Runx1 P1 is the dominant promoter in adult hematopoiesis

Utilizing the P1-GFP::P2-hCD4 reporter mouse model [29], we traced Runx1 expression for both promoters in vivo at a single cell level in adult mice (with flow cytometry gates based on the WT control tissues) (Fig 1A and 1B). We observed substantial heterogeneity of Runx1 expression within adult BM; approximately 55% of all BM cells were P1-GFP positive, almost 21% co-expressing P2-hCD4 (Fig 1B). Red blood cell lysis (using Ammonium-Chloride-Potassium (ACK) buffer) led to the depletion of P1-GFP^- P2-hCD4^- cells; 97% of remaining cells expressed P1-GFP with 20% co-expressing P2-hCD4. No P1-GFP^- P2-hCD4⁺ cells were observed. In the spleen, approximately 70% of cells expressed P1-GFP but only 1% co-expressed P2-hCD4, whereas in the thymus almost 100% of cells expressed P1-GFP, a quarter of which also expressed P2-hCD4. Altogether these results establish, in line with other reports, that P1 is the dominant Runx1 promoter in adult hematopoietic populations and that the activity of P2 is much more restricted [28].

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Fig 1. Runx1 promoter P1 and P2 expression in adult hematopoietic organs.

(A) Schematic diagrams of the Runx1 WT (top) and P1-GFP::P2-hCD4 dual reporter (GFP::hCD4, bottom) alleles. Expression of GFP is directed by Runx1 promoter P1 and a truncated hCD4 reporter is expressed under the control of Runx1 promoter P2. (B) Contour plots of Runx1 P1-GFP and P2-hCD4 expression in unfractionated total (far left panel) and ACK buffer lysed (middle left panel) adult BM, spleen (middle right panel) and thymus (far right panel) in WT (top) and P1-GFP::P2-hCD4/+ (bottom) mice. Representative data of three independent experiments are shown.

https://doi.org/10.1371/journal.pgen.1005814.g001

Taking advantage of our reporter model, we pursued a detailed examination of P1-GFP and P2-hCD4 expression in mature lymphoid and erythro-myeloid populations (Figs 2 and S1).

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Fig 2. Runx1 P1 and P2 expression in mature hematopoietic lineages.

(A, C, E, G, I, K) Contour plots of Runx1 P1-GFP and P2-hCD4 expression in BM erythroid (A), granulocytic/macrophage (C) and B lymphocyte populations (E), spleen B (G) or T (K) lymphocytes and thymocytes (I), as defined in S1 Fig. (B, D, F, H, J, L) Numbers of Runx1 P1^- P2^-, P1⁺ P2^- and P1⁺ P2⁺ cells as a proportion of defined BM erythroid (B), granulocytic/macrophage (D) and B lymphocyte populations (F), spleen B (H) or T (L) lymphocytes and thymocytes (J). Representative data of three independent experiments are shown.

https://doi.org/10.1371/journal.pgen.1005814.g002

Runx1 is expressed in definitive erythroid precursors, where it is involved in the regulation of erythroid gene expression as part of a core transcription factor complex, but is subsequently downregulated in mature erythrocytes [3–5,28,30]. Correspondingly, P1-GFP expression was restricted to 26% of the proerythroblast (CD71^high Ter119^int, ProE), 56% of the basophilic erythroblast (CD71^high Ter119^high FSC^high, EryA) and 4% of the late basophilic/polychromatic erythroblast (CD71^high Ter119^high FSC^low, EryB) fractions whilst being completely absent in the most mature CD71^low Ter119^high FSC^low (orthochromatic erythroblasts, reticulocytes, red blood cells, EryC) compartment (Figs 2A, 2B, and S1A). P2-hCD4 was expressed in less than 1% of Ter119⁺ erythroid cells, being apparently entirely dispensable for adult erythropoiesis. The low level of expression from both Runx1 P1 and P2 promoters, particularly the latter, in WT erythroid lineage cells was confirmed at the RNA level by qPCR (S2A Fig).

By contrast to the restricted expression observed in the erythroid lineage, Runx1 P1-GFP was expressed in almost 100% of mature myeloid CD11b⁺ BM cells (Figs 2C, 2D and S1B). Of these, P2-hCD4 was co-expressed in 12% of Gr1^high granulocytes, 39% of Gr1^-/low monocytic/immature granulocyte cells and 17% of F4/80⁺ macrophages. The decreased P2 activity in the more mature granulocytic/macrophage (GM) fractions suggests a diminished role for RUNX1B as myeloid differentiation progresses. This also appears to be the case for terminal lymphoid differentiation, as P2-hCD4 co-expression with P1-GFP was restricted in the B-cell lineage to 58% of the BM Pre-pro-B, almost half (49%) of the Pro-B and just 8% of the Pre-B progenitors (Figs 2E–2H, S1C and S1D). P1-GFP was expressed in over 90% of BM B cell progenitors but was reduced to approximately 80% of mature BM and spleen B cells. Finally, thymic T cells were highly enriched in the P1-GFP⁺ P2-hCD4^- fraction but P2-hCD4 activity appeared to peak in the CD4 CD8 double negative 2 (DN2) fraction at approximately 61% (Figs 2I, 2J and S1E). Interestingly, the more mature spleen CD4 and CD8 single positive (SP) T cell subsets displayed greater heterogeneity than their thymic counterparts; almost 100% of CD4 SP cells express P1-GFP whilst this is the case for only 20% of CD8 SP cells (Figs 2K, 2L and S1F). Relative quantitation of the Runx1 isoforms’ expression revealed comparatively high P1 and P2 activity in the GM, B and T lineages, peaking in the early thymic T cell CD4 CD8 DN population and provides direct evidence that the P1-GFP::P2-hCD4 reporters faithfully represent WT Runx1 expression throughout adult hematopoiesis (S2A–S2C Fig). Overall, P1 clearly dominates, accounting for over 80% of Runx1 expression in all analyzed lineage positive populations. Nonetheless, strong P2 expression was observed in CD11b⁺ GR1⁺ GM cells, Pre-pro/pro/pre-B cells and CD4 CD8 DN T cells, decreasing substantially in the more mature IgM⁺ B and CD4/CD8⁺ T cells. These results indicate that P1 is the dominant Runx1 promoter in terminally differentiated hematopoietic cells and suggest that downregulation of P2 is required for maturation to occur. We therefore decided to determine whether P2 expression has a greater prominence and significance in immature HSPC subsets.

Upregulation of Runx1 P2 in HSPCs marks a loss of erythroid potential

To examine the relative activities of the two Runx1 promoters in the most immature hematopoietic compartments, we separated the LSK fraction into phenotypic HSC and MPP fractions (Fig 3A). We observed that only the P1 promoter was active in the HSCs and CD48- MPPs (Fig 3C). The upregulation of CD48 expression coincides with the loss of long-term repopulating ability, the LSK CD48⁺ fraction consisting of a mixture of lymphoid and myeloid progenitors with varying multipotentiality. FMS-Like Tyrosine Kinase 3 (FLT3) expression marks a commitment to the GM and lymphoid lineages at the expense of Mk/Ery specification [31,32]. Increased GM/lymphoid lineage commitment appears to coincide with increased P2 activity, as the majority of cells in the lymphoid-primed multipotent progenitor (LMPP)-enriched FLT3⁺ and the common lymphoid progenitor (CLP, Fig 3B) subsets co-expressed P1-GFP and P2-hCD4 (Fig 3C). Therefore, although P1 is the dominant Runx1 promoter at the onset of adult hematopoiesis, our results suggest that P2 expression imparts or at least reflects distinct lineage commitment decisions in these immature hematopoietic compartments. Consistent with this theory, Runx1 P1 activity, as measured by quantitative RT PCR in WT BM HSPCs, peaked in the WT HSCs and decreased by approximately 50% in the FLT3⁺ MPPs, coinciding with the substantial increase in Runx1 P2 expression (S3 Fig). The overall result is that total Runx1 expression decreases only modestly in MPPs compared to HSCs but the relative contribution by P1 compared to P2 decreases substantially.

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Fig 3. Upregulation of Runx1 P2 in HSPCs marks a loss of erythroid potential.

(A–B) Contour plots showing immature adult BM Lineage (Lin) negative hematopoietic progenitors. (A) The LSK fraction can be separated into distinct HSCs, CD48^- MPPs and more mature CD48⁺ MPPs on the basis of CD150, CD48 and FLT3 expression. (B) The CLP is characterized as Lin^- CD127⁺ cKit^low Sca1^low. (C) Representative FACS plots of P1-GFP/P2-hCD4 expression in HSCs, CD48^- MPPs, CD48⁺ FLT3^- (LSK48F^-) MPPs, CD48⁺ FLT3⁺ (LSK48F⁺) MPPs and CLPs. (D) Numbers of B220⁺ CD19⁺ B cells and CD4⁺ CD8⁺ DP T cells produced following co-culture of WT, P1⁺ P2^- and P1⁺ P2⁺ LSK48F^- MPPs (top) and LSK48F⁺ MPPs (bottom) with OP9 (B cells) or OP9-DL1 (T cells) for 21 days. (n = 4). (E) CFU-C activity of WT, P1⁺ P2^- and P1⁺ P2⁺ LSK48F^- MPPs (top) and LSK48F⁺ MPPs (bottom) following culture in pro-myeloid semi-solid methylcellulose-based medium. (LSK48F^-, n = 7; LSK48F⁺, n = 6.) (F) Photographs of representative LSK48F^- derived methylcellulose colonies. (G) Representative CD11b/Gr1, CD41 and Ter119 FACS plots of OP9 co-cultured LSK48F^- MPPs isolated on day 8. (H) Quantification of CD11b⁺ Gr1⁺ granulocyte/macrophage (GM) cells, CD41⁺ megakaryocytes and Ter119⁺ erythrocytes in the progenitor/OP9 co-culture assays (n = 5).

https://doi.org/10.1371/journal.pgen.1005814.g003

When the differences in biological potential were directly assessed in FLT3⁺ MPPs, P2-hCD4^- and P2-hCD4⁺ subpopulations were capable of relatively similar levels of lymphoid and myeloid differentiation (Fig 3D and 3E bottom), although the increased GM:M CFU-C ratio in the P2-hCD4^- subset may suggest it represents a more immature population than its P2-hCD4⁺ counterpart. However, the difference in lineage output by the FLT3^- subsets was more marked; P2-hCD4^- LSK CD48⁺ FLT3^- (LSK48F^-) MPPs appeared to have reduced T cell output (Fig 3D top) but enhanced multilineage myeloid colony-forming unit potential at the expense of CFU-M output (Fig 3E and 3F). Co-culturing the LSK48F^- progenitors with the OP9 murine stromal cell line in myeloid differentiation media revealed that CD11b⁺/Gr1⁺ GM output was significantly decreased and CD41⁺ megakaryocytic (Mk) cell production was slightly increased in the P2-hCD4^- fraction as a proportion of total cells (Fig 3G and 3H). As a proportion of non-GM (CD11b^-) cells, CD41⁺ Mk cell output was in fact significantly increased in the P2-hCD4⁺ LSK48F^- fraction. Most strikingly, Ter119⁺ erythroid cell output was almost entirely restricted to the P2-hCD4^- fraction. Our phenotypic characterization of BM HSPCs therefore demonstrate that upregulation of Runx1 P2 not only occurs after loss of HSC activity but also coincides with a substantial decrease in erythroid specification.

To determine whether LSK48F^- P2-hCD4^- and P2-hCD4⁺ progenitors arise sequentially or independently in the hematopoietic hierarchy, sorted cells were cultured with pro-myeloid cytokines for up to 18 hours and immunophenotypically characterized (S4A and S4B Fig). Whereas the P2-hCD4⁺ fraction solely produced P2-hCD4⁺ LSK cells, P2-hCD4^- cultures yielded P2-hCD4^- and P2-hCD4⁺ LSK cells (S4B Fig). In addition, LSK48F^- P2-hCD4^- cultures produced more phenotypic erythroid (pre-erythroid colony-forming unit, PreCFUe or erythroid colony-forming unit, CFUe) or bi-potential PreMegE progenitors and fewer GM (Pre- Granulocyte-Macrophage progenitor, PreGM or Granulocyte-Macrophage Progenitor, GMP) and megakaryocyte progenitor (MkP) cells compared to LSK48F^- P2-hCD4⁺ cells. Altogether these data demonstrate a hierarchical relationship between an erythroid-biased P2-hCD4^- MPP population and increasingly pro-GM/Mk P2-hCD4⁺ progeny.

Runx1 P2 expression in GM-restricted progenitors enriches for monocyte/macrophage specification

Although Runx1 P2 expression appears to decrease as GM maturation proceeds, its expression in the earliest identified GM-restricted progenitors remained unknown. We found that approximately 80% of PreGMs and 70% of GMPs co-express P1-GFP and P2-hCD4 (Fig 4A and 4B). Moreover, we found P2 expression (as determined by qRT-PCR) to be higher in the PreGM and GMP than other analyzed WT BM HSPC populations but P1 expression was only 50% and 25% of the level observed in HSCs (S3 Fig). High P2 activity therefore appeared to be important for GM lineage commitment and we decided to investigate the functionality of the P2-hCD4⁺ and minority P2-hCD4^- GM progenitor populations. Interestingly, CFU-C activity was significantly higher in the P2-hCD4⁺ fractions, compared to the P2-hCD4^- populations, of both PreGM and GMP populations, reflecting higher CFU-M and CFU-GM frequencies (Fig 4C–4E). In particular, the P2-hCD4^- GMP fraction appeared to consist of monopotent granulocytic and monocytic/macrophage progenitors rather than bipotential GM progenitors. Liquid culture of the progenitors confirms an apparent bias against macrophage specification as F4/80⁺ cell numbers were significantly diminished in P2-hCD4^- PreGM and GMP cultures, whereas Gr1^high granulocyte output was unaltered (S5A and S5B Fig). Interestingly, production of CD11b^- cKit⁺ FcεR1α⁺ mast cells was also elevated in P2-hCD4^- cultures and more detailed analyses confirmed the absence of P2-hCD4 expression in immunophenotypic mast cell progenitors (MCp, S5A–S5C Fig). The decreased CFU-GM activity of the P2-hCD4^- GM progenitors implies that they reside later in the hematopoietic hierarchy than the P2-hCD4⁺ populations, but in vitro lineage tracing revealed that P2-hCD4^- GMPs gave rise to P2-hCD4⁺ GMPs (S5E Fig). P2-hCD4^- PreGMs gave rise to P2-hCD4⁺ PreGMs and subsequently to P2-hCD4^- and P2-hCD4⁺ GMPs, the latter dominating (S5D Fig). Therefore even within the GM lineage, differential Runx1 promoter activity appears to play a role in or at least correlate with crucial cell fate decisions.

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Fig 4. Runx1 P2 expression in GM-restricted progenitors enriches for monocyte/macrophage specification.

(A) Contour plots of adult BM Lin^- Sca1^- cKit⁺ (LK) GM progenitors. The GMP and PreGM progenitors can be distinguished on the basis of CD41, CD16/32, SLAMF1 (CD150) and Endoglin expression. (B) Representative FACS plots of P1-GFP/P2-hCD4 expression in PreGM and GMP cells. (C–E) CFU-C activity of WT, P1⁺ P2^- and P1⁺ P2⁺ PreGMs and GMPs following culture in pro-myeloid semi-solid methylcellulose-based medium. (C) Total CFU-C numbers (%). (D-E) Granulocyte (CFU-G), macrophage (CFU-M) and granulocyte/macrophage (CFU-GM) colony forming unit numbers per 100 plated PreGMs (D) or 500 plated GMPs (E). n = 4 independent experiments.

https://doi.org/10.1371/journal.pgen.1005814.g004

Distinct megakaryocytic and erythroid progenitors can be isolated on the basis of Runx1 P2-hCD4 expression

Erythropoiesis and megakaryopoiesis are highly similar developmental pathways, sharing numerous regulatory factors particularly at the point of lineage specification [33,34]. However, there are key differences and the specificity of a megakaryocyte maturation defect in Runx1-null adult BM implicates RUNX1 as a central player in Mk/Ery lineage determination [14]. Moreover, our observation that P2-hCD4^- and P2-hCD4⁺ MPPs have distinct Mk/Ery potential led us to investigate Runx1 promoter activity in Mk/Ery-restricted progenitors further (Fig 5A). We observed that erythroid restricted PreCFUe and CFUe progenitors expressed solely P1-GFP whereas the MkP was chiefly P1-GFP⁺ P2-hCD4⁺ (Fig 5B). Because mature megakaryocytes are scarce in adult mice, BM-derived megakaryocytes were obtained by culturing purified MkPs in vitro. CD41-expressing megakaryocytes expressed P2-hCD4 and a large fraction (60%) co-expressed P1-GFP (Fig 5C). Whilst lineage-restricted megakaryocytic and erythroid progenitors were highly homogeneous, the PreMegE fraction, which generates the MkP and PreCFUe populations, was markedly more heterogeneous; approximately 75% express solely P1-GFP whereas the remaining 25% were P1-GFP⁺ P2-hCD4⁺ (Fig 5B).

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Fig 5. Distinct megakaryocytic and erythroid progenitors can be isolated on the basis of Runx1 P2-hCD4 expression.

(A) Contour plots of adult BM LK Mk/Ery progenitors. The PreMegE, MkP, PreCFUe and CFUe populations can by distinguished on the basis of cKit, CD41, SLAMF1 (CD150), CD16/32 and Endoglin cell surface expression. (B) Representative FACS plots of P1-GFP/P2-hCD4 expression in PreMegE, MkP, PreCFUe and CFUe subsets. (n = 8). (C) P1-GFP/P2-hCD4 expression in BM MkP-derived cultured CD41⁺ megakaryocytes. (n = 4). (D–H) CFU-C activity of WT, P1⁺ P2^- and P1⁺ P2⁺ PreMegEs. Cells were cultured either in pro-myeloid semi-solid methylcellulose-based medium (D) or in pro-megakaryocytic collagen-based MegaCultTM medium (F). Photographs of representative PreMegE-derived methylcellulose (E) and MegaCultTM (G) colonies. (n = 5) (H) Numbers of megakaryocytes per MegaCultTM CFU-Mk colony from 3 independent experiments (mean ± SD, Mann-Whitney U test). (I) Contour plots of OP9 co-cultured PreMegE cells isolated on day 7 and stained with CD41 and Ter119 antibodies. (J) Quantification of CD41⁺ megakaryocytes and Ter119⁺ erythrocytes in the PreMegE/OP9 co-culture assays. (n = 5). (K) Representative FACS plots of PreMegE cells following short-term (12 hours) culture in pro-myeloid liquid medium. Top: CD150/CD41 expression of LK CD16/32^- progenitor cells. Middle: Endoglin/CD150 expression of LK CD41 negative CD16/32 negative progenitors. Bottom: P1-GFP/P2-hCD4 expression of immunophenotypic PreMegE (LK CD41- CD16/32- Endoglin- CD150⁺) cells (n = 3).

https://doi.org/10.1371/journal.pgen.1005814.g005

When compared to the relative homogeneity of the monopotent MkP, PreCFUe and CFUe populations, the heterogeneity of the PreMegE led us to consider the possibility of two functionally distinct and prospectively isolatable PreMegE subsets. We subsequently observed that erythroid CFU-C activity (CFUes and erythroid blast-forming units (BFUes)) was significantly enriched in the P2-hCD4^- PreMegE fraction compared to the P2-hCD4⁺ population (Fig 5D and 5E). By comparison, megakaryocyte CFU-C potential was highly enriched in the P2-hCD4⁺ fraction (Fig 5D–5H). In the MkP population, the P2-hCD4⁺ fraction possessed similar megakaryocytic CFU-C activity to its WT counterpart (S6A Fig), suggesting Runx1 haploinsufficiency did not significantly impair megakaryocyte colony formation. In addition to being more numerous, CFU-Mks derived from P2-hCD4⁺ PreMegEs were also larger than those derived from the P2-hCD4^- fraction, the median number of cells per colony being doubled (Fig 5E, 5G and 5H). To determine whether this was a result of increased proliferation in the P2-hCD4⁺ PreMegE fraction, we analyzed their cell cycle status by measuring 5’ethynyl-2’-deoxyuridine (EdU) incorporation and DNA content (S6B Fig). More P2-hCD4⁺ PreMegE cells were in the EdU⁺ DNA Synthesis (S) phase compared to their P2-hCD4^- counterparts, suggesting P2-driven RUNX1B expression may confer a proliferative advantage on PreMegE cells.

The distinct megakaryocytic and erythroid potential of the two PreMegE fractions was further confirmed following co-culture with OP9 cells. After 7 days, P2-hCD4⁺ PreMegE cultures contained significantly more CD41⁺ megakaryocytes and significantly fewer Ter119⁺ erythroid cells than the P2-hCD4^- PreMegE cultures (Fig 5I and 5J). In addition, we performed clonal analyses by plating single P2-hCD4^- and P2-hCD4⁺ PreMegEs with OP9 (S6C Fig). This demonstrated that although the two fractions had similar clonal output (28% and 24% positive wells respectively) the P2-hCD4⁺ fraction contained more bi-potent megakaryocytic/erythroid progenitors (42.9% versus 25% “Mk + Ery”) and more monopotent megakaryocyte-producing progenitors (31.4% versus 12.5% “Mk only”) than the P2-hCD4^- fraction. The P2-hCD4^- fraction was highly enriched for monopotent erythroid-producing progenitors (62.5% “Ery only” compared to 25.7% in the P2-hCD4⁺ co-cultures). Therefore, as in the immature LSK HSPC compartment, upregulation of P2 expression in PreMegEs appeared to coincide with a loss of erythroid and an enrichment of megakaryocytic specification.

To decipher their relative positions in the hematopoietic hierarchy, P2-hCD4^- and P2-hCD4⁺ PreMegE cells were cultured for up to 12 hours and analyzed (Fig 5K). We observed that P2-hCD4⁺ PreMegEs made a more rapid transition to an MkP immunophenotype than P2-hCD4^- cells. In addition P2-hCD4^- PreMegEs gave rise to both P2-hCD4^- and P2-hCD4⁺ fractions in vitro, whereas the P2-hCD4⁺ fraction did not appear to downregulate P2-hCD4. Taken together, these data suggest the P2-hCD4^- PreMegE can be placed earlier in the hematopoietic hierarchy, giving rise to the P2-hCD4⁺ PreMegE. Interestingly, we also observed that cultured P2-hCD4⁺ LSK48F^- cells produced only P2-hCD4⁺ immunophenotypic PreMegEs (S4B Fig). The differences in megakaryocytic and erythroid lineage potential in the P2-hCD4 negative and positive LSK48F^- fractions may therefore be due to the preferential downstream specification of distinct PreMegE subpopulations.

Global gene expression analysis of Runx1 P2-hCD4 positive and negative PreMegEs enables the identification of WT equivalents

To explore the distinct gene regulatory mechanisms involved in the bifurcation of the Mk/Ery pathway, and to identify candidate genes which may serve as markers to isolate the progenitors in WT BM, we performed global gene expression analysis by RNA-Seq (Fig 6A and S4 Table). The expression patterns of WT, P2-hCD4⁺ (P2+) and P2-hCD4^- (P2-) PreMegE samples were clearly separated based on principal component analysis, with WT cells clustering between the P2+ and P2- samples (S7A Fig). When directly comparing the P2+ and P2- populations, 4876 genes were found to be at least 2-fold differentially expressed (false discovery rate <0.05), 2681 being upregulated in P2+ and 2195 in P2- PreMegEs (Fig 6A and S4 Table). Gene Set Enrichment Analyses (GSEA) revealed a significant correlation between P2-hCD4 expression and activation of the Thrombopoietin (TPO) and Integrin pathways, both of which are crucial for megakaryopoiesis (Figs 6B and S7B–S7D) [35–37]. In line with the observed increased proliferative capacity of P2-hCD4⁺ PreMegEs, cell cycle regulators were also enriched in this population. Ingenuity pathway analysis (IPA) identified cell migration and blood cell recruitment as highly enriched functions and integrin signaling as the most significant activated pathway in P2-hCD4⁺ PreMegEs (S7E and S7G Fig). By contrast, functions and pathways associated with cell death and cell cycle inhibition were highly enriched in P2-hCD4^- PreMegEs (S7F and S7H Fig).

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Fig 6. Global gene expression analysis of Runx1 P2-hCD4+ and P2-hCD4- PreMegEs enables identification of WT equivalents.

(A) Heat map depiction of genes at least 2-fold differentially expressed between P1⁺ P2^- (P2^-) and P1⁺ P2⁺ (P2⁺) PreMegE samples, as determined by RNA Seq. Genes in red are upregulated and genes in blue are downregulated. (B) GSEA showing significantly enriched signaling pathways in the gene set upregulated in P2⁺ PreMegEs relative to P2^- PreMegEs. (C) Reads per kilobase per million mapped reads (RPKM) values of selected Mk/Ery- associated genes. (n = 3). (D) Quantitative PCR (qPCR) validation of expression of genes depicted in D. (n = 5). (E) Representative FACS plots of CD34 expression in P2⁺ and P2^- PreMegEs (top) and unsorted, purified CD34⁺ and purified CD34- WT PreMegEs (bottom). (P1-GFP::P2-hCD4 mice n = 3; WT mice n = 5). (F) CFU-C activity of CD34⁺ and CD34^- PreMegEs following culture in pro-myeloid semi-solid methylcellulose-based medium. (n = 5). (G) Contour plots of OP9 co-cultured CD34⁺ and CD34^- PreMegE cells isolated on day 7 and stained with CD41 and Ter119 antibodies. (H) Quantification of CD41⁺ megakaryocytes and Ter119⁺ erythrocytes in the PreMegE/OP9 co-culture assays (n = 4).

https://doi.org/10.1371/journal.pgen.1005814.g006

To further validate the distinct “pro-megakaryocytic” and “pro-erythroid” phenotypes of each PreMegE population, we screened, and validated by qPCR, the RNA Seq data for the expression of known Mk/Ery regulators and markers (Fig 6C and 6D). Early erythroid-associated factors, including Kruppel-like factor 1 (Klf1) and the Erythropoietin receptor (Epor) were significantly upregulated in P2-hCD4^- PreMegEs whereas numerous megakaryocyte-specific markers (Integrin alpha 2b (Itga2b or Cd41), Integrin beta 3 (Itgb3 or Cd61), Myeloproliferative Leukemia Virus Oncogene (Mpl) and Platelet Factor 4 (Pf4)) were enriched in the P2-hCD4⁺ fraction. Interestingly, the transcription factors GATA binding protein 1 (Gata1) and Growth Factor Independent 1B (Gfi1b) were upregulated in P2-hCD4^- PreMegEs. Both factors are crucial for the normal development of both megakaryocytic and erythroid lineages: deletion of either Gata1 or Gfi1b results in an early block in erythropoiesis at the PreCFUe stage whereas megakaryocytic maturation is impaired resulting in the accumulation of undifferentiated megakaryoblasts [38–41]. It would therefore appear that high Gfi1b and/or Gata1 expression promote erythroid specification whereas lower levels would favor megakaryocytic commitment, but ultimately an increase of both would be required for megakaryocytic maturation and thrombopoiesis. It is therefore highly likely that differential expression of Gata1 and Gfi1b at the PreMegE stage plays a role in Mk/Ery lineage determination and their differential expression may be driven by P2-driven RUNX1B.

In order to distinguish “pro-megakaryocytic” and “pro-erythroid” PreMegE subsets in WT mice by alternative means to our reporter line, we screened the list of differentially expressed genes in P2-hCD4⁺ and P2-hCD4^- PreMegEs for cell surface markers with commercially available antibodies validated for use in flow cytometry (S8A Fig). The majority of selected markers had low RPKM values, with the exception of Itgb3 (Cd61) and Cd34 (Figs 6C–6E and S8B). However, CD61 protein expression was not detected on P2-hCD4⁺ and P2-hCD4^- PreMegE cells by flow cytometry (S8C and S8D Fig). By contrast, CD34 expression was approximately 2-fold higher in P2-hCD4⁺ PreMegEs compared to P2-hCD4^- cells, both in terms of numbers of positive cells and median fluorescence intensity (MFI; Figs 6E and S8D). WT CD34⁺ and CD34^- PreMegEs were therefore FACS sorted to >95% purity (Fig 6E) and their lineage output and Runx1 isoform expression elucidated. Importantly, Runx1 P2 expression was substantially higher in the CD34⁺ PreMegEs compared to the CD34^- fraction (S3 Fig). Interestingly, P1 activity was also increased in the CD34⁺ cells, resulting in a 20% increase in total Runx1 expression. It is therefore unclear how important expression of the RUNX1B isoform is for the promotion of megakaryopoiesis compared to enhanced RUNX1 expression overall. Indeed, P2 transcripts were even more highly expressed in WT MkPs, contributing to the highest levels of total Runx1 in all analyzed HSPCs (S3C Fig). However, P1 expression was in fact decreased in MkPs compared to PreMegEs, offering additional evidence in favor of a specific pro-megakaryopoiesis role for the P2-specified RUNX1B protein.

Akin to P2-hCD4⁺ PreMegEs, WT CD34⁺ PreMegEs had enhanced CFU-Mk and diminished BFUe activity compared to CD34^- cells (Fig 6F). Myeloid co-culture with OP9 stromal cells confirmed these phenotypes, as CD34⁺ PreMegEs produced substantially more CD41⁺ megakaryocytes and fewer Ter119⁺ erythroid cells (Fig 6G and 6H). Single-cell OP9 co-culture revealed the CD34⁺ PreMegE compartment was highly enriched for monopotent “Mk only” progenitors (79.2% versus 41.7%) and bipotent “Mk + Ery” progenitors (20.8% versus 8.3%) compared to the CD34^- fraction (S8E Fig). By contrast, monopotent “Ery-only” progenitors accounted for 50% of the CD34^- PreMegE cultures but were apparently absent from the CD34⁺ fraction. We have therefore established the existence of prospectively isolatable “pro-megakaryocytic” CD34⁺ and “pro-erythroid” CD34^- PreMegE cells in WT mice.

Deletion of the dominant RUNX1C isoform does not ablate adult hematopoiesis but results in numerous lineage-specific defects

Having established that the P1-directed RUNX1C isoform is expressed throughout adult hematopoiesis, we decided to determine how its absence would impact the overall homeostasis of the adult blood system. Previously, we utilized the P1-GFP homozygous mouse to investigate the requirement for RUNX1C at the onset of hematopoiesis and found it to be dispensable for hematopoietic commitment [29]. However, this may be due to P2 being the dominant promoter at this stage. We therefore analyzed hematopoietic populations in adult WT, RUNX1C heterozygous (P1-GFP/+) and homozygous knockout (P1-GFP/GFP) mice (Fig 7A). Despite the high expression of P1 in erythroid, myeloid and lymphoid progenitors, we observed no significant perturbation of circulating red or white blood cell numbers upon performing automated cell counts (Fig 7B). Erythroid differentiation appeared to be normal, as peripheral blood hematocrit, hemoglobin concentration and the reticulocyte counts of the RUNX1C null mice were unaltered compared to their WT and heterozygous littermates (S9A–S9C Fig). Similarly, myelo-lymphoid cell fate decisions did not appear to be significantly affected, as the proportions of circulating monocytes, neutrophils and lymphocytes were unaffected (S9D-F). However, a modest but significant decrease in platelet numbers was observed in RUNX1C null mice compared to both the WT and heterozygous animals (Fig 7B). Their plateletcrit was also slightly decreased (albeit not to a significant extent) but the mean platelet volume was unaltered (S9G and S9H Fig). This suggests that, unlike in the conditional total Runx1 null adult mouse model, platelet maturation is not impaired but specification may be hampered.

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Fig 7. Impact of the absence of P1-directed Runx1 expression on adult hematopoiesis.

(A) Top: Schematic diagrams of the Runx1 WT (top) and P1-GFP (bottom) alleles. Expression of GFP is directed by Runx1 promoter P1 but expression of Runx1 from the P2 promoter remains intact. Bottom: Schematic diagram of the experimental design for the investigation of the impact of Runx1 P1 deletion on adult hematopoiesis. Peripheral blood, BM, thymus and spleen samples were collected from adult WT, P1-GFP heterozygous (P1-GFP/+) and homozygous (P1-GFP/GFP) adult mice. All samples were analyzed for mature blood cell surface marker expression. In addition, blood samples were subjected to automated cell counts (Sysmex) and CFU-C assays were performed on unfractionated BM. (B) Peripheral blood cell counts of WT, P1-GFP/+ and P1-GFP/GFP mice as determined by Sysmex automated cell counting. (C) Numbers of CD3e+ T cells, CD11b+ Gr1+ GM cells and B220+ CD19+ B cells as a proportion of total ACK-lysed blood cells from WT, P1-GFP/+ and P1-GFP/GFP mice. (D) CFU-C activity of WT, P1-GFP/+ and P1-GFP/GFP unfractionated ACK-lysed BM following culture in pro-myeloid semi-solid methylcellulose-based medium. (n = 4.) (E) Numbers of erythroid lineage (ProE, EryA, EryB and EryC) cells as a proportion of live unfractionated BM cells. (F) Numbers of T cell lineage populations as a proportion of live unfractionated thymus cells. (G) Numbers of CD4 SP and CD8 SP T cells as a proportion of live unfractionated spleen cells. (H) Ratio of splenic CD4 SP T cells to splenic CD8 SP T cells (n = 4).

https://doi.org/10.1371/journal.pgen.1005814.g007

FACS analysis of circulating blood cells and BM confirmed the presence of equal proportions of CD11b⁺ Gr1⁺ GM lineage and B220⁺ CD19⁺ B lymphoid cells in WT, P1-GFP/+ and P1-GFP/GFP mice (Figs 7C and S10). However, the numbers of CD3ε⁺ T cells were significantly reduced, suggesting that the absence of RUNX1C partially impairs T cell specification. We therefore analyzed the thymic T cell populations in greater detail and found that CD4/8 DN, DP and SP population numbers were not altered in P1-GFP/GFP mice (Fig 7F). However, the ratio of CD4 SP:CD8 SP T cells in the spleen was severely perturbed, as P1-GFP/GFP mice had considerably fewer CD4 SP and more CD8 SP T cells compared to WT littermates (Fig 7G and 7H). This therefore suggests that RUNX1C is dispensable for the DN to DP transition, observed to be blocked in total Runx1 deficient mice [13]. Nonetheless, the RUNX1C knockout recapitulates the defect in CD4 SP and CD8 SP T cell specification observed in total Runx1+/- mice, clearly demonstrating an important role for P1-driven RUNX1 activity in the T cell lineage [4,42,43].

To determine whether the absence of P1-directed RUNX1C expression impacts adult colony-forming HSPC populations, we performed myeloid CFU-C assays on unfractionated BM from WT, P1-GFP/+ and P1-GFP/GFP mice (Fig 7D). GM, MkE and multilineage GEMM colony numbers were unaffected, but RUNX1C null BM cells produced significantly more erythroid CFUe colonies than either the WT or P1-GFP/+ cultures. FACS analysis of unlysed BM revealed a significant expansion of the EryC population in the RUNX1C null mice, a stage which coincides with almost complete silencing of both the Runx1 P1 and P2 promoters (Fig 7E). In combination with the observed mild thrombocytopenia, it appears that the absence of RUNX1C may favor erythroid specification over megakaryopoiesis, a phenotype observed recently in mouse and human HSPCs depleted for total RUNX1 [44]. Overall, P1-directed RUNX1C activity may be dispensable for normal adult hematopoiesis but its absence nonetheless results in defects reminiscent of total RUNX1 deficiency.

Expression of the AML1-ETO9a oncogene preferentially induces Runx1 P2 expression

Increasingly it is becoming apparent that, in addition to a more classically defined tumor suppressor role, WT RUNX1 is required for the promotion of leukemogenesis in certain leukemia subtypes. Notably, AML1-ETO-driven CBF AML appears to be dependent on maintaining WT RUNX1 activity [21,22]. However, although AML1-ETO appears to promote RUNX1 expression, it is unclear whether AML1-ETO oncogene expression promotes the expression of one Runx1 promoter over another [45]. To address this question, we utilized a novel mouse model expressing a Doxycycline-inducible AML1-ETO9a transgene under the control of a Tetracycline Responsive Element (TRE, Fig 8A). The AML1-ETO9a oncogenic transcript is expressed in a majority of t(8;21) AML patients studied and encodes a truncated AML1-ETO protein with enhanced leukemogenic potential [46,47]. We therefore took advantage of our ability to induce AML1-ETO expression in adult mice (by administering Doxycycline in the food for 8 days) and studied the impact on Runx1 isoform expression in vivo by isolating AML1-ETO-expressing (AML1-ETO9a-IRES-GFP⁺) and non-expressing (AML1-ETO9a-IRES-GFP^-) BM HSPCs and quantitating Runx1 expression through qRT-PCR (Fig 8A). This allowed us to study the effect of AML1-ETO expression on WT Runx1 expression as one of the earliest events at the initiation of leukemogenesis.

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Fig 8. Effect of short-term induction of AML1-ETO9a expression on Runx1 expression in BM HSPCs.

(A) Top: Schematic representation of the Rosa26 and Hprt loci in the Doxycycline-inducible AML1-ETO9a GFP mouse model. The reverse tetracycline-controlled transcriptional activator (rtTA) is constitutively expressed under the control of the Rosa promoter. Upon binding to Doxycycline, the rtTA is capable of binding to and activating a tetracycline responsive element (TRE) located in the ubiquitously expressed Hprt locus, resulting in expression of the hemagglutinin (HA)-tagged AML1-ETO9a::IRES-GFP construct also incorporated into this locus. Bottom: Schematic diagram of the experimental design to induce short-term expression of AML1-ETO9a in adult mice by administering Doxycycline in food for 8 days. Bone marrow cells were the harvested and GFP⁺ and GFP^- HSPC populations were isolated by FACS sorting for RNA extraction and gene expression analysis. (B-E) Gene expression analysis by qPCR of AML1-ETO9a (B), total Runx1 (C), Runx1 P1 (D) and Runx1 P2 (E). (F) Ratio of Runx1 P1:P2 expression (n = 3).

https://doi.org/10.1371/journal.pgen.1005814.g008

Firstly, we confirmed the presence and absence of AML1-ETO9a expression in BM GFP⁺ and GFP^- HSPCs respectively (Fig 8B). We chose to analyze LSK, PreGM and GMP cells as the immature HSPC and GM-lineage progenitors contain the leukemia propagating cell fraction in numerous AML patient samples and in a previously described AML1-ETO mouse model [48,49]. Whilst in the PreGM, and GMP Runx1 P1 expression was unperturbed by the expression of AML1-ETO, it was in fact decreased by approximately 40% in LSK GFP⁺ cells compared to GFP^- (Fig 8D). In all three HSPC populations, however, the presence of AML1-ETO resulted in an upregulation of Runx1 P2 expression (Fig 8E), albeit not to a significant extent in PreGM cells. This resulted in an increase in total Runx1 expression in the GMP fraction (Fig 8C) but also a significant decrease in the P1:P2 ratio in all three populations, particularly in the LSK compartment, a phenotype associated with enhanced CFU-C activity, particularly in the GM lineage (Fig 8F).

Mice

P1-GFP::P2-hCD4 and P1-GFP mice have previously been described [29]. The AML1-ETO9a-IRES-GFP/rtTA mice were generated as follows: HA-tagged AML1-ETO9a cDNAs (provided by the Zhang laboratory [47]) were subcloned into a tet-ON vector in front of an IRES-GFP as described [69]. Ainv18 ES cells [69] (which constitutively express the rtTA under the control of the Rosa26 promoter) were then transfected with this tetracycline-inducible AML1-ETO9a construct by electroporation and stably transfected clones were selected with G418 (0.5mg/ml, Life Technologies) for 10–14 days. Chimeric mice were then generated by injecting AML1-ETO9a-IRES-GFP/rtTA ES cells into C57BL6J blastocysts. To induce AML1-ETO9a-IRES-GFP expression, 12 week-old mice were fed irradiated diet supplemented with 545mg/kg Doxycycline (ssniff Spezialdiäten GmbH) for 8 days prior to humane culling and tissue collection. All animal work was performed under regulations governed by UK Home Office Legislation under the Animals (Scientific Procedures) Act 1986. Details of animal husbandry and tissue collection are listed in S1 File.

Flow cytometry analysis and cell sorting

Dead cells were excluded using either 0.5μg/ml 7-Aminoactinomycin D (7-AAD, eBioscience) or 1μg/ml Hoechst 33258 (Life Technologies). Biotinylated antibody staining was detected by a secondary incubation step with fluorochrome-conjugated Streptavidin. Prior to flow sorting of HSPCs, bone marrow cells stained with biotinylated anti-lineage antibodies were lineage-depleted using anti-biotin-conjugated magnetic beads (Miltenyi) and then stained with additional antibodies, including conjugated streptavidin. Red blood cell depletion was performed by treatment with ACK lysis buffer (154mM ammonium chloride, 9.99mM potassium bicarbonate, 0.110mM EDTA) for 5 minutes at room temperature, followed by quenching with Phosphate-Buffered Saline (PBS).

Details of flow cytometry antibodies and reagents are listed in S1 Table. Details of flow cytometry antibody combinations used for each analysis or sort are listed in S2 Table.

For cell cycle analysis, in vivo EdU incorporation was performed by injecting 1.125mg EdU dissolved in PBS intraperitoneally into adult mice. After two hours, bone marrow was harvested and stained with hematopoietic stem and progenitor cell surface markers as detailed in S2 Table. Cells were then stained using the Click-iT EdU Alexa Fluor 647 Flow Cytometry Assay Kit (Life Technologies). Total DNA was stained with 1μg/ml FxCycle Violet Stain (Life Technologies).

Cells were analyzed using a LSR-II or LSR-II Fortessa analyzer, a FACSAria-II cell sorter or a FACSAria-III cell sorter (BD).

Automated peripheral blood counts

Tail vein blood (no more than 50μl per mouse) was sampled from 12 week old mice using heparinized end-to-end Micro Pipettes (Vitrex) and analyzed on a Sysmex XT 2000i analyzer, according to the manufacturer’s instructions.

Cell culture

Gene expression

Statistical analysis

Flow cytometry plots display the mean values of each indicated population. Unless otherwise indicated, data were evaluated using an Ordinary 2-way ANOVA and expressed as mean ± standard error of the mean (SEM). P<0.05 was considered statistically significant.

*P<0.05, **P<0.01, ***P<0.001, ****P<0.0001