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The Journal of Immunology, 2006, 176: 5267-5275.
Copyright © 2006 by The American Association of Immunologists

Notch Signaling Requires GATA-2 to Inhibit Myelopoiesis from Embryonic Stem Cells and Primary Hemopoietic Progenitors1

Renée F. de Pooter*, Thomas M. Schmitt*, José Luis de la Pompa{dagger}, Yuko Fujiwara{ddagger}, Stuart H. Orkin{ddagger} and Juan Carlos Zúñiga-Pflücker2,*

* Department of Immunology, University of Toronto, Sunnybrook and Women’s Research Institute, Toronto, Ontario, Canada; {dagger} Departamento de Immunología y Oncología, Centro Nacional de Biotecnología-Consejo Superior de Investigaciones Científicas, Universidad Autónoma de Madrid, Madrid, Spain; and {ddagger} Division of Hematology/Oncology, Children’s Hospital and Dana-Farber Cancer Institute, Harvard Medical School, and Howard Hughes Medical Institute, Boston MA 02115


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The bone marrow and thymus, although both hemopoietic environments, induce very distinct differentiation outcomes. The former supports hemopoietic stem cell self-renewal and multiple hemopoietic lineages, while the latter supports T lymphopoiesis almost exclusively. This distinction suggests that the thymic environment acts to restrict the hemopoietic fates available to thymic immigrants. In this study, we demonstrate that the addition of the Notch ligand Delta-like-1 (Dll-1) to an in vitro system that otherwise supports myelopoiesis, greatly reduces the myelopoietic potential of stem cells or uncommitted progenitors. In contrast, committed myeloid progenitors mature regardless of the presence of Dll-1. The block in myelopoiesis is the direct result of Notch signaling within the hemopoietic progenitor, and Dll-1-induced signals cause a rapid increase in the expression of the zinc finger transcription factor GATA-2. Importantly, in the absence of GATA-2, Dll-1-induced signals fail to inhibit commitment to the myeloid fate. Taken together, our results support a role for GATA-2 in allowing Dll-1 to restrict non-T cell lineage differentiation outcomes.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
In vertebrates, definitive hemopoiesis is characterized by multipotent hemopoietic stem cells (HSCs)3 (1), which possess both the potential for self-renewal and the ability to give rise to all the lineages of the hemopoietic system. The choice between self-renewal and differentiation, and between the multiple potential differentiated fates, is influenced by environmental cues. In the adult mammal, the bone marrow environment provides niches for both HSCs (2, 3) and their maturation to the majority of hemopoietic lineages (1). T cells are the exception, and develop primarily in the thymus.

One highly conserved signaling pathway, which controls multiple lineage fate decisions during development, and has been implicated in the ability of the thymus to support T lymphopoiesis, is the Notch pathway (4). In mammals, four Notch receptors (Notch-1, -2, -3, and -4) can be engaged by five ligands: Delta-like (Dll)-1, -3, and -4 and Jagged-1 and -2. Following ligand binding, two proteolytic cleavage events, mediated by a disintegrin and metalloprotease and {gamma}-secretase activity, release the intracellular domain of the Notch receptor (ICN), which translocates to the nucleus. In the canonical Notch-signaling pathway, ICN induces the expression of target genes, such as HES-1, via the DNA-binding protein CBF1/supressor of hairless/Lag-1 (CSL) (4). Alternate, CSL-independent pathways of Notch signaling have been proposed, but their importance in vertebrates is not clear (5) and deficiency in CSL, like deficiency in Notch 1, is embryonic lethal (6, 7, 8).

Within the hemopoietic system, Notch receptors and their ligands are expressed by both hemopoietic progenitors (HPC) and stromal cells (9, 10, 11). Notch receptor-ligand interactions in the thymus have been shown to favor T lymphopoiesis, and discourage commitment to other lineages (12, 13). Not surprisingly, bone marrow-derived stromal lines such as OP9 cells, which have been used to support robust in vitro myelopoiesis, erythropoiesis, and B and NK lymphopoiesis, from embryonic stem cell (ESC)-, fetal liver- and adult bone marrow-derived HPCs (14, 15, 16, 17), do not support T lymphopoiesis. In fact, HPCs are rapidly induced to commit to non-T cell fates in the OP9 stromal cell environment (18). Interestingly, neither OP9 cells nor bone marrow stroma express detectable levels of Dll-1 (19), in sharp contrast to the high levels of Dll-1 expression found on thymic epithelial cells (20, 21). We have recently demonstrated that when OP9 cells are retrovirally transduced to express Dll-1 (OP9-DL1 cells), they support T, but no longer B, lymphopoiesis from HPCs (22, 23).

In this study, we use this system to characterize the effect of Dll-1/Notch interactions on the commitment and maturation of myeloid cells, another hemopoietic lineage that is supported by the bone marrow, but not the thymic, microenvironment.

Although Pui et al. (12) observed no myelopoietic defect from bone marrow retrovirally transduced with ICN, others have reported that exposure to soluble or bound Notch ligands (24, 25, 26), or transduction with constitutively active Notch (27, 28, 29, 30) or its downstream target HES-1 (31), inhibits myelopoiesis from most hemopoietic cell lines and HPCs (32, 33). In particular, others have observed that the Jaggeds do not impede myelopoiesis (34, 35), but Dll-1 does (36, 37). In contrast, two groups found that Notch signaling promotes differentiation by the 32D cell line (38, 39, 40), while a separate report concluded that Notch signals are necessary for the differentiation of dendritic cells, but prevent their maturation (41). Similarly, Tan-Pertel et al. (40) found that constitutive Notch activation enhances granulopoiesis, but inhibits terminal differentiation. Thus, controversy still remains regarding the effect of Notch signals on the myeloid potential of uncommitted progenitors, and whether Notch signals deviate committed myeloid progenitors to other lineages (36), induce their apoptosis (42), or have no effect at all (12). Further, the targets of Notch signaling in multipotent HPCs are largely unknown.

In this study, we show that myelopoiesis from HPCs obtained from fetal liver, adult bone marrow, or ESCs is severely inhibited in OP9-DL1 vs OP9-control cell cultures. In contrast, committed myeloid progenitors complete their maturational program regardless of the presence of Dll-1. Further, we demonstrate that the Dll-1-induced inhibition of myelopoiesis is mediated by canonical Notch signals within the hemopoietic cell, as the developmental block is absent when CSL–/– ESCs are cultured on OP9-DL1 cells. In agreement with others (43, 44), we find that GATA-2 expression is maintained in HPCs undergoing differentiation in the presence of Notch signals. Importantly, we extend previous findings to establish that modulation of GATA-2 expression is required to mediate the Dll-1-induced defect in myelopoiesis, as HPCs that lack GATA-2 do not demonstrate a Notch/Dll-1-mediated inhibition of myelopoiesis. Thus, our results provide a potential mechanism for the observed function of Dll-1 in mediating the lineage restriction of differentiating HPCs within specific microenvironments.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Mice

Timed-pregnant Swiss.NIH and C57BL/6 CD45.1 congenic mice were obtained from the National Cancer Institute, Frederick Cancer Research and Development Center (Frederick, MD).

Differentiation of ESCs and cell culture

OP9 cells were originally obtained from Dr. T. Nakano (Osaka University, Osaka, Japan) and the Riken Cell Bank (Tsukuba, Japan). The generation of OP9-DL1 cells and OP9-control (GFP only) cells is described in Schmitt and Zúñiga-Pflücker (23). ESC/OP9 differentiation cocultures, using either OP9-DL1 or OP9-control cells and ESR1, CSL+/–, CSL–/–, GATA-2+/–, or GATA-2–/– ESCs were performed as previously described (15, 16). Briefly, 5 x 104 ESCs were seeded onto either OP9-DL1 or OP9-control cell monolayers in 10-cm dishes. After 5 days of coculture, cells were harvested and resuspended into single-cell suspensions by 0.25% trypsin treatment and vigorous pipetting. Cells were then washed and reseeded onto new OP9 cell monolayers, with the addition of Flt3L at a final concentration of 5 ng/ml (R&D Systems). Nonadherent cells were passed to fresh OP9 monolayers again at day 8 of coculture, and thereafter ESC/OP9 cocultures were maintained in the presence of 5 ng/ml Flt3L and 1 ng/ml IL-7 (R&D Systems) by changing media or passaging (without trypsin) to fresh OP9 monolayers every 2 days. GATA-2–/– and GATA-2+/–ESC cultures, because of the reduced hemopoietic efficiency of GATA-2–/– progenitors, were seeded onto OP9-control stroma and either maintained on OP9-control stroma or transferred to OP9-DL1 cells at day 8. This was to relieve the GATA-2–/– progenitors from the early proliferative inhibition observed in the presence of OP9-DL1 cells (R. de Pooter, unpublished observations). HPCs isolated from adult bone marrow or fetal liver were cocultured similarly to ESC-derived progenitors: in the presence of Flt3L at a final concentration of 5 ng/ml and IL-7 at 1 ng/ml.

For experiments using presenilin inhibitors, Presenilin inhibitor X (Calbiochem) was added to cultures of sorted CD117+Sca-1high fetal liver-derived HPCs on OP9-DL1 cells, at a final concentration of 0.33 or 1.0 µM. The same volume of DMSO was added to all cultures, including the control culture.

Isolation of HPCs

HPCs were isolated from day 14 fetal liver cells by enriching for CD24low lineage marker (Lin)– fetal liver cells by anti-CD24 Ab-/complement-mediated depletion (45), and sorting for the expression of CD117 and Sca-1. HPC from adult bone marrow were isolated by staining for Lin (Ter119, CD11b, Gr-1, CD4, CD8, CD45R), CD117, and Sca-1, and sorting based on a phenotype of Lin–CD117+Sca-1high. Equal numbers of CD117+Sca-1high HPC were seeded onto a ~80% confluent monolayer of either OP9-DL1 or OP9-control cells. To isolate common myeloid progenitors (CMP) and granulocyte-macrophage progenitors (GMP) from adult bone marrow (46), Lin+ (CD3, CD4, CD8, CD45R, Gr-1, Ter119, CD19, IgM), IL-7R+, and Sca-1+ cells were depleted by MACS (Miltenyi Biotec). The resulting populations were then sorted based on expression of CD16/32 and CD34: CMP as CD117+CD34+CD16/32int, and GMP as CD117+CD34+CD16/32+ (46).

For RT-PCR analysis, fetal liver cells were obtained from 13 days postcoitus (dpc) embryos, depleted of CD24high cells by anti-CD24 Ab-/complement-mediated depletion (45), stained with PE-conjugated anti-Sca-1 Ab (BD Biosciences) and anti-PE magnetic microbeads (Miltenyi Biotec), and Sca-1+ cells were isolated by two successive double-sensitive enrichments. Cocultures were harvested by vigorous pipetting at the indicated time points.

Flow cytometry and cell sorting

Flow cytometry was performed using a FACSCalibur instrument (BD Biosciences) as previously described (45). FITC-, PE-, biotin-, and allophycocyanin-conjugated Abs, and CyChrome-conjugated streptavidin were purchased from BD Biosciences. For analysis of hemopoietic cells, live cells were gated based on forward and side scatter, lack of propidium iodide uptake, and lack of GFP expression. Cells were sorted using a FACSDiVa instrument (BD Biosciences) as previously described. Sorted cells were >99% pure, as determined by postsort analysis.

MACS enrichment was performed according the manufacturer’s instructions (Miltenyi Biotec).

Limiting dilution assay

For the limiting dilution assay, equal numbers of sorted CD117+Sca-1high fetal liver-derived HPCs were cultured on either OP9-control or OP9-DL1 cells for 5 days, in the presence of Flt3L at a final concentration of 5 ng/ml. Following 5 days of culture, 12 replicates each of 100, 30, 10, 3, or 1 cell(s)/well were deposited by Clonecyte sorting into 96-well plates containing confluent monolayers of OP9-control cells. The fetal liver-derived cells were then cultured in the presence of Flt3L at a final concentration of 5 ng/ml and IL-3 at a final concentration of 1 ng/ml (R&D Systems), and analyzed by flow cytometry after 8 days.

PCR and RT-PCR

For RT-PCR analysis, cDNA was prepared from OP9-control and OP9-DL1 cells, freshly isolated ex vivo CD117+Sca-1high sorted fetal liver progenitors, freshly isolated ex vivo CD44+CD25–CD117+ (DN1 subset) and CD44+CD25+CD117+ (DN2 subset) day 14 fetal thymocytes (47), and Sca-1+ MACS-enriched fetal liver progenitors cultured for 37 h on either OP9-DL1 or OP9-control stroma. In the latter case, contamination by OP9 cells was minimized by sorting CD45+ cells directly into TRIzol (Invitrogen Life Technologies). OP9-control and OP9-DL1 cells were also collected using TRIzol. For the time course comparing GATA-2 expression patterns in CSL–/– vs CSL+/– ESC-derived hemopoietic cells, ESCs were seeded onto OP9-control cells as described above, and transferred to OP9-DL1 cells at day 8 of coculture. RNA was isolated following manufacturer’s instructions. All semiquantitative PCRs were performed using 3-fold serial dilutions of cDNA that had been normalized by beta-actin or CD45 expression, as indicated, using the following primers: beta-actin forward GTGGGCCGCTCTAGGCACCAA, beta-actin reverse CTCTTTGATGTCACGCACGATTTC (535 bp, 55°C); CD45 forward CTACGCAAAGCACGGCCTG, CD45 reverse TCGAGTCTGCGTTGTCCCAC (340 bp, 52°C); GATA-2 forward ACACAC CACCCGATACCCACCTAT, GATA-2 reverse CCTACGCCATGGCAGTCACCATGCT (720 bp, 60°C) (43); PU.1 forward ATGGAAGGGTTTTCCCTCACCGCC, PU-1 reverse GTCCACGCTCTGCAGCTCTGTGAA (216 bp. 61°C) (48); CD3 forward ATGGCCAAGAGCTGC, CD3 reverse AGAATACAGGTCCCGCT (384 bp, 57°C); CD25 forward GTTGGGGTTTCTCTCATTA, CD25 reverse GGTGGTGTTCTCTTTCATC (542 bp, 55°C); TCR C{alpha} forward, AGAACCTGCTGTGTACCAGTTAA, TCR C{alpha} reverse CATGAGCAGGTTAAATCCGGCT (331 bp, 60°C). The PCR program was as follows: 95°C for 2 min, and 30 cycles (beta-actin, CD45) or 34 cycles (all others) of 95°C for 30 s, annealing temperature for 30 s, 72°C for 30 s, followed by a 4-min extension at 72°C.

Immunohistochemistry

Cell smears were taken down to water, and endogenous peroxidase was blocked with 3% H2O2 for 15 min. Smears were then rinsed with water, followed by Tris buffer and blocked for 10 min with 10% nonimmune goat serum (Zymed Laboratories). Cell smears were stained with rabbit anti-myeloperoxidase at 1/2000 (DakoCytomation) at room temperature for 1 h. Cell smears were then washed three times with Tris, incubated with biotinylated goat anti-rabbit Ab (Zymed Laboratories) for 15 min at room temperature, and washed three more times. Cell smears were then labeled with peroxidase-conjugated streptavidin (Zymed Laboratories) and washed three more times with Tris before being developed in diaminobenzidine solution for 10 min. Cell smears were then washed with water and counterstained in Harris hematoxylin before being dehydrated, cleared, and mounted.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Defect in myelopoiesis in the presence of Dll-1

OP9 cells, despite their failure to produce M-CSF (17), support the production of myeloid cells from various sources of HPC (14, 15) (Fig. 1A, upper panels). This allowed us to examine the effects of Dll-1 on myelopoietic potential by coculturing uncommitted HPCs on either OP9-DL1 or OP9-control cells.


Figure 1
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  		FIGURE 1. Myelopoiesis by uncommitted progenitors is inhibited in the presence of Dll-1. A, Day 15 ESC-, day 7 fetal liver-, and day 8 adult bone marrow-derived HPCs, cocultured with either OP9-control cells or OP9-DL1 cells, were analyzed for CD45 and CD11b expression. Data shown are representative of at least three independent experiments, with the fold changes in myelopoiesis between OP9-DL1 and OP9-control cultures ranging from: ESCs 6- to 20-fold, fetal liver 12- to 40-fold, bone marrow 5- to 50-fold. B, Fetal liver HPCs cultured on either OP9-control or OP9-DL1 cells for 8 days were analyzed for expression of myeloperoxidase, an enzyme associated with neutrophil granules. Slides shown at x40 magnification.

 
We found that OP9-control cells supported robust myelopoiesis from ESC-, fetal liver-, and bone marrow-derived hemopoietic cells, as defined by the expression of CD45 and high levels of CD11b (Fig. 1A, upper panels). In contrast, there was a marked absence of CD45+CD11bhigh myeloid cells on OP9-DL1 cells (Fig. 1A, lower panels). Instead, as we have previously shown (23, 49), the majority of progenitors cultured on OP9-DL1 cells gave rise, at later time points, to T lineage cells (data not shown). In addition, we examined the expression of myeloperoxidase, an enzyme contained within the granules of neutrophils and their progenitors. Although positive cells were clearly present in OP9-control cultures, they were not found in OP9-DL1 cocultures (Fig. 1B). Furthermore, when we compared ESC- and fetal liver-derived culture on OP9-control or OP9-DL1 stroma for their expression of other myeloid-restricted surface markers, we found that while OP9-control cultures contained populations of cells with the CD11bhighCD11c+ phenotype characteristic of myeloid dendritic cells, and the Gr-1+CD11b+ phenotype associated with granulocytes and immature monocytes, cocultures with OP9-DL1 cells did not (Fig. 2A). Most strikingly, the more mature myeloid phenotype characterized by high levels of MHC class II (MHC II) expression was completely absent in OP9-DL1 cocultures, but comprised over one-quarter of the CD11bhigh population present on OP9-control cells (Fig. 2A). Finally, we observed no CD11b+CD19+ cells, indicating that, not only do our CD11b+ cells express other characteristic myeloid markers, but they do not express a definitive B cell marker. This supports the conclusion that our observations reflect a loss of myeloid cells in OP9-DL1 cocultures, not just a down-regulation of CD11b, or a loss of CD11b-expressing B cells (Fig. 2B).


Figure 2
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  		FIGURE 2. Myeloid subsets are inhibited in the presence of Dll-1. A, Day 15 ESC and day 8 fetal liver cultures were compared for their expression of various surface markers of myeloid cells, when cultured in the presence of either OP9-control or OP9-DL1 cells. B, Day 16 ESC cultures were analyzed for the presence CD11b and CD19 surface staining. Data shown are representative of at least three independent experiments.

 
To address whether the block in myelopoiesis was due only to a lack of proliferation by myeloid progenitors on OP9-DL1 cells, we performed a limiting dilution assay to determine the frequency of cells with myeloid potential. Fetal liver-derived HPCs were cultured on either OP9-control or OP9-DL1 cells for 5 days, and then decreasing numbers of cells per well were deposited onto OP9-control cells in 96-well plates. Following an additional 8 days of coculture with OP9-control cells, the contents of each well were analyzed by flow cytometry. The limiting dilution assay demonstrated that the frequency of cells with myeloid potential was 1 in 65 progenitors from cultures on OP9-control cells, but only 1 in 404 in cultures from OP9-DL1 cells (Table I). Although an apparent reduction in myeloid potential could result if the same absolute number of myeloid progenitors arose on either stroma, but were greatly outnumbered by T cell progenitors in OP9-DL1 cocultures, this was not the case, as progenitors on OP9-control cultures actually expanded 2- to 3-fold more than the same number of progenitors cultured on OP9-DL1 cells. Taken together, these results show that the environment provided by OP9-DL1 cells strongly inhibits the generation of myeloid progenitors from uncommitted primary or ESC-derived HPCs, in contrast to that provided by OP9-control cells.


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  		Table I. Myeloid progenitor frequency analysis for fetal liver HPCs cultured on OP9-control or OP9-DL1 cells

 
Committed immature myeloid progenitors are refractory to a Notch signaling-mediated block in maturation

To address whether the OP9-DL1 environment could inhibit the maturation of committed myeloid progenitors, bone-marrow-derived Lin–CD117+Sca-1–CD34+ CMP and GMP, as defined by Akashi et al. (46) on the basis of CD16/32 expression, were cultured on either OP9-control or OP9-DL1 cells for 3 days. CD16/32int CMPs and CD16/32+ GMPs appeared to be unimpeded in their ability to generate CD11bhigh myeloid cells in OP9-DL1 cultures (Fig. 3), although CMPs showed a slight 2-fold reduction in cell yields. Thus, committed myeloid progenitors are not sensitive to an OP9-DL1-induced developmental block, as they can mature to express high cell surface levels of CD11b in the presence of either OP9-DL1 or OP9-control cells. No evidence for apoptosis was observed, as these populations persisted on both stroma for at least 7 days (data not shown).


Figure 3
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  		FIGURE 3. Committed immature myeloid progenitors are refractory to a Notch signaling-mediated block in maturation. GMPs (A) and CMPs (B), cultured on either OP9-control cells or OP9-DL1 cells, were analyzed for CD45 and CD11b expression. Data shown are representative of three independent experiments.

 
The block in myelopoiesis is due to Notch signaling within the HPC

To confirm that the observed defect in myelopoiesis was due to Notch signaling, we cultured fetal liver HPCs on OP9-DL1 cells in the presence or absence of a Presenilin inhibitor specific for both Presenilins-1 and -2. We found that the percentage of CD11bhigh cells increased with increasing concentrations of this inhibitor (Fig. 4A).


Figure 4
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  		FIGURE 4. The block in myelopoiesis is due to Notch signaling. A, The percentage of CD45+CD11bhigh cells was analyzed in day 7 fetal liver cocultures on OP9-DL1 in the presence of increasing concentrations of presenilin inhibitor. B, Day 16 CSL+/– and CSL–/– ESC/OP9-DL1 were analyzed for the expression of CD45 and CD11b. Plots shown are gated on the CD45+ population. Data shown are representative of at least three independent experiments.

 
This observation indicated that Notch signaling was involved in the myelopoietic defect, but could not distinguish whether this defect was the direct result of Notch signaling intrinsic to the HPCs, or resulted indirectly from a Notch signaling-mediated effect on the OP9 cells, which altered their expression of some other unidentified factor critical to myeloid development.

To distinguish between these two possibilities, we used HPCs derived from CSL–/– ESCs. When cocultured with OP9-DL1 cells, CSL+/– ESC-derived HPCs behaved as wild-type cells (Fig. 1A), exhibiting a profound defect in myelopoiesis (Fig. 4B), and giving rise to T lineage cells at later time points (data not shown). In contrast, CSL–/– ESCs produced robust populations of myeloid cells in the presence of Dll-1 (Fig. 4B). These results demonstrate that Notch signaling within the HPC is sufficient and required to mediate the block in myelopoiesis observed in cocultures with OP9-DL1 cells, and that the induced inhibition is downstream of the canonical Notch-signaling pathway mediated by CSL.

GATA-2 is differentially expressed in OP9-DL1 vs OP9-control cocultures

Our results showed that the diminished myelopoiesis from HPCs cultured on OP9-DL1 cells was the result of Notch signals within the hemopoietic cells. Thus, to address the mechanism of this block, we used semiquantitative RT-PCR to examine the expression of several genes known to be involved in hemopoiesis. To this end, RNA was isolated from fetal liver HPCs cultured on either OP9-control or OP9-DL1 cells for 37 h. This early time point was chosen to focus on events proximal to Notch signals. Although several genes were examined, strikingly, analysis by densitometry showed that GATA-2 expression was up-regulated ~3.5-fold in HPCs cultured on OP9-DL1 cells, compared with those on OP9-control cells (Fig. 5A). In contrast, PU.1 expression was not differentially affected in HPCs cultured on either stroma at this early time point (Fig. 5A). In addition, we also examined whether GATA-2 is expressed by thymocyte progenitors. Fig. 5B shows that the earliest thymic immigrants, which are contained within the CD117+CD44+CD25– subset (DN1 cells), express detectable levels of GATA-2, in contrast to the T cell-specified CD117+CD44+CD25+ subset (DN2 cells). This observation suggests in vivo relevance for the increased expression of GATA-2 by fetal liver HPCs cultured on OP9-DL1 vs OP9-control cells. We then examined the expression of GATA-2 by ESC-derived cells in the presence or absence of Notch signals, by comparing GATA-2 mRNA levels in CSL+/– or CSL–/– ESC-derived HPCs cultured on OP9-control cells for 8 days, and then transferred to OP9-DL1 cells to induce Notch signals. GATA-2 expression was analyzed from the ESC cocultures at the indicated culture periods, and normalized to CD45 expression levels as a gauge of hemopoietic cell contribution. Our results show that, similar to fetal liver-derived hemopoietic cells, ESC-derived HPCs sustained GATA-2 expression in the presence (CSL+/–), but not the absence (CSL–/–), of Notch signals (Fig. 5C). These initial results suggested that sustained GATA-2 expression in HPCs, as a consequence of Dll-1-induced Notch signals, could be proximally involved in the myelopoietic block observed on OP9-DL1 cells.


Figure 5
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  		FIGURE 5. GATA-2 is differentially expressed in OP9-DL1 vs OP9-control cocultures, and early thymocyte subsets. A, For RT-PCR analysis, cDNA was prepared from OP9-control and OP9-DL1 cells, freshly isolated ex vivo CD117+Sca-1high fetal liver HPCs, and Sca-1+ MACS-enriched fetal liver progenitors cultured for 37 h on either OP9-DL1 or OP9-control stroma, and analyzed for the expression of the indicated gene products. Normalized against the intensity of CD45 by densitometric analysis, hemopoietic cells cultured in the presence of OP9-DL1 cells expressed 3.5-fold more GATA-2. B, cDNA was prepared from sorted DN1 (CD44+CD25–CD117+) and DN2 (CD44+CD25+CD117+) day 14 fetal thymocyte subsets and total fetal thymus, and analyzed for the expression of the indicated gene products. C, cDNA was prepared from CSL+/– or CSL–/– ESC-derived cells from OP9 cocultures as described in Materials and Methods and analyzed for the expression of the indicated gene products.

 
GATA-2 is required to mediate the block in myelopoiesis

The observed Notch signaling-dependent increase in GATA-2 expression by fetal liver- or ESC-derived HPCs was consistent with results from Kumano et al. (43) using the 32D myeloid cell line transduced with ICN. Given that overexpression of GATA-2 did not cause a maturational block in this cell line model system, we asked whether GATA-2 is required for the Dll-1-induced block of myelopoiesis observed in primary cells. GATA-2 deficiency is embryonic lethal before 10.5 dpc and embryos exhibit severe anemia due to the requirement for GATA-2 for efficient expansion of HSCs, precluding the use of GATA-2–/– fetal liver or adult bone marrow (50). Nevertheless, GATA-2–/– ESCs give rise to myeloid lineage cells when cultured on either OP9-control or OP9-DL1 cells (Fig. 6A). This is in marked contrast to the case of wild-type ESCs (Fig. 1A) or GATA-2+/– ESCs (Fig. 6A), which showed a dramatic reduction in myelopoiesis when cultured on OP9-DL1 cells, as compared with OP9-control cells (Fig. 6). The ability of GATA-2–/– ESCs to give rise to myeloid cells is not due simply to their inability to respond to the OP9-DL1 environment, as these cells give rise to T cells when cultured on OP9-DL1 cells (Fig. 6B). Although, in keeping with their reduced potential for early hemopoietic expansion, fewer T cells were produced in the absence of GATA-2, the percentages of CD4+CD8+ cells that were generated were similar to wild-type ESC cultures.


Figure 6
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  		FIGURE 6. GATA-2 is required to inhibit myelopoiesis in the presence of Dll-1, but is not required for T lymphopoiesis. A, Day 17 GATA-2+/– and GATA-2–/– ESC/OP9-control and OP9-DL1 cocultures were analyzed for the presence of CD45+CD11bhigh myeloid cells. Plots shown are gated on CD45+ population. B, Day 19 cocultures of GATA-2+/.+ and GATA-2–/– were analyzed for the expression of CD4 and CD8, indicative of the presence of T lineage cells. Data shown are representative of three independent experiments.

 
Taken together, our findings indicate that modulation of GATA-2 expression by Notch signals is required to mediate the defect in HPC commitment to the myeloid lineage, and that this defect and the commitment of progenitors to the T cell fate appear to be distinct events.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Environmental signals play an important role in the outcome of hemopoiesis. Interestingly, while thymic epithelial cells express high levels of Dll-1 (20, 21), bone marrow stroma does not (19), although the levels of Dll-1 to which progenitors are exposed in the thymus, vs various niches within the bone marrow, remains unresolved. In this study, we demonstrate that OP9-DL1 cells inhibit myelopoiesis from HPCs, in contrast to OP9-control cells (23). Our findings, taken together with the observation that the thymic environment actively discourages the adoption of non-T cell fates (51, 52), strongly support a physiological role for Notch receptor/Dll-1 interactions in preventing significant myelopoiesis from occurring in the thymus from thymus-seeding progenitor cells, which are known to possess myelopoietic potential (53, 54). Furthermore, our findings suggest that any myelopoiesis taking place within the thymus is likely derived from incoming myeloid committed cells.

Although it has been suggested that signals downstream of Notch are involved in inhibiting myelopoiesis, there are conflicting reports regarding this and potential underlying mechanisms. Our results suggest that some of these contradictions might have arisen because only uncommitted progenitors are susceptible to a Notch-mediated inhibition in myelopoiesis. Indeed, two recent findings using the same model system, but starting with different types of HPC, have shown differing degrees of myeloid inhibition in response to Notch-induced signals (44, 55). On one hand, fetal liver CD117+ progenitors showed nearly a 3-fold reduction in myelopoiesis (44), while purified bone marrow CD117+Sca-1high HSCs showed considerably greater 14-fold change between OP9-DL1 and OP9-control cultures (55). This discrepancy in fold change is probably the result of committed myeloid progenitors present in the CD117+Sca-1– population, consistent with our unpublished observations that this population contains sufficient committed myeloid progenitors to obscure any defect in myelopoiesis in the cultures with OP9-DL1 stroma. Furthermore, other models using the 32D cell line have yielded disparate conclusions regarding the role of Notch in myelopoiesis. However, as it is unclear where the 32D cell line falls on the gradient between specification and commitment to the myeloid lineage, it is possible that the conflicting results obtained using this cell line, finding either that Notch signals inhibit (26, 29, 30, 43) or promote myeloid differentiation (38, 39, 40), may reflect clonal variation in this regard. The requirement to transfect 32D cells with exogenous Notch to render them responsive to Notch ligands, despite the presence of endogenous Notch expression (40), might also reflect a resistance to physiological levels of Notch signaling.

Although previous analysis of Notch1 deficient in vivo models did not report an accumulation of myeloid cells, this does not exclude a physiological relevance for Dll-1-mediated signals in inhibiting myeloid commitment, as such an accumulation might not have been examined. Further, in the absence of Notch signaling, the B cell fate is available to progenitors entering the thymus, and in the IL-7-rich lymphopoietic thymic environment, B cells would have a proliferative advantage over myeloid cells.

An important aspect of the OP9 model system is differentiation kinetics, as HSCs do not undergo long-term self-renewal in this environment, and the various lineages arise in successive waves. Specifically, erythroid and myeloid lineages appear first, followed by lymphoid cells (16). Notably, at the time points shown here, neither culture is composed predominantly of lymphocytes, thus the defect in myelopoiesis observed in OP9-DL1 cocultures is not due to the same number of myeloid cells being masked by large numbers of T cell progenitors. Additionally, the block in myelopoiesis is not due to a rapid commitment of HPCs to the T cell fate on OP9-DL1 cells, as the ability of GATA-2–/– ESCs to give rise to both myeloid and T lineage cells on OP9-DL1 cells demonstrates that the defect in myelopoiesis and commitment to the T cell fate are mechanistically distinct events. Nor does the defect in myelopoiesis appear to be one of delayed kinetics, despite the evidence that Notch signals can delay cell cycle (32, 56), as myeloid populations do not arise even at later time points. Neither is Dll-1 simply preventing all hemopoietic differentiation, as OP9-DL1 cocultures produce robust populations of T lineage cells at later time points (23, 49).

Our results showing that GATA-2 expression is up-regulated by fetal liver- or ESC-derived HPCs in the presence of Dll-1 agree with observations by Kumano et al. (43), who also found that myeloid maturation in the 32D myeloid cell line could be blocked by transduction with ICN or HES-1. However, while this inhibition could be overcome by further transducing 32D cells with either a dominant-negative form of GATA-3 (DN-GATA3), which inhibits all GATA family members, or PU.1, overexpression of GATA-2 itself did not block myeloid maturation. As neither DN-GATA3 nor PU.1 is a specific inhibitor of GATA-2 function, these results, while suggestive, did not allow the conclusion that Notch acts through GATA-2 to interfere with normal myelopoiesis (43).

Recent findings in both nonhemopoietic systems (57) and peripheral T cells (58) have demonstrated regulatory interactions between Notch signals and GATA family transcription factors during differentiation. Importantly, given our hypothesis that Notch signaling-induced modulation of GATA-2 expression is involved in restricting non-T lineage fate choices within the thymus, we find that DN1 thymocytes express GATA-2. As GATA-2 expression is lost by the T cell-committed DN2 stage of thymocyte development, other mechanisms must later assume the role of excluding the myeloid fate, indicating that GATA-2 is not required for maintenance of the T cell fate. In fact, we have demonstrated that GATA-2 is not required for T lymphopoiesis, as GATA-2–/– ESCs produce CD4+CD8+ T lineage cells when cultured on OP9-DL1 cells (Fig. 6). In contrast, we have directly assessed the role of GATA-2 in mediating the Dll-1/Notch-induced block of myelopoiesis, and demonstrate that the absence of GATA-2 alleviates the defect in myelopoiesis observed in the presence of Dll-1. Although our results do not allow us to draw conclusions regarding how Notch signals control GATA-2 up-regulation, recent work by others has demonstrated the presence of CSL-binding sites in the regulatory regions of the GATA-2 gene (57), suggesting that ICN can directly up-regulate GATA-2 expression.

We chose to examine a system in which GATA-2 was absent, instead of one in which GATA-2 was overexpressed. This avoided the inherent, confounding problems of forced overexpression of GATA-2 in hemopoiesis (59, 60).

Although GATA-2 is required for the efficient expansion of the HSC pool during development, and its homeostasis in the adult, GATA-2-deficient cells appear to be capable of the full range of hemopoietic differentiation (50, 61, 62), suggesting that hemopoietic commitment in the absence of GATA-2 is not sufficiently perturbed to preclude the conclusions drawn from our model system.

A role for GATA-2 in inhibiting myelopoiesis is consistent with the ability of GATA-2 to physically interact with the Ets transcription factor PU.1 and interfere with its ability to transactivate target genes (60, 63, 64). This suggests that the inability of Notch signals to inhibit myelopoiesis from committed progenitors might be due to the high levels of PU.1 expressed by committed myeloid progenitors, which could preclude effective inhibition by GATA-2. Alternatively, myeloid progenitors may no longer be responsive to Notch signals. Initially, GATA-2 does not appear to down-regulate the expression of PU.1, but rather to interfere with its function (64), although GATA-2 may inhibit PU.1 expression at later time points (60). This mechanism is consistent with our observation that PU.1 expression is equivalent between HPCs cultured on either OP9-control or OP9-DL1 cells, and with antagonism between PU.1 and another GATA family member, GATA-1, during erythropoiesis (64, 65, 66, 67). Furthermore, while GATA-2 expression is required for efficient early hemopoiesis (50, 68, 69), it is dispensable for myeloid terminal differentiation (69), and is antagonized by PU.1 during macrophage differentiation (63).

In turn, while PU.1–/– embryos lack significant numbers of T cells, B cells, and macrophages (70, 71), PU.1 seems to be only absolutely required for B lymphopoiesis and myelopoiesis, as adult PU.1–/– HPCs are capable of low levels of T cell development (72). Thus, taken with other findings demonstrating that graded expression of PU.1 is involved in B cell vs macrophage lineage commitment (73), it is possible that Notch/Dll-1 interactions could inhibit PU.1 function to below the level required for B and myeloid cell production without abrogating T cell development.

Taken together, our results, and the existing literature on the role of GATA-2 in hemopoiesis and its interactions with other hemopoietic transcription factors (74), lead us to propose a model in which sustained GATA-2 expression downstream of Dll-1/Notch receptor interactions may contribute to the lineage restriction of early thymic immigrants.


   Acknowledgments
 
We thank Dr. Janet Rossant (Mt. Sinai Hospital, Toronto, Ontario, Canada) for her helpful advice, suggestions, and gift of reagents; Dr. Michele Anderson (Sunnybrook & Women’s Research Institute, Toronto, Ontario, Canada) for valuable discussion and suggestion; Gisele Knowles for her expert assistance with flow cytometry and cell sorting; and Kevin Kwok for his expert assistance with immunohistochemistry. R. de Pooter thanks Georges de Pooter (1914–2004) for his encouragement.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by the National Cancer Institute of Canada, with funds from the Canadian Cancer Society. R.F.d.P. is supported by a Doctoral Research Award from the Canadian Institute of Health Research. J.C.Z.-P. is supported by a Canada Research Chair in Developmental Immunology. Back

2 Address correspondence and reprint requests to Dr. Juan Carlos Zúñiga-Pflücker, Department of Immunology, University of Toronto, Sunnybrook & Women’s Research Institute, 2075 Bayview Avenue, Room A-331, Toronto, Ontario M4N 3M5, Canada. E-mail address: jczp{at}swri.ca Back

3 Abbreviations used in this paper: HSC, hemopoietic stem cell; Dll, Delta-like; ESC, embryonic stem cell; HPC, hemopoietic progenitor; CMP, common myeloid progenitor; CSL, CBF1/supressor of hairless/Lag-1; dpc, days postcoitus; GMP, granulocyte-macrophage progenitor; ICN, intracellular Notch. Back

Received for publication December 12, 2005. Accepted for publication February 20, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

  1. Morrison, S. J., N. Uchida, I. L. Weissman. 1995. The biology of hematopoietic stem cells. Annu. Rev. Cell Dev. Biol. 11: 35-71. [Medline]
  2. Zhang, J., C. Niu, L. Ye, H. Huang, X. He, W. G. Tong, J. Ross, J. Haug, T. Johnson, J. Q. Feng, et al 2003. Identification of the haematopoietic stem cell niche and control of the niche size. Nature 425: 836-841. [Medline]
  3. Calvi, L. M., G. B. Adams, K. W. Weibrecht, J. M. Weber, D. P. Olson, M. C. Knight, R. P. Martin, E. Schipani, P. Divieti, F. R. Bringhurst, et al 2003. Osteoblastic cells regulate the haematopoietic stem cell niche. Nature 425: 841-846. [Medline]
  4. Artavanis-Tsakonas, S., M. D. Rand, R. J. Lake. 1999. Notch signaling: cell fate control and signal integration in development. Science 284: 770-776. [Abstract/Free Full Text]
  5. Martinez Arias, A., V. Zecchini, K. Brennan. 2002. CSL-independent Notch signalling: a checkpoint in cell fate decisions during development?. Curr. Opin. Genet. Dev. 12: 524-533. [Medline]
  6. Conlon, R. A., A. G. Reaume, J. Rossant. 1995. Notch1 is required for the coordinated segmentation of somites. Development 121: 1533-1545. [Abstract]
  7. Swiatek, P. J., C. E. Lindsell, F. F. del Amo, G. Weinmaster, T. Gridley. 1994. Notch1 is essential for postimplantation development in mice. Genes Dev. 8: 707-719. [Abstract/Free Full Text]
  8. Oka, C., T. Nakano, A. Wakeham, J. L. de la Pompa, C. Mori, T. Sakai, S. Okazaki, M. Kawaichi, K. Shiota, T. W. Mak, T. Honjo. 1995. Disruption of the mouse RBP-J{kappa} gene results in early embryonic death. Development 121: 3291-3301. [Abstract]
  9. Walker, L., A. Carlson, H. T. Tan-Pertel, G. Weinmaster, J. C. Gasson. 2001. The Notch receptor and its ligands are selectively expressed during hematopoietic development in the mouse. Stem Cells 19: 543-552. [Medline]
  10. Singh, N., R. A. Phillips, N. N. Iscove, S. E. Egan. 2000. Expression of notch receptors, notch ligands, and fringe genes in hematopoiesis. Exp. Hematol. 28: 527-534. [Medline]
  11. Jonsson, J.-I., Z. Xiang, M. Pettersson, M. Lardelli, G. Nilsson. 2001. Distinct and regulated expression of Notch receptors in hematopoietic lineages and during myeloid differentiation. Eur. J. Immunol. 31: 3240-3247. [Medline]
  12. Pui, J. C., D. Allman, L. Xu, S. DeRocco, F. G. Karnell, S. Bakkour, J. Y. Lee, T. Kadesch, R. R. Hardy, J. C. Aster, W. S. Pear. 1999. Notch1 expression in early lymphopoiesis influences B versus T lineage determination. Immunity 11: 299-308. [Medline]
  13. Radtke, F., A. Wilson, G. Stark, M. Bauer, J. van Meerwijk, R. MacDonald, M. Aguet. 1999. Deficient T cell fate specification in mice with an induced inactivation of. Notch1. Immunity 10: 547-558. [Medline]
  14. Nakano, T., H. Kodama, T. Honjo. 1996. In vitro development of primitive and definitive erythrocytes from different precursors. Science 272: 722-724. [Abstract]
  15. Nakano, T., H. Kodama, T. Honjo. 1994. Generation of lymphohematopoietic cells from embryonic stem cells in culture. Science 265: 1098-1101. [Abstract/Free Full Text]
  16. Cho, S. K., T. D. Webber, J. R. Carlyle, T. Nakano, S. M. Lewis, J. C. Zúñiga-Pflücker. 1999. Functional characterization of B lymphocytes generated in vitro from embryonic stem cells. Proc. Natl. Acad. Sci. USA 96: 9797-9802. [Abstract/Free Full Text]
  17. Kodama, H., M. Nose, S. Niida, S. Nishikawa. 1994. Involvement of the c-kit receptor in the adhesion of hematopoietic stem cells to stromal cells. Exp. Hematol. 22: 979-984. [Medline]
  18. de Pooter, R. F., S. K. Cho, J. R. Carlyle, J. C. Zúñiga-Pflücker. 2003. In vitro generation of T lymphocytes from embryonic stem cell-derived pre-hematopoietic progenitors. Blood 102: 1649-1653. [Abstract/Free Full Text]
  19. Bertrand, F. E., C. E. Eckfeldt, J. R. Fink, A. S. Lysholm, J. A. Pribyl, N. Shah, T. W. LeBien. 2000. Microenvironmental influences on human B-cell development. Immunol. Rev. 175: 175-186. [Medline]
  20. Anderson, G., J. Pongracz, S. Parnell, E. J. Jenkinson. 2001. Notch ligand-bearing thymic epithelial cells initiate and sustain Notch signalling in thymocytes independently of T cell receptor signaling. Eur. J. Immunol. 31: 3349-3354. [Medline]
  21. Parreira, L., H. Neves, S. Simoes. 2003. Notch and lymphopoiesis: a view from the microenvironment. Semin. Immunol. 15: 81-89. [Medline]
  22. Zúñiga-Pflücker, J. C.. 2004. T-cell development made simple. Nat. Rev. Immunol. 4: 67-72. [Medline]
  23. Schmitt, T. M., J. C. Zúñiga-Pflücker. 2002. Induction of T cell development from hematopoietic progenitor cells by Delta-like-1 in vitro. Immunity 17: 749-756. [Medline]
  24. Varnum-Finney, B., C. Brashem-Stein, I. D. Bernstein. 2003. Combined effects of Notch signaling and cytokines induce a multiple log increase in precursors with lymphoid and myeloid reconstituting ability. Blood 101: 1784-1789. [Abstract/Free Full Text]
  25. Masuya, M., N. Katayama, N. Hoshino, H. Nishikawa, S. Sakano, H. Araki, K. Mitani, H. Suzuki, H. Miyashita, K. Kobayashi, et al 2002. The soluble Notch ligand, Jagged-1, inhibits proliferation of CD34+ macrophages progenitors. Int. J. Hematol. 75: 269-276. [Medline]
  26. Li, L., L. A. Milner, Y. Deng, M. Iwata, A. Banta, L. Graf, S. Marcovina, C. Friedman, B. J. Trask, L. Hood, B. Torok-Storb. 1998. The human homolog of rat Jagged1 expressed by marrow stroma inhibits differentiation of 32D cell through interaction with Notch1. Immunity 8: 43-55. [Medline]
  27. Ye, Q., J.-H. Shieh, G. Morrone, M. A. S. Moore. 2004. Expression of constitutively active Notch4 (Int-3) modulates myeloid proliferation and differentiation and promotes expansion of hematopoietic progenitors. Leukemia 18: 777-787. [Medline]
  28. Stier, S., T. Cheng, D. Dombkowshi, N. Carlesso, D. T. Scadden. 2002. Notch 1 activation increases hematopoietic stem cell self renewal in vivo and favors lymphoid over myeloid outcome. Blood 99: 2369-2378. [Abstract/Free Full Text]
  29. Milner, L. A., A. Bigas, R. Kopan, C. Brashem-Stein, I. D. Berstein, D. I. K. Martin. 1996. Inhibition of granulocytic differentiation by mNotch1. Proc. Natl. Acad. Sci. USA 93: 13014-13019. [Abstract/Free Full Text]
  30. Bigas, A., D. I. Martin, L. A. Milner. 1998. Notch1 and Notch2 inhibit myeloid differentiation in response to different cytokines. Mol. Cell. Biol. 18: 2324-2333. [Abstract/Free Full Text]
  31. Kawamata, S., C. Du, K. Li, C. Lavau. 2002. Overexpression of the Notch target genes Hes in vivo induces lymphoid and myeloid alterations. Oncogene 21: 3855-3863. [Medline]
  32. Carlesso, N., J. C. Aster, J. Sklar, D. T. Scadden. 1999. Notch1-induced delay of human hematopoietic progenitor cell differentiation is associated with altered cell cycle kinetics. Blood 93: 838-848. [Abstract/Free Full Text]
  33. Walker, L., M. Lynch, S. Silverman, J. Fraser, J. Boulter, G. Weinmaster, J. C. Gasson. 1999. The Notch/Jagged pathway inhibits proliferation of human hematopoietic progenitors in vitro. Stem Cells 17: 162-171. [Medline]
  34. Karanu, F. N., L. Yuefei, L. Gallacher, S. Sakano, M. Bhatia. 2003. Differential response of primitive human CD34– and CD34+ hematopoietic cells to the Notch ligand Jagged-1. Leukemia 17: 1366-1374. [Medline]
  35. Varnum-Finney, B., L. E. Purton, M. Yu, C. Brashem-Stein, D. Flowers, S. Staats, K. A. Moore, I. Le Roux, R. Mann, G. Gray, et al 1998. The Notch ligand, Jagged-1, influences the development of primitive hematopoietic precursor cells. Blood 91: 4084-4091.
  36. Ohishi, K., B. Varnum-Finney, R. E. Serda, C. Anasetti, I. D. Berstein. 2001. The Notch ligand, Delta-1, inhibits the differentiation of monocytes into macrophages but permits their differentiation into dendritic cells. Blood 98: 1402-1407. [Abstract/Free Full Text]
  37. Ohishi, K., B. Varnum-Finney, I. D. Berstein. 2002. Delta-1 enhances marrow and thymus repopulating ability of human CD34+CD38– cord blood cells. J. Clin. Invest. 110: 1165-1174. [Medline]
  38. Schroeder, T., U. Just. 2000. Notch signalling via RBP-j promotes myeloid differentiation. EMBO J. 19: 2558-2568. [Medline]
  39. Schroeder, T., H. Kohlhof, N. Rieber, U. Just. 2003. Notch signaling induces multilineage myeloid differentiation and up-regulates PU.1 expression. J. Immunol. 170: 5538-5548. [Abstract/Free Full Text]
  40. Tan-Pertel, H. T., L. Walker, D. Browning, A. Miyamoto, G. Weinmaster, J. C. Gasson. 2000. Notch signaling enhances survival and alters differentiation of 32D myeloblasts. J. Immunol. 165: 4428-4436. [Abstract/Free Full Text]
  41. Cheng, P., Y. Nefedova, L. Miele, B. A. Osborne, D. Gabrilovich. 2003. Notch signaling is necessary but not sufficient for differentiation of dendritic cells. Blood 103: 3980-3988.
  42. Ohishi, K., B. Varnum-Finney, D. Flowers, C. Anasetti, D. Myerson, I. D. Bernstein. 2000. Monocytes express high amounts of Notch and undergo cytokine specific apoptosis following interaction with the Notch ligand, Delta-1. Blood 95: 2847-2854. [Abstract/Free Full Text]
  43. Kumano, K., S. Chiba, K. Shimizu, T. Yamagata, N. Hosoya, T. Saito, T. Takahashi, Y. Hamada, H. Hirai. 2001. Notch1 inhibits differentiation of hematopoietic cells by sustaining GATA-2 expression. Blood 98: 3283-3289. [Abstract/Free Full Text]
  44. Taghon, T. N., E. S. David, J. C. Zúñiga-Pflücker, E. V. Rothenberg. 2005. Delayed, asynchronous, and reversible T-lineage specification induced by Notch/Delta signaling. Genes Dev. 19: 965-978. [Abstract/Free Full Text]
  45. Carlyle, J. R., J. C. Zúñiga-Pflücker. 1998. Requirement for the thymus in {alpha}beta T lymphocyte lineage commitment. Immunity 9: 187-197. [Medline]
  46. Akashi, K., D. Traver, T. Miyaamoto, I. L. Weissman. 2000. A clonogenic common myeloid progenitor that gives rise to all myeloid lineages. Nature 404: 193-197. [Medline]
  47. Schmitt, T. M., M. Ciofani, H. T. Petrie, J. C. Zúñiga-Pflücker. 2004. Maintenance of T cell specification and differentiation requires recurrent notch receptor-ligand interactions. J. Exp. Med. 200: 469-479. [Abstract/Free Full Text]
  48. Anderson, M. K., G. Hernandez-Hoyos, R. A. Diamond, E. V. Rothenberg. 1999. Precise developmental regulation of Ets family transcription factors during specification and commitment to the T cell lineage. Development 126: 3131-3148. [Abstract]
  49. Schmitt, T. M., R. F. de Pooter, M. A. Gronski, S. K. Cho, P. S. Ohashi, J. C. Zúñiga-Pflücker. 2004. Induction of T cell development and establishment of T cell competence from embryonic stem cells differentiated in vitro. Nat. Immunol. 5: 410-417. [Medline]
  50. Tsai, F.-Y., G. Keller, F. C. Kuo, M. Weiss, J. Chen, M. Rosenblatt, F. W. Alt, S. H. Orkin. 1994. An early hematopoietic defect in mice lacking the transcription factor GATA-2. Nature 371: 221-226. [Medline]
  51. Spangrude, G. J., R. Scollay. 1990. Differentiation of hematopoietic stem cells in irradiated mouse thymic lobes: kinetics and phenotype of progeny. J. Immunol. 145: 3661-3668. [Abstract]
  52. Abboud, S. L., M. Bunegin, N. Ghosh-Choudhury, K. Woodruff. 2003. Analysis of the mouse CSF-1 gene promoter in a transgenic mouse model. J. Histochem. Cytochem. 51: 941-949. [Abstract/Free Full Text]
  53. Porritt, H. E., L. L. Rumfelt, S. Tabrizifard, T. M. Schmitt, J. C. Zuniga-Pflucker, H. T. Petrie. 2004. Heterogeneity among DN1 prothymocytes reveals multiple progenitors with different capacities to generate T cell and non-T cell lineages. Immunity 20: 735-745. [Medline]
  54. Katsura, Y.. 2002. Redefinition of lymphoid progenitors. Nat. Rev. Immunol. 2: 127-132. [Medline]
  55. Huang, J., K. P. Garrett, R. Pelayo, J. C. Zuniga-Pflucker, H. T. Petrie, P. W. Kincade. 2005. Propensity of adult lymphoid progenitors to progress to DN2/3 stage thymocytes with Notch receptor ligation. J. Immunol. 175: 4858-4865. [Abstract/Free Full Text]
  56. Schroeder, T., U. Just. 2000. mNotch1 signaling reduces proliferation of myeloid progenitor cells by altering cell-cycle kinetics. Exp. Hematol. 28: 1206-1213. [Medline]
  57. Robert-Moreno, A., L. Espinosa, J. L. de la Pompa, A. Bigas. 2005. RBPj{kappa}-dependent Notch function regulates Gata2 and is essential for the formation of intra-embryonic hematopoietic cells. Development 132: 1117-1126. [Abstract/Free Full Text]
  58. Amsen, D., J. M. Blander, G. R. Lee, K. Tanigaki, T. Honjo, R. A. Flavell. 2004. Instruction of distinct CD4 T helper cell fates by different notch ligands on antigen-presenting cells. Cell 117: 515-526. [Medline]
  59. Persons, D. A., J. A. Allay, E. R. Allay, R. A. Ashmun, D. Orlic, S. M. Jane, J. M. Cunningham, A. W. Nienhuis. 1999. Enforced expression of the GATA-2 transcription factor blocks normal hematopoiesis. Blood 93: 488-499. [Abstract/Free Full Text]
  60. Kitajima, K., M. Tanaka, J. Zheng, H. Yen, A. Sato, D. Sugiyama, H. Umehara, E. Sakai, T. Nakano. 2006. Redirecting differentiation of hematopoietic progenitors by a transcription factor, GATA-2. Blood 107: 1857-1863. [Abstract/Free Full Text]
  61. Ling, K. W., K. Ottersbach, J. P. van Hamburg, A. Oziemlak, F. Y. Tsai, S. H. Orkin, R. Ploemacher, R. W. Hendriks, E. Dzierzak. 2004. GATA-2 plays two functionally distinct roles during the ontogeny of hematopoietic stem cells. J. Exp. Med. 200: 871-882. [Abstract/Free Full Text]
  62. Rodrigues, N. P., V. Janzen, R. Forkert, D. M. Dombkowski, A. S. Boyd, S. H. Orkin, T. Enver, P. Vyas, D. T. Scadden. 2005. Haploinsufficiency of GATA-2 perturbs adult hematopoietic stem-cell homeostasis. Blood 106: 477-484. [Abstract/Free Full Text]
  63. Walsh, J. C., R. P. DeKoter, H.-J. Lee, E. D. Smith, D. W. Lancki, M. F. Gurish, D. S. Friend, R. L. Stevens, J. Anastasi, H. Singh. 2002. Cooperative and antagonistic interplay between PU.1 and GATA-2 in the specification of myeloid cell fates. Immunity 17: 665-676. [Medline]
  64. Zhang, P., G. Behre, J. Pan, A. Iwama, N. Wara-Aswapati, H. S. Radomska, P. E. Auron, D. G. Tenen, Z. Sun. 1999. Negative cross-talk between hematopoietic regulators: GATA proteins repress PU.1. Proc. Natl. Acad. Sci. USA 96: 8705-8710. [Abstract/Free Full Text]
  65. Zhang, P., X. Zhang, A. Iwama, C. Yu, K. A. Smith, B. U. Mueller, S. Narravula, B. E. Torbett, S. H. Orkin, D. G. Tenen. 2000. PU.1 inhibits GATA-1 function and erythroid differentiation by blocking GATA-1 DNA binding. Blood 96: 2641-2648. [Abstract/Free Full Text]
  66. Rekhtman, N., F. Radparvar, T. Evans, A. I. Skoultchi. 1999. Direct interaction of hematopoietic transcription factors PU.1 and GATA-1: functional antagonism in erythroid cells. Genes Dev. 13: 1398-1411. [Abstract/Free Full Text]
  67. Nerlov, C., E. Querfurth, H. Kulessa, T. Graf. 2000. GATA-1 interacts with the myeloid PU.1 transcription factor and represses PU.1-dependent transcription. Blood 95: 2543-2551. [Abstract/Free Full Text]
  68. Minegishi, N., N. Suzuki, T. Yokomizo, X. Pan, T. Fujimoto, S. Takahashi, T. Hara, A. Miyajima, S.-I. Nishikawa, M. Yamamoto. 2003. Expression and domain-specific function of GATA-2 during differentiation of the hematopoietic precursor cells in midgestation mouse embryos. Blood 102: 896-905. [Abstract/Free Full Text]
  69. Tsai, F. Y., S. H. Orkin. 1997. Transcription factor GATA-2 is required for proliferation/survival of early hematopoietic cells and mast cell formation, but not for erythroid and myeloid terminal differentiation. Blood 89: 3636-3643. [Abstract/Free Full Text]
  70. Scott, E. W., M. C. Simon, J. Anastasi, H. Singh. 1994. Requirement of transcription factor PU.1 in the development of multiple hematopoietic lineages. Science 265: 1573-1577. [Abstract/Free Full Text]
  71. Scott, E. W., R. C. Fisher, M. C. Olson, E. W. Kehrli, M. C. Simon, H. Singh. 1997. PU.1 functions in a cell autonomous manner to control the differentiation of multipotential lymphoid-myeloid progenitors. Immunity 6: 437-447. [Medline]
  72. Spain, L. M., A. Guerriero, S. Kunjibettu, E. W. Scott. 1999. T cell development is PU.1-deficient mice. J. Immunol. 163: 2681-2687. [Abstract/Free Full Text]
  73. DeKoter, R. P., H. Singh. 2000. Regulation of B lymphocyte and macrophage development by graded expression of PU.1. Science 288: 1439-1441. [Abstract/Free Full Text]
  74. Ranganath, S., K. M. Murphy. 2001. Structure and specificity of GATA proteins in Th2 development. Mol. Cell. Biol. 21: 2716-2725. [Abstract/Free Full Text]



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The OP9-DL1 System: Generation of T-Lymphocytes from Embryonic or Hematopoietic Stem Cells In Vitro
CSH Protocols, February 1, 2009; 2009(2): pdb.prot5156 - pdb.prot5156.
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 BloodHome page
N. P. Rodrigues, A. S. Boyd, C. Fugazza, G. E. May, Y. Guo, A. J. Tipping, D. T. Scadden, P. Vyas, and T. Enver
GATA-2 regulates granulocyte-macrophage progenitor cell function
Blood, December 15, 2008; 112(13): 4862 - 4873.
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 Mol. Cell. Biol.Home page
M.-J. Yoon, B.-K. Koo, R. Song, H.-W. Jeong, J. Shin, Y.-W. Kim, Y.-Y. Kong, and P.-G. Suh
Mind bomb-1 Is Essential for Intraembryonic Hematopoiesis in the Aortic Endothelium and the Subaortic Patches
Mol. Cell. Biol., August 1, 2008; 28(15): 4794 - 4804.
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L. Zhou, L. W. Li, Q. Yan, B. Petryniak, Y. Man, C. Su, J. Shim, S. Chervin, and J. B. Lowe
Notch-dependent control of myelopoiesis is regulated by fucosylation
Blood, July 15, 2008; 112(2): 308 - 319.
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 Proc. Natl. Acad. Sci. USAHome page
M. Sakata-Yanagimoto, E. Nakagami-Yamaguchi, T. Saito, K. Kumano, K. Yasutomo, S. Ogawa, M. Kurokawa, and S. Chiba
From the Cover: Coordinated regulation of transcription factors through Notch2 is an important mediator of mast cell fate
PNAS, June 3, 2008; 105(22): 7839 - 7844.
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 JEMHome page
M. L. Caton, M. R. Smith-Raska, and B. Reizis
Notch-RBP-J signaling controls the homeostasis of CD8- dendritic cells in the spleen
J. Exp. Med., July 9, 2007; 204(7): 1653 - 1664.
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 Mol. Cell. Biol.Home page
X. Ma, M. J. Renda, L. Wang, E.-c. Cheng, C. Niu, S. W. Morris, A. S. Chi, and D. S. Krause
Rbm15 Modulates Notch-Induced Transcriptional Activation and Affects Myeloid Differentiation
Mol. Cell. Biol., April 15, 2007; 27(8): 3056 - 3064.
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