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a Department of Biochemistry, Dartmouth Medical School, Hanover, New Hampshire, USA;
b (Current address for Dr. de Bruijn) MRC Molecular Haematology Unit, Weatherall Institute of Molecular Medicine, John Radcliffe Hospital, Oxford, United Kingdom
Key Words. Runx1/AML1/CBF
2 • Hematopoietic stem cells • Gene expression • Transcription factor
Marella F.T.R. de Bruijn, Ph.D., MRC Molecular Haematology Unit, Weatherall Institute of Molecular Medicine, John Radcliffe Hospital, Oxford OX3 9DS, United Kingdom. Telephone: 44-1865-222397; Fax: 44-1865-222500; e-mail: marella.debruijn{at}imm.ox.ac.uk
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ABSTRACT |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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We previously generated a Runx1 allele in which two coding exons were replaced with lacZ coding sequences (Runx1lz) [14]. The Runx1-ß-galactosidase fusion protein produced from this modified Runx1lz allele contains the N-terminus of Runx1 through amino acid 242 (64 amino acids C-terminal to the DNA-binding Runt domain). Runx11z is a nonfunctional Runx1 allele [14]. The fusion of ß-galactosidase to the DNA-binding domain of Runx1 could potentially interfere with endogenous core-binding factors by creating a protein that can bind DNA but not activate transcription. The fidelity of Runx1-ß-galactosidase expression is supported by the observation that it correlates with the presence of Runx1 protein and mRNA in Runx1lz/+ embryos, as detected by immunohistochemistry and in situ hybridization [15, 16]. Here, mice heterozygous for this Runx1lz allele were used to isolate and characterize Runx1-expressing cells. We show that in the adult mouse, Runx1 was expressed in all BM hematopoietic stem cells (HSCs), in spleen colony-forming units (CFU-S), and in the majority of progenitor cells generating myeloid, erythroid and mixed colonies in culture. We also characterized Runx1 expression in more differentiated hematopoietic cells by flow cytometry and compared the BM and peripheral blood compositions of normal mice with those of mice heterozygous for two different mutated Runx1 alleles to determine which lineages are affected by reduced Runx1 dosage.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Immunofluorescent Labeling and Flow Cytometry
Nucleated BM, spleen, and thymus cells were counted using a Neubauer hemocytometer (VWR Scientific; Mississauga, Ontario, Canada; http://www.vwrsp.com) and Türk’s solution. Peripheral blood leukocyte counts were performed on whole blood using a Pentra 60 (ABI Diagnostics; Columbia, MD; http://www.abionline.com). Immunofluorescent labeling and flow cytometry were performed as described [14], with the modification that BM, spleen, and peripheral blood erythrocytes were lysed prior to labeling by a 2-minute incubation in ammonium chloride buffer (155 mM NH4Cl, 0.1 mM EDTA, 10 mM Tris-HCl, pH 7.5). All antibodies were obtained from Pharmingen (San Diego, CA; http://www.bdbiosciences.com/pharmingen) and were directed against c-kit (CD117; clone ACK45), Sca-1 (Ly-6A/E; D7), Mac-3 (M4/84), Gr-1 (Ly-6G; RB6-8C5), TER-119 (Ly-76; TER-119), Ly-6C (AL-21), BP-1 (Ly-51; 6C3), IgD (11-2c.2a), IgM (R6-60.2), NK1.1 (Ly-55; PK136), TCRß (H57-597), TCR
(GL3), CD3 (145-2C11), CD4 (RM4-5), CD8 (53-6.7), CD11b (Mac-1; M1/70), CD11c (HL3), CD19 (1D3), CD24 (HSA; M1/69), CD25 (interleukin-2R; PC61), CD31 (PECAM-1; MEC 13.3), CD41 (MWReg30), CD43 (Ly-48; S7), CD44 (Pgp-1; IM7), B220 (CD45R; RA3-6B2), CD62P (RB40.34), and CD71 (C2). Fluorescein-di-ß-D-galactopyranoside (FDG; Molecular Probes; Eugene, OR; http://www.probes.com) was used as a fluorescent substrate for ß-galactosidase as described [18]. Briefly, cells were suspended in phosphate-buffered saline (PBS) supplemented with 10% fetal calf serum (FCS) and penicillin/streptomycin (P/S) and incubated for 1 minute precisely with an equal volume of FDG (2 mM stock in water) in a 37°C water bath. FDG loading was stopped by adding excess ice-cold PBS (with 10% FCS and P/S), and cells were kept on ice for 1 hour to recover prior to antibody labeling and throughout the entire labeling procedure. To determine the time span in which the fluorescent form of FDG could be detected, we measured its fluorescent product from 1 to 6 hours after loading at 1 hour intervals by flow cytometer. No changes in FDG signal were detected in this time span (data not shown), and cells were always analyzed within 2 to 3 hours of FDG loading. Wild-type hematopoietic cells were also loaded with FDG and served as negative controls for FDG signal. For antibody staining, appropriate isotype controls were used, as described previously [18]. A fluorescence-activated cell sorter, FACStar Plus (BD Biosciences; San Jose, CA; http://www.bdbiosciences.com), was used for cell sorting, and a CyAn (Dako-Cytomation; Carpinteria, CA; http://www.dakocytomation.com) and FACS Calibur (BD Biosciences) were used for analysis. Propidium iodide, Hoechst 33258, or TO-PRO3 (Molecular Probes) was used to exclude dead cells from analysis and cell sorting.
Analysis of Hematopoietic Activity
Long-term multilineage repopulation, CFU-S, and culture CFU (CFU-C) activity were determined as described [14, 18, 19] with minor modifications. BM cell populations were assayed for HSC activity by intravenous transfer into irradiated (900 rad, split dose) 3- to 6-month-old (129S3/SvImJ x C57BL/6J) F1 recipients. Recipients were analyzed for donor cell contribution at 1 month and 4–6 months post-transplantation by polymerase chain reaction of peripheral blood genomic DNA using primers for lacZ and myogenin sequences as described [18]. Multilineage engraftment was determined by flow cytometric analysis of BM cells as described [18]. For analysis of CFU-S11 activity, irradiated recipients (1,000 rad, split dose) were injected intravenously with sorted BM cells. Control mice were not injected. Spleens were harvested 11 days after transplantation and immersed in Tellyesniczky’s solution (63% EtOH, 5% glacial HOAc, 2% formaldehyde in water). Colonies were counted by visual inspection. For analysis of CFU-C activity, sorted FDG-loaded BM cells and unsorted (not FDG loaded) BM cells were plated in 35-mm suspension culture dishes at 2 x 104 cells/dish in 1% methylcellulose in Iscove’s medium (MethoCult 3231; StemCell Technologies; Vancouver, Canada; http://www.stemcell.com), supplemented with 50 ng/ml stem cell factor, 20 ng/ml interleukin (IL)-3, 25 ng/ml IL-6, 5 ng/ml GM-CSF, 2 ng/ml G-CSF, and 2 U/ml erythropoietin (all cytokines from R&D Systems; Minneapolis, MN; http://www.rndsystems.com). Cultures were grown at 37°C in 5% CO2, and hematopoietic colonies were counted after 11 days using an inverted microscope.
Western Blot Analysis
Single-cell suspensions were prepared from whole thymuses, lysed, and the nuclei collected and extracted in high salt buffer (20 mM HEPES, pH 7.5; 25% glycerol; 0.42 M NaCl; 3 mN MgCl2; 0.2 mM EDTA; 1 mM dithiothreitol; protease inhibitor cocktail [Sigma; St. Louis, MO; http://www.sigmaaldrich.com]). Nuclear proteins were separated on 12% SDS PAGE gels and transferred to nitrocellulose. Runx1 proteins were detected with mouse monoclonal antibody 3.2.5 and enhanced chemiluminescence reagents (ECL; Amersham; Piscataway, NJ; http://www.apbiotech.com).
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RESULTS |
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) using a monoclonal antibody that recognizes the Runt domain. We found that the Runx1-ß-galactosidase fusion protein did not accumulate to higher levels than the endogenous Runx1 protein.
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and 2B; Table 1). Runx1-expressing (Runx1+) and nonexpressing (Runx1-) cells, isolated from total BM and from the c-kit+ and Lin-c-kit+ BM fractions, were assayed for the presence of HSCs. Upon intravenous transfer of 105 sorted cells into irradiated adult recipients, we found long-term multilineage high-level hematopoietic reconstitution, indicative of HSC activity, only in the Runx1+ populations (Table 1). The extent of the hematopoietic reconstitution was similar to that observed after transplantation of 105 unsorted BM cells (7 of 10 recipients reconstituted, 45%-73% donor-derived BM cells; not shown). No detectable donor cell reconstitution was observed after transfer of Runx1- cells. Thus, virtually all long-term repopulating HSCs express Runx1, beginning with their generation in ontogeny [18] and extending into adult life. We also determined that all progenitor cells capable of forming spleen colonies (CFU-S11) expressed Runx1 (Table 1). In addition, in vitro clonogenic progenitors such as CFU-granulocyte-erythrocyte-monocyte-megakaryocyte (CFU-GEMM), CFU-granulocyte-macrophage (GM), and BFU-E, were found almost exclusively among Runx1+ cells. The small number of CFU-C in the Runx1- fraction may be due to contamination with Runx1+ cells, or may reflect the presence of some CFU-C (sort purity was 98%).
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), while no significant decrease was observed in the spleen. In peripheral blood, no significant decrease was found either (Table 2), although we observed that four of seven mice analyzed had leukocyte counts between 1.1 and 1.8 x 106/ml, low counts that were not observed among the six wild-type mice analyzed.
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). In addition, the decrease in spleen cell count observed in the Runx1rd/+ mice versus wild-type mice was statistically significant. The difference in peripheral blood leukocyte counts between the Runx1rd/+ mice and wild-type mice was, like the difference between Runx1lz/+ and wild-type counts, not significant. However, we did find low leukocyte counts (1.1 and 1.6 x 106 nucleated cells/ml) in two of five Runx1rd/+ mice analyzed.
Small changes were seen in the cellular composition of BM and peripheral blood from Runx1lz/+ and Runx1rd/+ mice compared with BM and blood from wild-type mice (Tables 3 and 4). In BM there was a small but significant increase in the proportions of monocyte and granulocyte precursors (Mac-1 and Gr-1 cells), while for most other populations there were decreases in frequency (Table 3). In addition, we tested the effect of Runx1 gene dosage on early progenitor cells in functional assays. Similar frequencies of CFU-S, CFU-GEMM, CFU-GM, and BFU-E were obtained from both wild-type and Runx1lz/+ BM (Table 5). The adult stem cell compartment was not examined.
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). This is consistent with the reported broad expression of Runx1 mRNA and CBF family proteins in BM myeloid cells [10, 20] and the high frequency of Runx1+ cells detected among Mac-1+ and Gr-1+ cells in BM and peripheral blood (Tables 3 and 4). Notably, the level of Runx1 expression in BM granulocytes was approximately 4.5-fold lower than in monocytes (Fig. 2C). Runx1+ cells were also found among mature Mac3+ macrophages and CD11c+ dendritic cells in BM, albeit at lower frequencies (on average 43%-58%; Table 3). In the megakaryocytic lineage, on average, 50% of BM CD62P+ megakaryocytes (Table 3) and 59% ± 12% of peripheral blood CD41+ platelets (n = 4; not shown) expressed Runx1. CBFß expression has also been reported in BM myeloid cells from CFU-GM through fully differentiated cells [13], supporting a role for Runx1-CBFß throughout myeloid differentiation.
Runx1 Expression Decreases Upon Erythroid Maturation
Almost all BFU-E expressed Runx1 (Table 1), yet erythroid cells in BM are reported to express little or no RUNX1 protein and mRNA [10, 11] or other proteins of the CBF family [13, 20]. Indeed, only 26% ± 3% of TER-119+ cells were found to express Runx1 (Table 3). Examination of Runx1 expression during erythroid maturation showed that while, on average, 64% of proerythroblasts still expressed Runx1, this percentage decreased upon maturation to only 16% ± 4% of normoblasts (Fig. 2D, Table 6); the levels of Runx1 expression in individual cells also decreased (Fig. 2D). A similar decrease in CBFß expression was reported [13]. Thus, Runx1-CBFß could be involved in the initial activation of erythroid-specific genes at the CFU-GEMM or BFU-E stages [21], but is probably not required for terminal erythroid maturation [11, 22].
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T-cell development in the thymus is characterized by the maturation of CD4 and CD8 single-positive (SP) cells from a lymphoid precursor population. Hayashi et al. [9] showed an effect of decreased Runx1 dosage in thymocyte maturation characterized by reduced numbers of CD4 and CD8 SP T cells and a relative increase in the percentage of CD4/CD8 double-positive (DP) T cells. In accordance with this, we observed similar changes in CD4/CD8 populations in thymocytes and reduced numbers of CD4 and CD8 SP cells in BM and peripheral blood from Runx1lz/+ mice (Tables 3, 4, and 6). We examined Runx1 expression within the distinct CD4/CD8 thymocyte populations. On average, 44% ± 11% of double negative (DN) cells expressed Runx1 (Fig. 2E; Table 6). The majority of cells in the first three DN stages expressed Runx1 (Fig. 2G), with no observed differences in levels of Runx1 expression between the three subsets (data not shown). In the fourth DN stage, the percentage of Runx1-expressing cells decreased dramatically (Figure 2G). The percentage of Runx1+ cells increases at later stages of T-cell differentiation (Fig. 2E and 2G; Tables 3, 4, and 6).
A role for Runx1 in B lymphopoiesis is suggested by the association of the t(12;21) TEL-Runx1 fusion protein with pediatric acute B-lymphocytic leukemia [1]. We found Runx1 expressed at all stages of differentiation (Fig. 2F and 2H; Table 6) and in peripheral blood B cells (Table 4). During B-cell differentiation in the BM, on average, 48%-68% of the cells are Runx1+ (Fig. 2F and 2H; Table 6), with no obvious differences in the levels of Runx1 expression between immature and more mature B-cell populations (Fig. 2F, and data not shown).
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DISCUSSION |
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The basis for the varying percentages of Runx1+ and Runx1- cells in the lymphoid subpopulations is currently unclear. It may reflect differences in Runx1 expression in cycling versus resting cells. Another possibility is that Runx1 expression in lymphoid cells is stochastic, such that only a certain percentage of cells are at any time Runx1+. Or the various populations we examined may be heterogeneous and consist of specific subpopulations of Runx1+ or Runx1- cells that could be further segregated based on the expression of additional markers. For example, it was recently shown that T helper (Th) 1 and Th2 cells differ in Runx1 expression [24]. Since CBFß is expressed uniformly throughout T-cell development [13], it will be of interest to examine whether Runx1- cells express Runx2 and/or Runx3, the other two core-binding factor family members for which important roles in T-cell differentiation have been reported [12, 23, 30]. The highest proportion of Runx1- cells in the lymphoid lineages was observed in the last DN stage. It is possible that this may reflect the recently reported function of Runx1 in CD4 repression [12, 23], in that the decrease in Runx1 expression might allow the cells to proceed to the CD4+CD8+ stage. However, substantiation of this notion awaits more detailed studies.
We showed by Western blot analysis that the Runx1-ß-galactosidase fusion protein is expressed at similar or slightly lower steady-state levels than endogenous Runx1 protein. It remains possible that the Runx1-ß-galactosidase protein is degraded at a different rate than endogenous Runx1. Human RUNX1 is degraded via the ubiquitin-proteasome pathway, and heterodimerization of RUNX1 with CBFß, through the Runt domain, protects RUNX1 from degradation [31]. At least five of the lysine residues targeted by ubiquitin-mediated degradation are conserved in mouse Runx1 [31] and are maintained in Runx1-ß-galactosidase. Furthermore, Runx1-ß-galactosidase still contains the Runt domain [14], and CBFß was reported to be expressed in all hematopoietic cell types in which we detected Runx1 [13]. Thus, two important aspects influencing Runx1 protein stability are still available to the fusion protein. Uncertainties about protein turnover would apply to any of the currently available methodologies used to study gene expression with fluorescent proteins. For example, expression of ß-galactosidase or green fluorescent protein from the Runx1 locus via an internal ribosome entry site rather than as a fusion protein would bypass the intricate translational control to which Runx1 mRNAs are normally subjected [32], and the stability of the marker protein would not necessarily mimic that of endogenous Runx1. Comparison of enzymatic or fluorescent activity with Runx1 protein levels is also difficult due to differences in the sensitivity of detection. The approach we have taken would overestimate the population of Runx1+ cells if the fusion protein turns over less rapidly than endogenous Runx1. The observation that all HSCs and virtually all committed progenitors fall into the Runx1+ population suggests it is unlikely that we are underestimating the percentage of cells in which Runx1 is expressed. The heterogeneity in levels of Runx1 expression detected by flow cytometry in total BM Runx1+ cells could not be unambiguously related to one or more specific cell types or differentiation stage(s). Instead, the variation was seen in most Runx1+ cell populations.
BM and thymus of Runx1lz/+ mice consistently showed a decreased cellularity. In addition, small but significant changes in the frequencies of the T-cell and monocyte/granulocyte lineages were observed. Decreased cellularity of hematopoietic tissues and changes in their cellular composition were also seen in Runx1rd/+ mice, which are heterozygous for the Runx1rd allele from which a DNA-binding defective, unstable, or no protein would be produced (Tables 3 and 4), indicating that the effects are not specific to the Runx1lz allele. Similar effects on the frequencies of cells in the T-cell or monocyte/granulocyte lineages were found in Runx1lz/+ and Runx1rd/+ mice. In the T-cell lineage, reduced frequencies of mature SP CD4 and CD8 T cells were observed, as reported previously [9]. In contrast, in the myeloid lineage, frequencies of monocytes (BM) and granulocytic cells (BM and peripheral blood) were elevated. As the frequency of Runx1lz/+ CFU-GM was comparable to wild type, the relative increase in myeloid cells cannot be explained by an increase in progenitor cells. Indeed, an increase in frequency was observed only in the terminal stages of BM monocytic (CD31-Ly-6Chi) and granulocytic (CD31-Ly-6Cmed) maturation, but not in the stage before (CD31+Ly-6C+; Table 6 and Fig. 2C). Elucidation of the mechanism(s) behind the observed perturbations in hematopoiesis awaits future studies. The expression of Runx1 in functional HSCs and its continued expression in adult myeloid and lymphoid lineage cells support a role for Runx1 in HSC biology and terminal differentiation of the myeloid and lymphoid lineages. Studies using Runx1 conditional knock-out mice will help to determine at precisely what stage(s) Runx1 is required, and to define whether Runx1 acts as a transcriptional repressor and/or activator in those cells.
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ACKNOWLEDGMENT |
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REFERENCES |
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CD71hi TER-119+