcl-2 cDNA is driven by the promoter of the vav gene, were provided by J. Adams (Walter and Eliza Hall Insitute of Medical Research, Victoria, Australia) and K. Macleod (University of Chicago, Chicago, IL) (39). To generate Mx1-Cre/sur^fl/+ mice, sur^fl/fl mice were crossed to Mx1-Cre transgenic mice. Mx1-Cre/sur^fl/+ mice were subsequently bred to sur^fl/fl mice to produce littermates with the following genotypes: sur^fl/+, sur^fl/fl, Mx1-Cre/sur^fl/+, and Mx1-Cre/sur^fl/fl. To generate Mx1-Cre/sur^fl/fl; Bcl-2 mice, Mx1-Cre/sur^fl/fl mice were crossed to Bcl-2 transgenic mice to obtain Mx1-Cre/sur^fl/+; Bcl-2 mice, which were then bred to sur^fl/+ mice. All animal research was approved by the University of Chicago and Northwestern University Institutional Animal Care and Use Committees.

Mouse genotyping was performed by PCR of tail DNA. For genotyping the Mx1-Cre transgene, the Mx1-Cre forward (5′-gcctgcattaccggtcgatgcaacga-3′) and Mx1-Cre reverse (5′-gtggcagatggcgcggcaacaccatt-3′) primers were used to amplify a 725-bp PCR product. Three primers (Adv17, Adv25, and Adv28) were used to genotype the wild-type, floxed, and Cre-deleted survivin allele, as previously described (31). For genotyping the vav-Bcl-2 transgene, primers against the SV40 poly(A) DNA sequence were used (39).

Newborn (10–14 d) mice received two intraperitoneal injections (13 μg/g body weight) of pI-pC (GE Healthcare) every other day to induce survivin deletion. Sternum BM and spleen were isolated 10 d later, fixed overnight in formalin (Sigma-Aldrich), and processed for histological analysis. Serial sections were stained with hematoxylin and eosin (HE), and peripheral blood smears were stained with May-Grünwald Giemsa using standard protocols. Immunohistochemistry was performed on deparaffinized spleen sections with anti-Ter119 antibody (BD Biosciences).

To obtain BM cells, mouse femurs and tibiae were flushed with PBS, and cells were separated by passage through a 20-gauge syringe. For spleen cells, spleens were cut into small pieces and subsequently disaggregated with an 18-gauge syringe. Cells were resuspended in PBS containing 1 mM EDTA and 1% bovine serum albumin (Sigma-Aldrich) to reduce aggregates and nonspecific staining, respectively. The cells were stained with different combinations of c-kit, Gr-1, Mac-1, CD41, CD71, and Ter119 antibodies, conjugated to APC, PE, or FITC (BD Biosciences). Finally, surface marker expression was analyzed using a FACSCanto flow cytometer (BD Biosciences) and FlowJo software. To isolate discrete erythroid populations, spleen cells were first stained with FITC-conjugated CD71 and PE-conjugated Ter119 antibodies. Then, based on the CD71 and Ter119 expression level, erythroid subpopulations were purified using a MoFlo-HTS cell sorter (DakoCytomation). FACS-sorted cells were cytospun onto slides and stained with either May-Grünwald Giemsa or benzidine Wright Giemsa for morphological evaluation.

Adult (6–8 wk) mice were injected intraperitoneally with one dose of pI-pC. BM cells were isolated 48 h later and stained with FITC anti-CD71 and PE anti-Ter119 antibodies. Cells were fixed with 2% paraformaldehyde (Sigma-Aldrich) and 70% ethanol to preserve cell surface antibody-conjugated fluorochrome staining. Subsequently, cells were treated with 2 N HCl/0.5% Triton X-100, followed by 0.1 M sodium tetraborate, and DNA was stained with DAPI (Sigma-Aldrich) at 4°C overnight. The cell cycle profile of CD71⁺ Ter119⁺ erythroid cells was analyzed using a LSR-II flow cytometer (BD Biosciences) and FlowJo software.

Primary mouse BM cells were collected from vav-Bcl-2 transgenic mice and their wild-type littermates and enriched for progenitors with the EasySep negative-selection mouse hematopoietic progenitor-enrichment kit (StemCell Technologies). The cells were either left untreated or irradiated with 200 or 300 rads in a Cesium-137 irradiator. To evaluate erythroid colony formation, 25,000 cells were plated in MethoCult 3234 (StemCell Technologies), supplemented with IL3, SCF, and 10 U/ml EPO. Pure burst-forming unit erythroid and mixed colonies were enumerated after 10 d.

C57BL/6 CD45.1⁺ congenic recipients (6–8 wk) were purchased from The Jackson Laboratory and placed on Bactrim-containing water 1 wk before lethal irradiation. Mice were irradiated with a total of 1,200 rads in split doses (800 rads, and then 400 rads 4 h later). Total nucleated BM cells (10⁶), marked by the CD45.2⁺ allele from untreated control, Mx1-Cre/sur^fl/+, or Mx1-Cre/sur^fl/fl mice, along with CD45.1⁺/CD45.2⁺ support cells (0.5 × 10⁶), were injected retroorbitally into lethally irradiated CD45.1⁺ recipients that were anesthetized with Nembutal (Abbott Laboratories). CD45.2⁺ and CD45.1⁺/CD45.2⁺ BM cells were allowed to engraft for 5 wk after transplantation. After the engrafting, peripheral blood from recipient mice was obtained by tail vein bleeding, subjected to red blood cell lysis, and stained with PE-Cy5.5–conjugated CD45.1 (eBioscience) and FITC-conjugated CD45.2 (BD Biosciences) antibodies to verify donor cell engraftment by flow cytometric analysis. A subset of the recipients was injected with 2 doses of pI-pC, whereas the remainder served as untreated controls. BM cells from pI-pC–treated and control animals were isolated 2 wk after pI-pC administration and analyzed for the presence of CD45.2⁺ and CD45.1⁺/CD45.2⁺ donor cells. For HSC analysis, BM cells from transplanted mice, 10 wk after 3 doses of pI-pC, were depleted of mature hematopoietic lineages, expressing surface antigens CD5, CD11b, B220, Gr-1, Ter119, and 7–4, with EasySep (StemCell Technologies). Lineage-negative cells were stained for APC anti–c-kit, PE-Cy7 anti–Sca-1 (eBioscience), FITC anti-CD45.2, and PE-Cy5.5 anti-CD45.1 antibodies and analyzed by FACSCanto flow cytometer.

The authors thank Dr. Astar Winoto for the survivin-floxed mice, Drs. Jerry Adams and Kay Macleod for the vav-bcl2 transgenic mice, Dr. Hanno Hock for advice on Mx1-Cre targeting, The University of Chicago Cancer Research Center, and the Flow Cytometry Core Facility for technical expertise. Additional thanks to Mitchell Weiss, Iannis Aifantis, Jonathan Licht, and Hanno Hock for critical review of the manuscript.

This research was supported, in part, by the Elsa U. Pardee Foundation and by National Institutes of Health grants CA101774 and DK074693. J.D. Crispino is a Scholar of the Leukemia and Lymphoma Society.

The authors have no conflicting financial interests.