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Limited responsiveness to inflammatory cytokines is a feature of adult hematopoietic stem cells, and contributes to the relative quiescence and durability of the stem cell population in vivo. Here we report that the executioner Caspase, Caspase-3, unexpectedly participates in that process. Mice deficient in Caspase-3 had increased numbers of immunophenotypic long-term repopulating stem cells in association with multiple functional changes, most prominently cell cycling. While these changes were cell autonomous, they reflected altered activation by exogenous signals. Caspase-3−/− cells exhibited cell type specific changes in phosphorylated members of the Ras-Raf-MEK-ERK pathway in response to specific cytokines while, notably, members of other pathways such as pSTAT3, pSTAT5, pAKT, pp38 MAPK, pSmad2 and pSmad3 were unaffected. Caspase-3 contributes to stem cell quiescence, dampening specific signaling events and thereby cell responsiveness to microenvironmental stimuli.
In organs with a high turnover of cells, stem cells must be strictly regulated to ensure the balance between the preservation of the stem cell pool and differentiation to continuously replace mature cells lost due to injury, disease, or wear. Continuous and precisely controlled elimination of cells by programmed cells death (apoptosis) prevents excess accumulation of normal cells and enables culling of defective cells. The balance between self-renewal, differentiation and apoptosis at the stem cell level can affect tissue function and is critical for the survival of the organism. We are interested in how these processes are integrated through a limited number of molecular events in these generally quiescent cell types. We focused on Caspase proteases as a group of enzymes capable of modulating multiple protein targets.
Caspase-3 was selected because of its central role in the apoptotic process as an executioner protease and because several lines of evidence implicate it in non-apoptotic functions. For example, altering the expression level or enzymatic activity of Caspase-3 in vitro has implicated Caspase-3 in specific stages of erythroid and megakaryocytic differentiation (Carlile et al., 2004; de Botton et al., 2002; Zermati et al., 2001). Intracellular Caspase-3 activation has been observed during differentiation in the absence of apparent apoptotic induction in neural and hematopoietic cells (Fernando et al., 2005; de Botton et al., 2002). In addition, Caspase-3 has a cell type-dependent effect on cell proliferation. While splenic B cells showed hyperproliferation in absence of Caspase-3 (Woo et al., 2003), bone marrow stromal stem cells exhibited reduced cycling activity (Miura et al., 2004). Yet, Caspase-3 deficiency did not alter the frequency of apoptosis in either of the cell types.
We report here that alteration in Caspase-3 expression perturbs the homeostasis of primitive hematopoietic cells. Caspase-3 deficient stem cells displayed accelerated proliferation and retarded differentiation. At the molecular level Caspase-3 altered the sensitivity of primary hematopoietic cells to cytokine stimulation through modulation of specific signaling pathways. The data indicate a novel role for Caspase-3 as a governor, down-modulating stem cell responsiveness to environmental signals and point to proteases as central to the processes regulating the stem cell compartment.
To establish the potential for Caspase-3 to play a regulatory role in primitive hematopoietic cell function, we assayed wild-type populations of blood cells for the expression of Caspase-3 message and protein. We detected high levels of Caspase-3 mRNA in sorted primitive bone marrow compartments defined as Lineage negative, cKit positive and Sca-1 positive population (LK+S+), enriched for stem cells, and also in the more mature progenitor population, defined as Lineage negative, cKit positive, Sca-1 negative (LK+S−) (Figure 1A). Caspase-3 protein was also detectable in the LK+S+ and LK+S− populations of mouse bone marrow assayed by flow cytometry (Figure 1B).
To analyze whether Caspase-3 affected hematopoiesis in vivo, we used mice engineered to be deficient for Caspase-3 (Woo et al., 2003). Analysis of peripheral blood cell counts revealed unchanged levels of red blood cells and platelets, and a slight but not significant decrease in the total white blood count. Differential analysis of white blood cells showed a significant reduction in both B and T lymphocytes with no significant reduction in myeloid cells in Caspase-3−/− (KO) animals (Figure S1). KO mice were smaller in body size than were their wild type littermates, as also noted previously by others (Kuida et al., 1996; Woo et al., 1998; Miura et al., 2004). The total bone marrow cellularity per femur and the normalized, relative cell number per gram of body weight were both significantly reduced in KO mice (Figure S2).
The effect of Caspase-3 deficiency on primitive subpopulations in the bone marrow was then examined. We observed a doubling in the frequency of LK+S+ cells in KOs, compared with their wild type littermates. When the LK+S+ population was further subfractionated into the long term HSC-containing population LK+S+CD34lowFlk2low, the short term repopulating HSC population LK+S+CD34+Flk2low and the multipotent progenitor population LK+S+CD34+Flk2+, an increase in the frequency of all three populations in the absence of Caspase-3 was noted. Most striking was a ~3 to 4-fold increase in the immunophenotypic long term repopulating HSC fraction (Figure 1C). In contrast, no increase was observed in the frequency of the progenitor-containing population (LK+S−), nor was there evidence for an increase in the frequency of colony forming cells (CFC), an in vitro functional read-out of progenitor cells (Figure S3). Analysis of different subpopulations within the progenitor pool revealed similar frequencies of the granulocyte-macrophage progenitors (GMP) and megakaryocyte-erythroid progenitors (MEP) between the genotypes, but a significant decrease in the common myeloid progenitors (CMP) in KO mice (Figure 1C). No differences in the frequency of common lymphoid progenitors (CLP) between the genotypes were observed. These data indicate a preferential expansion of primitive hematopoietic populations with a concomitant reduction in early myeloid progenitors. Further quantitation of the number of stem cells by competitive, limit dilution transplantation was not possible because we defined abnormalities in the proliferation and differentiation of primitive cells in the absence of Caspase-3 as described below. These functional changes make the standard quantitation of stem cells by measuring the contribution of donor cells to the blood of recipients, uninterpretable.
Given that Caspase-3 is a known effector of apoptosis, we questioned if the observed differences in primitive subpopulations in KO mice could be accounted for by altered susceptibility to apoptotic stimuli. Using the AnnexinV/DAPI assay, we analyzed freshly isolated bone marrow cells for their expression of AnnexinV and the exclusion of the nuclear dye DAPI. No significant difference in the frequency of AnnexinV+/DAPI− cells within the stem and progenitor cell-containing populations was observed (Figures 2A and S4). The TUNEL assay, which detects apoptosis at the later DNA fragmentation stage, is not reliable in vivo due to the elimination of dying cells by resident macrophages. We used this assay in vitro to measure susceptibility to apoptosis inducing signals using cytokine starvation, gamma-irradiation or heat-shock (Williams et al., 1990; Domen et al., 2000; Kadhim et al., 1995) of FACS-sorted LK+S+ cells; hematopoietic stem cells do not express the Fas receptor (Aguila and Weissman, 1996; Bryder et al., 2001). With cytokine starvation or gamma irradiation, we detected a reduction in the frequency of apoptotic cells in the absence of Caspase-3 (Figures 2B and S5); however, no difference was observed after heat shock treatment (data not shown). Given the lack of evidence for increased apoptosis in vivo and the conflicting results of the in vitro assays, we regarded apoptosis as being unlikely by itself to account for the phenotypic changes observed in Caspase-3 deficient mice.
We therefore analyzed whether Caspase-3 affects proliferation of primitive hematopoietic cells in vivo. Using a BrdU incorporation assay we determined the frequency of LK+S+ cells that have initiated at least one cell division within 24 hours of BrdU administration. LK+S+ cells from Caspase-3−/− mice displayed a significantly higher proliferation rate than their wild type counterparts (Figure 2C). The analysis of different stages of the cell cycle within the LK+S+ population revealed a shift from G0 towards G1, ultimately resulting in a higher percentage of cells that were in mitotic (S/G2/M) stages of the cell cycle (Figure 2D). This increase by fourfold in cells in G1 is dramatic and of greater magnitude than that previously observed by us and others in other genetic mutants affecting hematopoietic stem cell cycling (Cheng et al., 2000; Hock et al., 2004; Janzen et al., 2006). These data demonstrate that Caspase-3 limits cell cycling in primitive hematopoietic cells.
Caspase-3 KO mice display growth retardation due to a delay in bone formation (Miura et al., 2004). To examine whether microenvironmental changes present in Caspase-3 deficient mice are responsible for the phenotype within the stem cell compartment we transplanted KO and WT bone marrow cells into lethally irradiated WT recipients and monitored their peripheral blood and bone marrow reconstitution. A congenic mouse system was utilized such that donor cells all expressed the pan-hematopoietic surface marker CD45.1 and were transplanted into CD45.2 allele-bearing hosts (Figure 3A). While we noted significant reductions in peripheral blood myeloid and lymphoid cells at all time points examined (Figure 3B), a consistent increase in the primitive LK+S+CD34low cell population was observed in the bone marrow of WT recipients of a Caspase-3 null graft (Figure 3C). In contrast, the progenitor containing population (LK+S−) was reduced in recipients of mutant bone marrow, a phenotype reminiscent of the distribution of these populations observed in KO bone marrow prior to transplantation. To further exclude the effect of the Caspase-3−/− bone marrow microenvironment on adult hematopoiesis, we transplanted wild type bone marrow into lethally irradiated KO and littermate control mice. Analysis of recipient peripheral blood revealed no differences in engraftment between the genotypes at all time points examined (Figure 3D), indicating that a Caspase-3 deficient microenvironment does not impact the engraftment kinetics of hematopoietic stem cells.
Several proteins involved in cell cycle control have been demonstrated to be essential for the preservation of the repopulation ability of hematopoietic stem cells (Cheng et al., 2000; Hock et al., 2004; Janzen et al., 2006). To analyze whether Caspase-3 deletion has an impact on the maintenance of the stem cell pool we performed sequential bone marrow transplantations (Figure 4A). While a consistently reduced number of mature blood cells were detected in recipients of the Caspase-3−/− bone marrow at each round of the transplant, a significantly larger pool of mutant LK+S+ cells was apparent even after the 4th serial transplant (Figure 4B). These data indicate a preserved, cell autonomous, self renewal ability of immunophenotypic hematopoietic stem cells in the absence of Caspase-3, despite the observed elevation of proliferation rate within this population.
Next we sought to investigate the ability of KO HSCs to repopulate irradiated hosts in the presence of competing wild-type donor cells. Peripheral blood and bone marrow samples were assayed 16 weeks following transplantation for the relative contribution of donor cells to the blood and bone marrow based on their expression of the CD45.1 or CD45.2 antigens (Figure 4C). While we observed comparable peripheral blood cell production and distribution between the genotypes, a disproportionate increase in mutant LK+S+ cells was observed, reflecting either the increase in stem cells present in the original graft or increased reconstitution of the stem cell compartment by higher proliferating Caspase-3−/− cells (Figure 4D). Of note, the increase in Caspase-3−/− primitive cells was paired with a relative reduction of more mature cells. In association with unaltered apoptosis, these data are consistent with a negative impact on the differentiation of immature cells with the loss of Caspase-3 expression.
The analysis of peripheral blood cells from unmanipulated Caspase-3−/− mice revealed a striking reduction in mature B lymphocytes. The deficit of mature B cells in the absence of Caspase-3 was exacerbated in the various transplant settings studied (Figures S1, 3B, 4B). The regulatory role of Caspase-3 on splenic B cell proliferation has been established previously (Woo et al., 2003). However, the requirement for Caspase-3 in the differentiation of earlier B cell subpopulations has not been explored. Available markers can distinguish steps in early B lineage differentiation in a precise manner, and we, therefore, analyzed KO bone marrow for its composition of B cell precursors. We observed a relative increase in the pro-B cell population, and a reduction in the subsequent pre-B population of the B lineage (Figure 5A), consistent with a partial developmental block at this important transition. The molecular processes responsible for the pro-B to pre-B cell differentiation step are well characterized and require intact signaling through the interleukin-7 (IL-7) receptor activating the extracellular signal-regulated kinase (ERK) signaling pathway (Fleming and Paige, 2001; Milne et al., 2004). The Ras-Raf-MEK-ERK signaling pathway controls fundamental cellular processes in a cascade of phosphorylation events (O’Neill et al., 2004). Membrane-proximal activation of Ras family members in turn phosphorylate Raf, followed by the serine-threonine kinase MEK, which finally phosphorylates ERK1/2. Numerous receptors utilize this pathway and it is activated in hematopoietic precursors following stimulation with specific cytokines (Uchida et al., 2001; Akbarzadeh et al., 2002). Thus, we addressed whether Caspase-3 deficiency affects the IL-7-mediated ERK signaling cascade. Using IL-7-dependent WT and Caspase-3−/−pro-B cell lines, we examined the kinetics of ERK phosphorylation after stimulation with IL-7. While we observed phosphorylation of ERK1/2 in wild-type cells, no pERK was detectable in Caspase-3 null cells (Figure 5B). However, a similar effect was not noted in the JAK-STAT pathway as levels of pSTAT5 were unaffected by the absence of Caspase-3.
Cytokines known to alter stem cell behavior, including kit ligand, flt-2 ligand, IL-3 and TPO utilize receptors that activate the Ras-Raf-MEK-ERK pathway (Jalal et al., 2004; Nonami et al., 2004; Robinson et al., 1998; Rouyez et al., 1997). We therefore examined whether Caspase-3 modulates the intensity of signal transduction within cells enriched for HSCs to account for the differences in proliferation and differentiation in the knock-out animals. Bone marrow LK+S+ cells from WT and Caspase-3 KO mice were isolated and subjected to cytokines with recognized effects on hematopoietic stem cells including kit ligand or stem cell factor (SCF) interleukin-3 (IL-3), thrombopoetin (TPO), interleukin-6 (IL-6), Stromal Derived Factor (SDF), and Angiopoietin-1 (Ang1). Specific cytokines demonstrated altered activation of ERK in the setting of Caspase-3 deficiency. Elevated phosphorylated ERK in Caspase-3 deficient LK+S+ cells compared with WT controls was noted at multiple time points after stimulation by SCF or IL-3 (Figure 6A). Of note, however, the effects on pERK levels were highly cytokine specific as TPO resulted in a decrease in pERK levels in knock-out LK+S+ cells (Figure 6A) while IL-6, SDF and Ang1 had no activating effect on the ERK pathway (Figure 6B). These data indicate that the effects of Caspase-3 are not common to all activators of the ERK pathway. Therefore, Caspase-3 alters signaling dependent upon both the cell type and the specific stimulus.
To determine the critical step at which the Caspase-3 affects ERK signaling we examined the activity of proteins upstream of ERK. Analysis of MEK phosphorylation upon stimulation of WT and Caspase-3−/− BM cells with SCF revealed elevated levels of activated MEK in Caspase-3 null cells compared with wild-type controls (Figure 6C), indicating that the effect of Caspase-3 was upstream of both ERK and MEK.
Next we assessed whether the differences noted were restricted to the ERK pathway, or if Caspase-3 deletion affects signaling across different pathways. We analyzed the activation of p38MAPK with cytokines that did (SCF) or did not (IL-6) have altered ERK phosphorylation in the KO LK+S+ cells. No changes in p38MAPK phosphorylation were noted at multiple time points (Figure S6). Similarly, when we examined pathways activated with particular cytokines such as AKT activation after SCF stimulation or Stat3 activation after IL-6 or TPO signaling, we could detect no effect of Caspase-3 deficiency (Figure 6D).
To further establish the mechanism by which Caspase-3 affects signal transduction, we asked whether Caspase-3 is activated immediately following binding of a growth factor to its corresponding receptor. Such a finding would suggest that Caspase-3 is actively involved in signal transduction. We examined if Caspase-3 cleavage was detectable in WT LK+S+ cells following stimulation with cytokines. As shown in Figure S7, no cleaved Caspase-3 was observed after stimulation of bone marrow cells with SCF, despite apparent activation of ERK. To exclude the possibility that low levels of Caspase-3 activation occurred, but were undetectable due to technical limitations, we utilized Caspase-3 specific inhibitors to eliminate its enzymatic activity. However, short-term treatment of WT bone marrow cells with the Caspase-3 inhibitor z-DEVD.fmk or the pan-Caspase inhibitor z-VAD.fmk did not prevent ERK activation in LK+S+ cells (Figure S8) or IL-7 dependent pro-B-cells (Figure S9). One could also speculate that Caspase-3 activation during specific cellular processes, as it was reported in other cells types (Carlile et al., 2004; Fernando et al., 2005), may lead to a certain degree of protein turnover. In Caspase-deficient cells the lack of this enzymatic activity may cause an accumulation of some mediators of cell signaling. However, no differences in the expression levels of some selected participants of different signaling pathways were detectable at steady state (Figure S10). These data suggest that Caspase-3 has an indirect effect on ERK signal transduction, rather than a direct, immediate effect mediated by its enzymatic activity.
Others have reported that TGF-beta may alter growth factor responsiveness, so we decided to analyze effects of Caspase-3 on TGF-beta response. Using the recently described assay to visualize signal activity in single cells by immunostaining (Yamazaki et al., 2006; Ema et al., 2006) we demonstrated low endogenous TGF-beta signaling in primitive cells independent of the Caspase-3 genotype (Figure S11). Additional stimulation with TGF-beta increased the phosphorylation of Smad2 and 3 with no significant differences between wildtype and KO cells (Figure S11 and S12). To test whether TGF-beta treatment would affect the clonal expansion of primitive hematopoietic cells, we sorted single wildtype and Caspase-3−/− LK+S+ cells and cultured them in presence of different cytokine cocktails. No significant differences were noted in the effects of TGF-beta between the wild type and knock-out cells (Figure S 13). Therefore, Caspase-3 seems to selectively affect the Ras-Raf-MEK-ERK pathway in a cell type and cytokine specific manner. Limiting cell numbers precludes further detailed analysis of the affected events.
The processes of cell cycle control, self-renewal, differentiation and apoptosis are closely intertwined in controlling stem cell fate during development and in adult homeostasis. Here we demonstrate that changes in expression of a central member of the apoptosis machinery, the protease Caspase-3, have multiple effects on hematopoietic stem cell homeostasis. Mice engineered to be deficient for Caspase-3 demonstrated a striking increase in the abundance of the most immature hematopoietic cells by the only measure feasible in this model, immunophenotype. However, the accumulation of primitive hematopoietic cells was most likely not due to the changes in the rate of apoptosis, since we did not observe a difference in the frequency of apoptotic events in any subfraction of the primitive compartment. Although, we demonstrated a reduction in the susceptibility to forced induction of apoptosis in an in vitro assay, the physiological relevance of these results remain unclear. We can not definitely rule out that differences in the frequency of apoptotic events occur in vivo may escape our detection and yet still have a cumulative effect on stem cell homeostasis. However, this is likely to only partly explain the phenotype we observed in Caspase-3 mutant HSCs.
The increased magnitude of the HSC-containing compartment was primarily associated with accelerated proliferation of KO cells. Previously, we and others had shown that cytokine receptors were expressed on hematopoeitic stem cells and yet these cells were known to be cytokine resistant. (Berardi et al., 1995; Cheng et al., 1996; Ivanova et al. 2002; Kiel et al. 2005) We had hypothesized and shown that this was associated with increased levels of cyclin dependent kinase inhibitors and when these molecules were deleted, stem cell cycling increased. (Cheng et al., 2000) However, the ability of cells to enter into cycle with that change may not be reflective of why stem cells are relatively cytokine resistant. The data presented here suggest another possible mechanism: Caspase-3 limits their ability to signal via specific pathways of activation.
Entry into cycle was not the only abnormality of the Caspase-3 null cells and increased cycling does not necessarily increase the stem cell pool. Deletion of the cell cycle regulator p21Cip1/Waf1 leads to increased stem cell proliferation and pool size in some mouse strains, but this is not sustained and stem cells prematurely exhaust under stress (Cheng et al., 2000). An accelerated proliferation of hematopoietic stem cells due to deletion of PTEN, a negative regulator of the PI3K-AKT signaling pathway, is associated with the depletion of hematopoietic stem cell pool within weeks after PTEN deletion (Yilmaz et al., 2006). However, increased proliferation in Caspase-3 deficient cells was associated with a preserved stem cell pool even under the stress conditions of transplantation. Our data suggest that Caspase-3 deficient cells exhibit differentiation delay at specific stages of hematopoiesis and it may be that this effect contributes to the increased primitive cell population.
The molecular basis for the phenotype of Caspase-3 null cells is at least in part due to an increased sensitivity of mutant primitive cells to exogenous signals. We defined the Ras-Raf-MEK-ERK pathway to be altered in at least two types of KO hematopoietic cells, lymphocytes and stem cells. The direction of modulation was curiously opposite in the two cell types. This coupled with the data that Caspase-3 activity at the time of cell stimulation is not critical for the altered signaling phenotype, suggest that Caspase-3 exerts its effect indirectly. The indirect effects are cell type- and cytokine-specific and appear to not overlap with the AKT or JAK-STAT pathways and not to be due to changes in TGF-beta signaling as these were not different between the genotypes.
Indirect effects of Caspase-3 on activating the Ras-Raf-MEK-ERK cascade proteins may be due to multiple actions. Accumulation of a number of different Caspase-3 target proteins is most likely and may still occur, despite the lack of short term effects by the enzymatic inhibitors. In mesenchymal stem cells, Miura et al noted increased Smad-2 in the absence of Caspase-3 (Miura et al., 2004). It is tempting to speculate that known targets of Caspase-3 such as Raf-1 (Widmann et al., 1998), may be the basis for the altered cytokine responsiveness we observed, but this is one of many possibilities and broadly examining protein modifications in a stem cell population is not currently feasible. Our data from the IL-7 stimulation in B cell progenitors suggest that the signaling alterations are highly cell type specific. Given the cell context dependence and multiple known targets of Caspase-3, it is not our contention that the Ras-Raf-MEK-ERK pathway is the only mechanism for the Caspase-3 effects. Rather, we posit that this pathway is a contributor to altered cytokine sensitivity and that cytokine response can account for some, if not all, of the functional changes observed.
In summary, the data presented here indicate that Caspase-3 is involved in cytokine-mediated signaling by different receptors in primitive hematopoietic cells. Caspase-3 alters signal transduction by limiting activation of the Ras-Raf-MEK-ERK pathway. We postulate that Caspase-3 prevents the accumulation of activated components of the pathway and thereby restricts cytokine response, serving as a molecular component of the known quiescence and cytokine resistance of hematopoietic stem cells. As such, it is a mediator of stem cell homeostasis, evident in the alteration of function of the stem cell compartment in its absence. Alteration of Caspase-3 levels either pharmacologically or genetically may change the size and activity of the stem cell compartment by means not reflected in vivo by measures of apoptosis. We therefore conclude that Caspase-3 modulates stem cell responsiveness to the cytokine cues of the stem cell microenvironment selectively dampening specific signaling pathways to maintain stem cell quiescence.
C57Bl/6 wild type and Caspase-3 mice were bred in-house in a pathogen-free environment. The Caspase-3 KO mouse was generated as previously described (Woo et al., 1998) and backcrossed to C57Bl/6 for more than 8 generations. The Subcommittee on Research Animal Care of the Massachusetts General Hospital (MGH) approved all animal work according to federal and institutional policies and regulations.
WT and Caspase-3−/− IL-7 dependent pro-B cell lines were generated and maintained as described previously (Fleming and Paige, 2001). Bone marrow was harvested as previously described (Cheng et al., 2000) and cultured in CFU-C assays according to the manufacturers’ protocols (Stem Cell Technologies). Sorted L−K+S+ cells were cultured in HSC medium: X-Vivo 15™ (Cambrex) supplemented with 10% detoxified BSA (StemCell Technologies), 50 U/ml penicillin (BioWhittaker), 50 U/ml streptomycin (Cellgro), 2 mM L-glutamine (BioWhittaker), and 0.1 mM 2-mercaptoethanol (Sigma-Aldrich). Prior apoptosis induction LK+S+ cells were cultured in presence of 50 ng/ml rmSCF, 50 ng/ml rmTPO, 50 ng/ml rmFlt-3L (all from PeproTech) and 20 ng/ml rmIL3 (R & D Systems). 48 h after initial stimulation cells were maintained in culture in presence of 10 ng/ml rmSCF and 10 ng/ml rmTPO. After total culture time of 96 h cells were washed with sterile PBS and subject to apoptosis induction. Cytokine withdrawal assay was performed by washing cells twice with PBS and placing in HSC medium with antibiotics and BSA but without cytokines. Detection of apoptotic events was performed 48 hours and 72 post cytokine withdrawal with AnnexinV-DAPI and TUNEL respectively. Mitogenic stimulation assay was performed by continuously culturing of LK+S+ cells in HSC medium in presence of 10 ng/ml rmSCF and 10 ng/ml rmTPO. Detection of apoptotic events was performed 4 and 5 days after beginning of cell culture with AnnexinV-DAPI and TUNEL assays respectively. Induction of apoptosis by DNA damage was performed by gamma-irradiation with 2 Gy and additional culturing of cells in HSC medium in presence of 10 ng/ml rmSCF and 10 ng/ml rmTPO. Detection of apoptotic events was performed 24 and 48 hours after irradiation with AnnexinV-DAPI and TUNEL assay respectively. Heat shock treatment was performed by incubation of the cells for 60 min at 42 °C and subsequently culturing in HSC medium in presence of 10 ng/ml rmSCF and 10 ng/ml rmTPO prior to detection of apoptosis 24 and 48 hours after heat shock treatment with AnnexinV-DAPI and TUNEL assay respectively. Clonal analysis were performed by sorting single LK+S+ cells in 96 well plates in HSC-medium supplemented with 50 ng/ml SCF, 50 ng/ml TPO, and 50 ng/ml Flt3-ligand, or with 50 ng/ml SCF, 50 ng/ml IL-6 (PeproTech) and 10 ng/ml IL-3, where indicated 5 ng/ml TGF-beta (R & D Systems) was added. Colonies were scored at day 5 and day 11 for the presence of proliferating clones by light microscopy.
Lineage staining utilized a cocktail of biotinylated anti-mouse antibodies to Mac-1α (CD11b), Gr-1(Ly-6G & 6C), Ter119 (Ly-76), CD3ε, CD4, CD8a (Ly-2), and B220 (CD45R) (BD Biosciences). For detection and sorting we used streptavidin conjugated with PE/Cy7, c-Kit-APC (CD117), Flk2-PE (CD135), CD34-FITC (all from BD Biosciences) and Sca1-PE/Cy5.5 (Ly 6A/E, Caltag). For congenic strain discrimination anti-CD45.1-PE and anti-CD45.2 FITC antibodies (BD Biosciences) were used. To assess the cell cycle in the primitive population, BM cells were stained with lineage antibodies and c-Kit as above, anti-Sca-1 PE-Cy-7 (BD-Biosciences), Pyronin Y (RNA dye) and Hoechst 33342 (DNA dye) as described (Cheng et al., 2000). For BrdU incorporation we used the APC-BrdU Flow Kit (BD Biosciences) following a single intraperitoneal injection of BrdU (1 mg per 6g of body weight) and an additional administration of of 1 mg/ml of BrdU (Sigma) mixed to drinking water for approximately 24 hours. Surface staining for lineage markers was performed as above, Sca1-PE/Cy5.5, c-Kit-APC/Cy7 (eBioscience), and CD34-FITC. For the apoptosis assay we used DAPI (Molecular probes) and AnnexinV-APC (BD Biosciences). To asses late stages of apoptosis the APO-BrdU TUNEL Assay Kit from Molecular Probes was used according to the manufacturers’ protocol. For the intracellular detection of Caspase-3, cleaved Caspase-3 (both from Cell Signaling Technology) bone marrow cells were fixed and permeabilized using BD Cytofix/Cytoperm Fixation/Permeabilization Solution Kit (BD Biosciences) according to the manufacturers recommendations.
Prior to stimulation, pro-B cells were washed in OptiMEM + 0.5% FCS to remove IL-7 and starved for 4–6 hr in a 37°C humidified incubator at 5 × 106 cells/ml. Cells were resuspended at 107 cells/ml at 37°C and stimulated by the addition 10–25 ng/ml murine IL-7, for the times indicated. Cells were lysed at 5 × 107 cells/ml in 1% NP40, 150 mM NaCl, 20 mM Tris-HCl (pH 7.4), 1 mM EDTA, 10 mM sodium fluoride, 1 mM sodium orthovanadate, 5 mM sodium pyrophosphate, 1 mM PMSF, and 5 μg/ml aprotinin and leupeptin (Boehringer Mannheim) on ice for 20 min. The detergent insoluble fraction was removed by centrifugation. Protein samples (equivalent of 6 × 105–1.2 × 106 cells) were mixed with 4× NuPAGE sample buffer (Invitrogen) and 0.7 M 2-ME, resolved on a gradient NuPAGE gel, and transferred to a PVDF membrane in 20 mM Tris/150 mM glycine/20% methanol. Detection of phosphorylated ERK1/2 and STAT5 was performed according to the manufacturer’s instructions (NEB). For loading controls, membranes were stripped in 6.25 mM TRIS (pH 6.8), 2% SDS, and 100 mM 2-ME (50°C, 25 min), washed in TBST, and re-probed with actin as above. Bone marrow cells were harvested and pre-cultured in HSC Medium (see above) without cytokines for 1–2h and stimulated with the implied cytokines for the time indicated. Subsequently the cells were fixed and permeabilized using Phosflow Lyse/Fix and Perm/Wash buffers (both from BD-Biosciences) according to manufacturer’s instructions. Cells were than stained for surface markers to define the HSC containing population as described above and with antibodies against pERK1/2, p-p38 (both from BD Biosciences), pAKT, pMEK1/2, pStat3, pSmad-2, pSmad-3 or active (cleaved) Caspase-3 (all from Cell Signaling). Proteolytic inhibition of Caspase-3 activity was performed by incubation of bone marrow cells with the irreversible Caspase-3 selective inhibitor z.DEVD.fmk or the general Caspase inhibitor z.VAD.fmk (both from BD Biosciences) at a concentration of 20μM for 60 min prior to stimulation with cytokines. Single cell immunostaining was performed according to the protocol described by Ema et al Nat Protoc 2006.
For serial transplantation 2x106 whole bone marrow cells from 8 to12 weeks old WT and Caspase-3 KO (CD45.2) littermates were injected into lethally irradiated (9,5 Gy) female recipient Bl6-SJL (CD45.1) mice. Engraftment efficiency in recipients has been monitored by donor contribution of CD45.2 positive cells using FACS analysis. Peripheral blood cell counts were obtained by tail vein nicking 3, 8 and 16 weeks post transplantation. Eight weeks post transplantation recipients were used as donors for the next transplantation cycle and for assessment for the contribution of donor derived primitive BM subpopulations. Transplants were discontinued after 4th round of transplantation. For the competitive repopulation assay with bone marrow from young mice we used 5x105 WT or Caspase-3 KO whole bone marrow cells from CD45.2 littermates (8 weeks old) mixed with 5x105 CD45.1 (competitor) WT cells and injected into lethally irradiated CD45.1 recipient mice. Repopulation was assessed by flow cytometry at 16 weeks post transplant for multilineage reconstitution.
Caspase-3 genotyping was done as described previously (Woo et al., 1998). RNA was isolated from FACS sorted bone marrow subpopulations using the PicoPure Kit (Arcturus Bioscience). Pre designed assays for Caspase-3 and Hprt-1 were purchased from Applied Biosystems (assay Ids Mm01195082 and Mm00446968, respectively).
The Student t-test was used to determine the significance of the difference between the means; significance was set at P values less than 0.05.
The authors thank the National Institutes of Health (D.T.S., H.F.), Dr. Mildred Scheel Foundation for Cancer Research (V.J.), the fortüne program of the University of Tübingen (V.J.), and The Burroughs Wellcome Foundation (D.T.S.). The authors also thank David M. Dombkowski, Michael T. Waring, and Hans-Jörg Bühring for cell sorting and assistance with mulitparameter flow cytometry. In addition the authors would like to thank Kathleen M. Leahy for mouse colony maintenance and Bettina Kirchner for technical assistance. We (GK and SK) would like to thank Dr Hiro Nakauchi, Dr Hideo
Ema and Satoshi Yamazaki, Tokyo University, as well as Falk Hertwig, Lund Stem Cell Center, for their generosity, technical advise, and inspiring discussions.
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