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Notch/Delta4 Interaction in Human Embryonic Liver CD34⁺ CD38^– Cells: Positive Influence on BFU-E Production and LTC-IC Potential Maintenance

^a U362 Inserm, PR1, Institut Gustave Roussy, and
^b Department of Clinical Biology, Institut Gustave Roussy, Villejuif, France;
^c U506 Inserm, Hôpital Paul Brousse, Villejuif, France;
^d Children’s Hospital of Pittsburgh–Rangos Research Institute, Pittsburgh, PA, USA

Key Words. Hematopoietic stem cell • Erythropoiesis • Human embryo • Embryonic liver

Correspondence: E. Lauret, Ph.D., U362 Inserm, Institut Gustave Roussy, PR1, 94800 Villejuif, France. Telephone: 33-1-42-11-42-33; Fax: 33-1-42-11-52-40; e-mail: elauret{at}igr.fr

	ABSTRACT

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We investigated whether Notch signaling pathways have a role in human developmental hematopoiesis. In situ histochemistry analysis revealed that Notch1, 2, and 4 and Notch ligand (Delta1–4, and Jagged1) proteins were not expressed in the yolk sac blood islands, the para-aortic splanchnopleure, the hematopoietic aortic clusters, and at the early stages of embryonic liver hematopoiesis. Notch1–2, and Delta4 were eventually detected in the embryonic liver, from 34 until 38 days postconception. Fluorescence-activated cell sorter analysis showed that first-trimester embryonic liver CD34⁺CD38^low cells expressed both Notch1 and Notch2. When these cells were cultured on S17 stroma stably expressing Delta4, a 2.6-fold increase in BFU-E number was observed at day 7, as compared with cultures with control stroma, and this effect was maintained for 2 weeks. Importantly, exposure of these cells to Delta4 under these conditions maintained the original frequency and quality of long-term culture-initiating cells (LTC-ICs), while control cultures quickly resulted in the extinction of this LTC-IC potential. Furthermore, short-term exposure of embryonic liver adherent cells to erythropoietin resulted in a dose-dependent increase in Delta4 expression, almost doubling the expression observed with untreated stroma. This suggests that Delta4 has a role in the regulation of hematopoiesis after a hypoxic stress in the fetus.

	INTRODUCTION

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The hematopoietic system of higher vertebrates emerges in a series of finely controlled spatial and temporal events that occur through the sequential appearance and colonization of specific embryonic territories [1]. Hematopoietic cells are first detected in the yolk sac, defined as the site of primitive hematopoiesis, which is responsible for the preliminary wave of circulating blood cells but does not contain long-term repopulating hematopoietic stem cells (HSCs). Definitive hematopoiesis, which implies the stem cells that give rise to the hematopoietic system in the adult, appears in the embryo, within the aorta-gonads-mesonephros (AGM) region. We previously mapped the emergence of definitive hematopoiesis in the human embryo to the truncal part of the dorsal aorta and vitelline artery within the AGM, from 27–40 days postconception (dpc) [2–4]. From 31 dpc onward, hepatic hematopoiesis commences, which remains the major hematopoietic organ for the first trimester, an event which we have suggested to occur through the colonization of the liver by the AGM-derived HSCs.

Notch was first identified in Drosophila [5], in which it was demonstrated to specifically control wing development but was later determined to also influence the development of many other tissues [6]. Notch-mediated signal transduction represents a highly conserved series of events with implications in asymmetric cell division [7, 8], lateral inhibition [9, 10], and cell fate determination [11–13] in both developmental and adult processes.

As a result of the direct influence on cell fate, the hematopoietic system from both mice and humans has received extensive study to determine if activation of the Notchpathway can influence HSC fate. Although knockout studies have not shown a clear role for Notch in adult HSCs, experiments implying Notch activation show that Notch does modulate adult HSC fate. The constitutive expression of an active form of Notch1 in murine hematopoietic progenitors promotes HSC self-renewal [14, 15]. Notch ligands, Jagged1–2, and Delta1 inhibit differentiation of hematopoietic progenitors [16–19]. Furthermore, Jagged1 and Delta1 were shown to be growth factors for hematopoietic progenitors [20–23]. In contrast, we and others have documented a role for Delta4 and Jagged1 as negative regulators of the cell cycle [24, 25].

Previous work has argued a role for Notch signaling in the development of the murine hematopoietic system. Coculture of AGM or d11 fetal liver CD34⁺c-kit⁺ cells with S17 stroma expressing Jagged1 increased the committed colony-forming potential of the output cells [17]. Recent studies in mice implicate Notch1 in the generation of the earliest embryonic HSCs [26]. Splanchnopleural explant cultures from Notch1-deficient but not Notch2-deficient mice were markedly impaired in their ability to generate hematopoietic colonies.

In the study reported here, we addressed the role of the Notch signaling pathway in human hematopoietic development. Using, in parallel, immunohistochemistry and in vitro analysis of embryonic hematopoietic progenitors cocultured with Notch ligand–expressing stroma, we have documented a role for Notch activation during the first trimester of human embryonic hematopoietic emergence. Chronologically, Notch expression in the major hematopoietic sites was only detected after the establishment of definitive hematopoiesis, while functional studies of the Notch ligands detected by immunohistochemistry showed that Notch activation through Delta4 induces fetal CD34⁺CD38^– cells to generate erythroid progenitors, while maintaining their long-term culture-initiating cell (LTC-IC) potential, thus preventing progenitor cell exhaustion in human embryonic development.

	MATERIALS AND METHODS

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Human Tissues
Human embryos from 21–55 dpc (Table 1) were obtained after voluntary terminations of pregnancy induced with the RU 486 anti-progestative compound. Alternatively, embryos up to 10.5 weeks of development were obtained after voluntary termination of pregnancy by aspiration. In all cases, informed consent for use of embryonic tissue for research was obtained from the patient. All work on human tissues was performed according to the guidelines and with the approval of both our national (Comité National d’Ethique) and institutional (Comité Opérationnel pour L’ethique en Sciences) ethics committees. Gestational age was estimated according to previously described criteria [4].

Antibodies Used for Histochemical Analysis
Anti-Notch1, Notch2, and Notch4 antibodies specific for the extracellular-domain epitopes and anti-Delta1, intracellular-domain epitope–specific antibody (Santa Cruz Biotechnologies, Santa Cruz, CA, http://www.scbt.com) were used at 0.8 µg/ml, 0.15 µg/ml, 0.8 µg/ml, and 0.25 µg/ml, respectively. An anti-Jagged1 antibody recognizing an extracellular-domain epitope (R&D Systems, Minneapolis, http://www.rndsystems.com) was used at 2 µg/ml, and an anti-Delta4 antibody recognizing an intracellular-domain epitope (Millenium Inc., Cambridge, MA, http://www.mlnm.com) was used at 0.3 µg/ml. Anti-human CD34 antibodies (Immunotech, Marseille, France, http://www.sfrl.fr/qui/ad_immunotech.html) were used undiluted, and anti-human CD45 (Immunotech) was used at a 1/100 dilution. Secondary species-specific biotin-conjugated antibodies were anti-mouse at 1/500 (Immunotech), anti-goat at 10 µg/ml (Jackson Laboratory, Bar Harbor, ME, http://www.jax.org), and anti-rabbit at 1/400 (Immunotech). Negative-control stained samples were incubated with the secondary antibody only.

Cell Lines
S17 [27] and MS-5 [28] cell lines were grown in minimal essential medium (-MEM) supplemented with 10% fetal calf serum (FCS; Invitrogen, Carlsbad, CA, http://www.invitrogen.com).

Generation of S17 Stromal Cell Populations
Generation and characterization of the S17 stromal cell control (C/S17), and S17 stromal cell stably engineered to express the membrane-bound form of Delta4 (mb4/S17) have been previously described [25].

Cell Preparation and Labeling
Following isolation from the embryonic body by micro dissection, first-trimester human embryonic livers were quickly dissociated by pipetting the tissue repeatedly in a 250 U/ml concentration of collagenase I solution (Sigma Chemical Corp., St. Louis, http:// www.sigma-aldrich.com). The cell suspension was then immediately subjected to a density-gradient separation, and the mononuclear cells were collected and frozen in a solution mixture of 90% FCS and 10% dimethyl sulfoxide (DMSO) at –80°C using a step-down freezing device, then stored in nitrogen until required. Following thawing, sorting of the desired CD34⁺CD38^– population (defined as the 5% of the CD34⁺ cell population expressing low CD38) was performed using a FACSVantage flow cytometer (Becton, Dickinson, Franklin Lakes, NJ, http://www.bd.com), as previously described. CD34⁺CD38^– cell populations were isolated using anti-CD34 phycoerythrin (PE)-Cy5 (Beckman-Coulter, Miami, http://www.beckman.com) and anti-CD38 PE antibodies (Becton, Dickinson).

Analysis of Notch1 or Notch2 Expression in CD34⁺CD38^– Cells
Mononuclear cells isolated from embryonic livers were stained with anti-CD34 and anti-CD38 antibodies, as described above. In addition, each sample was stained with a rabbit immunoglobulin G (IgG) control, the anti-Notch1, or the anti-Notch 2 antibody, all at 4 µg/ml. After 1 hour on ice, the samples were washed and then stained with an anti-rabbit IgG-fluorescein isothiocyanate–conjugated antibody. After a further hour, the samples were washed and analyzed on a FACSort, equipped with an argon laser and a diode tuned to 485 nm and 605 nm, respectively, by gating on the CD34⁺CD38^– population and then cross analyzing with the third color.

Coculture Experiments
Cultures were initiated by plating 1,000 CD34⁺CD38^– cells in 24-well plates that had been precoated with C/S17, or mbD4/ S17, in 1 ml of -MEM containing 10% FCS, pegylated-recombinant human (rhu)–megakaryocyte growth and development factor (MGDF; 50 ng/ml; Kirin Brewery, Tokyo, http://www.kirin.co.jp/english), rhu–interleukin-3 (IL-3; 100 U/ml; Novartis International, Basel, Switzerland, http://www.novartis.com), rhu–FMS-like tyrosine kinase 3 (FLT-3) ligand (100 ng/ml; Amgen, Thousand Oaks, CA, http://www.amgen.com), and rhu–stem cell factor (SCF; 50 ng/ml; Amgen). At day 7, non-adherent cells were harvested, and the CD34⁺ mononuclear cell fraction was sorted. This enabled us to determine the percentage of output CD34⁺ cells in the coculture. Purified CD34⁺ cells were analyzed for colony-forming cell (CFC) and LTC-IC potential, while 1,000 output CD34⁺ cells were replated onto fresh stromal layers in the cytokine-rich medium for a further week. This procedure was performed over 2 weeks. Some cocultures were performed in the presence of 10 µM DAPT (a gamma secretase inhibitor: N-[N-(3,5-difluorophenacetyl)-L-alanyl]-(S)-phenylglycine t-butyl ester; Calbiochem, San Diego, http://www.bioresearchonline.com/storefronts/calbiochem.html) or DMSO as control. At day 7, cell populations were analyzed for their BFU-E potential.

Clonal analysis was performed by depositing one CD34⁺CD38^– cell per well on either control or Delta4-expressing S17 stroma in 96-well plates in the presence of cytokines for 1 week, followed by plating in methylcellulose.

Clonogenic Progenitor Assays
Quantification of CFCs was performed using standard methylcellulose colony assays following previously described criteria [29].

Assessment of LTC-IC Potential
Input CD34⁺CD38^– cells from fresh tissue samples, or output CD34⁺ cells from the coculture with control or Notch ligand–expressing stroma, were transferred to 96-well plates precoated with MS-5 cells and incubated in standard LTC medium (-MEM containing 12.5% horse serum, 12.5% FCS, 10^–4 mol/L 2-ß-mercaptoethanol), at a limiting dilution. Plates were incubated at 33°C, in air atmosphere with 5% CO₂, with weekly medium demidepletion as previously described. After 5 weeks in culture, wells were trypsinized and then plated in methylcellulose in clonogenic progenitor assay to determine both frequency and quality of the LTC-IC population for each condition. Frequency was determined by the Poisson distribution, and the LTC-IC quality was determined by their individual proliferative capacity (i.e., CFC number per one LTC-IC).

Embryonic Liver Adherent Cell Analysis
CD34⁺CD45^– cells were purified from the first-trimester human embryonic liver and plated into 24-well plates precoated with 0.2% w/v gelatin, in endothelial cell growth medium (BioWhittaker Molecular Applications, Rockland, ME, http://www.bmaproducts.com). After 3 days, the wells received a medium exchange, which was performed weekly until a clearly visible adherent layer had formed. Cells were then transferred and expanded in -MEM containing 10% v/v FCS for further experimentation. For erythropoietin (EPO) sensitivity experiments, 100,000 adherent cells seeded 24 hours previously were incubated with -MEM containing 10% v/v FCS, supplemented with human EPO or vascular endothelial growth factor (VEGF; R&D Systems, Minneapolis, http://www.rndsystems.com) at increasing concentrations. After 2–2.5 hours, the cells were trypsinized, fixed, permeabilized in Ortheapermafix reagent (Ortho-Clinical Diagnostics, Raritan, NJ, http://orthoclinical.com), and stained with antibodies specific for the intracellular domains of Delta1, Delta4, or a rabbit polyclonal IgG isotype control (Sigma), all at 4 µg/ml. Sequentially, the samples were stained with an anti-rabbit, PE-conjugated antibody at 5 µg/ml (SBA Products, Birmingham, AL, http://www.southernbiotech.com) and analyzed for both forms of Delta expression on a FACSort (Becton, Dickinson). Oxygen sensitivity experiments were performed in a similar mode, except the cells were incubated in either 21% or 7% O₂ humidified incubators for 2–2.5 hours prior to analysis.

Statistical Analysis
Statistical analysis was performed using Student’s t-test (equal variant analysis); data were considered significant at p < .05.

	RESULTS

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Detection After Commencement of Definitive Hematopoiesis in the Embryonic Liver
We first focused on in situ analysis of Notch and Notch ligand protein expression (Notch1, Notch2, and Notch4; Delta1 and Delta4; Jagged1) to the preliminary sites of hematopoietic development. Extraembryonic blood islands, which appear in the human yolk sac from 16–17 dpc and give rise to the first wave of blood cell circulation, did not express any of the Notch or Notch ligands tested (data not shown). We analyzed extraembryonic tissues from 20–25 dpc (n = 6), representing Carnegie stages 9 to 11 (Table 1), and never detected expression in the blood islands. Furthermore, no expression of Notch or Notch ligands was detected anywhere within the embryos analyzed.

From 27–40 dpc, hematopoiesis switches to the embryo proper on the ventral wall of the dorsal aorta in the AGM region. The CD34⁺CD45⁺ cell clusters found here also did not express any of the Notch and Notch ligands tested, although a weak Jagged1 expression encircling the aorta was observed, while rare cells expressing Notch1 were found both anterior and posterior to the aorta (data not shown).

We next switched our focus to the embryonic liver, which represents the major hematopoietic organ within the developing embryo and fetus in the first trimester of development. The onset of CD34⁺ cell-mediated hematopoiesis commences here at approximately 30 dpc, and we were only able to detect Notch1, Notch2, and Delta4 expression from 34 dpc onward, and then only at a low frequency. No Notch or Notch ligand expression was detected in the liver prior to this time. We continued to observe the rare expression of these proteins for the remainder of the gestational period analyzed (Table 1). In one embryo at 36 dpc, in addition to Notch1, Notch2, and Delta4, we observed the rare appearance of cells expressing Delta1 (Fig. 1). In all cases, the major proteins expressed were Notch1, Notch2, and Delta4, all at low frequency, which prevented us from comparing sequential tissue sections and from clearly determining whether the positive cells were also expressing CD34 or CD45 or were part of the liver vasculature. It is noteworthy that at no time in any of the tested hematopoietic tissues did we observe expression of the endothelium-associated Notch4 protein or Jagged1. However, we could rarely detect Notch4 in other vascular tissues, while Jagged1 was highly expressed in the neural tube, mesonephros, and hepatic ductal plate and rarely in the vasculature (data not shown).

View larger version (164K): [in this window] [in a new window]   Figure 1. Notch and Notch ligand expression in the embryonic human liver. The 36-day liver sections are shown, stained with antibodies to (A) CD34, (B) CD45, (C) Notch1, (D) Notch2, (E) Notch4, (F) Jagged1, (G) Delta1, (H) Delta4. Scale bar: 25 µm.

View larger version (18K): [in this window] [in a new window]   Figure 2. Notch1–2 protein expression on embryonic liver CD34+CD38– cells. Mononuclear cells from first-trimester embryonic liver were stained for CD34, CD38, and Notch molecules. Following gating on the CD34+CD38– population, cells were analyzed for coexpression of CD34 and Notch1 or Notch2.

The role of Delta4 on HSCs was analyzed as follows. CD34⁺CD38^– mononuclear cells obtained from human embryonic liver at 6.5–9.5 weeks of gestation were cultured with either S17 stroma stably transfected with an empty vector (C/S17) or S17 stroma stably transfected with a construct coding for the membrane-bound form of Delta4 (mb4/S17), in a cytokine-rich milieu known to support the proliferation of primitive HSCs, for 7 days. Output CD34⁺ mononuclear cells were then purified by cell sorting, analyzed for hematopoietic characteristics, or replated onto fresh stromal layers for a further 7 days.

Following culture of 1,000 CD34⁺CD38^– cells with either the C/S17 or mb4/S17 stroma (n = 12), no significant differences were observed in the number of output CD34⁺ mononuclear cells at weeks 1 and 2 (28 ± 23 x 10³ versus 12 ± 7 x 10³ for C/S17 and 11 ± 7 x 10³ versus 8 ± 4 x 10³ for mb4/S17), as well as the percentage of CD34⁺ cells at weeks 1 and 2 (16% ± 12% versus 15% ± 11% for C/S17 and 10% ± 5% versus 9% ± 6% for mb4/S17). In contrast, clonogenicity of the output CD34⁺ cells revealed that mb4/S17 significantly augmented the total colony number per 1,000 output CD34⁺ cells at week 1 (104 ± 68 colonies for mb4/S17 versus 68 ± 31 for C/S17; p = .03) (Fig. 3A). Dissection of the colony type generated revealed that the C/S17 and mb4/S17 stromas generated equivalent numbers of nonerythroid myeloid colonies (67 ± 18 colonies for mb4/S17 versus 59 ± 17 for C/S17) (Fig. 3B). More striking was the effect of mb4/S17 on the erythroid cell–forming capacity of CD34⁺ cells. After 7 days of culture with mb4/S17, output CD34⁺ cells gave rise to 35% of BFU-E (36 ± 10 colonies for mb4/S17 versus 14 ± 12 for C/S17 per 1,000 output CD34⁺ cells; p = .001) (Fig. 3C). A similar significantly higher number of BFU-E was observed with mb4/S17 at week 2 (16 ± 7 colonies for mb4/ S17 versus 4 ± 3 for C/S17 per 1,000 output CD34⁺ cells; p = .001) (Fig. 3C). No differences in myeloid colony type were detected at week 2 of culture between mb4/S17 and C/S17 (Fig. 3B).

View larger version (21K): [in this window] [in a new window]   Figure 3. Effect of Delta4 on hematopoietic progenitors. Following coculture of CD34+CD38– mononuclear cells with either control (white) or Delta4 (gray) stroma, output CD34+ cells were assessed for their direct colony-forming capacity. (A): Frequency of total colonies. (B): Frequency of nonerythroid colonies. (C): Frequency of erythroid colonies, all at weeks 1 and 2. (D): Following coculture of CD34+CD38– cells with either control (white) or Delta4 (gray) stroma, in the presence of 10 µM DAPT (a secretase inhibitor: N-[N-(3,5-difluorophenacetyl)-L-alanyl]-(S)-phenylglycine t-butyl ester) or dimethyl sulfoxide (DMSO) (as control) for 1 week, output nucleated cells were assessed for their BFU-E content. Asterisks indicate statistical significance: *p < .05; **p < .001.

The CD34⁺CD38^– cell population, though representing purified primitive cells, possesses significant amounts of functional diversity. It therefore became important to determine whether Delta4 was expanding progenitors already committed toward the erythroid lineage or favoring differentiation toward erythropoiesis. To address this question, CD34⁺CD38^– cells were seeded at 1 cell per well on either the C/S17 or mb4/S17 stroma in the presence of cytokines. After 1 week, the total cells were assessed for their erythroid colony-forming potential. Both the C/S17 and mb4/S17 stroma conditions yielded equivalent frequencies of wells that could generate colonies (16.2% ± 2.8% and 17.8% ± 1.1%, respectively; n = 4) (Table 3), while the frequency of wells containing erythroid colonies was augmented in the mb4/S17 condition (42% ± 7% for mb4/S17 compared with 31% ± 5% for C/S17). No differences were observed in the frequency of wells giving rise to myeloid colonies. Additionally, the fact that no well gave rise to exclusive erythroid colonies and that the number of erythroid colonies generated per single cell was almost identical

Delta4 and LTC-IC Frequency
In parallel to the study on committed progenitors, we also examined the effect of Delta4 on more primitive progenitors, as measured using the in vitro LTC-IC assay. Following the initial 7-day coculture period with either the control or the Delta4 stroma, the output CD34⁺ cells were plated on MS-5 for 5 weeks and then cultured in methylcellulose to analyze the LTC-IC–derived CFCs. The frequency of LTC-ICs in output CD34⁺ cells was maintained following mb4/S17 coculture (Fig. 4) at a level similar to the one of input CD34⁺CD38^– cells (frequency of 0.04 versus 0.01, respectively). In contrast, coculture with the C/S17 stroma led to a rapid and significant decrease in the LTC-IC frequency (0.0043), which was 10-fold lower than the LTC-IC frequency of output CD34⁺ cells exposed to mb4/S17 (p = .03) and threefold lower than the LTC-IC frequency of input CD34⁺CD38^– cells (p = .0003). No differences in the number of clonogenic progenitors, either total or BFU-E, generated per LTC-IC were observed between the mb4/S17 and C/S17 conditions. The decrease in the LTC-IC frequency of output CD34⁺ cells continued into the second week of coculture, dropping below the sensitivity limit of the assay following C/S17 coculture (p<.0002), and being only just detectable following mb4/S17 coculture (Fig. 4). At no time point tested did the output CD34^– mononuclear cell population generate detectable LTC-ICs (data not shown).

View larger version (12K): [in this window] [in a new window]   Figure 4. Effect of Delta4 on long-term culture-initiating cell (LTC-IC) potential. Following coculture of CD34+CD38– mononuclear cells with either control (white) or Delta4 (gray) stroma, output CD34+ cells were assessed for their primitive LTC-IC characteristics: LTC-IC frequency of input CD34+CD38– cells and LTC-IC frequency of output CD34+ cells at week 1 or week 2. Results are shown as the mean ± SE from six embryonic livers. Asterisks indicate statistical significance: *p < .05.

View larger version (32K): [in this window] [in a new window]   Figure 5. Effect of hypoxia-associated growth factors on Delta expression on embryonic liver–adherent cells. Embryonic liver–adherent cells were exposed to increasing concentrations of either erythropoietin (EPO) or vascular endothelial growth factor (VEGF). Delta expression was measured after a continuous 2.5-hour growth factor exposure. (A): Delta4 expression of embryonic liver–adherent cells exposed to increasing concentrations of EPO in one representative experiment. (B): (a) Delta4 expression following EPO exposure; (b) Delta1 expression following EPO exposure; (c) Delta4 expression following VEGF exposure; (d) Delta1 expression following VEGF exposure. The mean result from five independent liver samples is shown ± SE.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

In contrast to the murine model [26] was the absence of Notch- or Delta-type ligand protein expression in the AGM region and the surrounding aorta-associated hematopoietic cell clusters. Jagged1 RNA expression has still been detected around the aorta in the human AGM region [39], and we also could find Jagged1 expression by immunohistochemistry, though the protein signal was weak, especially in comparison with the high Jagged1 expression in the neural tube in the same embryos (data not shown). With specific regard to the human hematopoietic development, Notch and Notch ligand proteins were only detected after the initiation of definitive hematopoiesis in the liver after 34 days. Continued protein expression of both Notch and Notch ligands was observed throughout the first trimester, although the frequency of positive cells was low.

Because Delta4 was expressed in the liver from 34–38 dpc, we examined its potential role in the regulation of fetal liver HSCs. Interestingly, culture of fetal liver CD34⁺CD38^– cells on S17 stroma stably expressing Delta4 did not result, as previously demonstrated with adult HSCs, in a reduction of cell proliferation. Indeed, human CD34⁺CD38^low cells (harvested from cord blood, bone marrow, or cytapheresis) exposed to membrane-bound Delta4 or an immobilized purified Delta4-Fc protein proliferated less, a higher proportion of cells being maintained in G0/G1 phase [25]. This discrepancy can be due to the more active cell cycle status of embryonic cells.

Nevertheless, fetal liver HSCs shared several responses to Delta4 with adult HSCs. One of the striking observations was the counteracting in the loss of the LTC-IC potential induced by cytokines in output fetal CD34⁺ cells. Importantly, exposure of fetal CD34⁺CD38^– cells to Delta4 during 7 days maintained the original frequency and high proliferative capacity of LTC-ICs, while culturing with control stroma quickly resulted in the extinction of this LTC-IC potential. These observations are in keeping with the activity of Delta4 on adult HSCs. Indeed, whatever its mode of presentation, as membrane bound or as purified protein immobilized on the plastic, Delta4 was demonstrated to rescue the loss of the LTC-IC potential in adult CD34⁺CD38^low cells, induced by cytokines. Such delay in cell differentiation has been previously documented in a report describing that overexpression of mbDelta-4 in murine HSCs prevents their reconstitutive ability by blocking their differentiation into pre-CFU-spleen [40]. Interestingly, the fact that Delta4 does not alter cell proliferation of fetal CD34⁺ cells allowed us to conclude that Delta4 directly counteracts cell differentiation events triggered by cytokines, independent of the mitotic history, and thus promotes self-renewing divisions. Such a conclusion could not be drawn with adult HSCs, since it was difficult to determine whether Delta4 was maintaining the LTC-IC potential by a reduction in cell proliferation or by alternative mechanisms independent of the mitotic history. All these observations are in keeping with the general property that Notch activation favors HSC self-renewal in both mice and humans [41].

Besides its activity in maintaining the LTC-IC potential of fetal CD34⁺ cells, Delta4 appeared as an actor in the generation of erythroid progenitors. Culture of fetal CD34⁺CD38^– cells with Delta4 led to a 2.6-fold increase in the number of BFU-E observed at day 7, as compared with the fresh sample, an effect which was maintained for at least 2 weeks. This increased production of BFU-E was the result of an enhanced capacity of single CD34⁺ CD38^– cells to generate BFU-E. This effect, previously observed when cord blood CD34⁺ CD38^low cells were cultured in the presence of Delta4 [25], seems specific for Delta4 since Delta1 did not elicit such activity on BFU-E production (data not shown). Nevertheless, previous work has documented such a role for Jagged1 when HSCs were cultured in the presence of SCF alone [24].

The clear push toward erythrogenesis of fetal liver progenitors following culture with Delta4 and the expression of Delta4 in the embryonic liver, combined with the fact that Delta4 mRNA was upregulated in endothelial cells following exposure to extreme basal oxygen concentrations [42], prompted us to investigate a role of Delta4 in hypoxia during early development. Since standard responses to lowering oxygen tension include increases in the levels of EPO and VEGF, we addressed the question whether such cytokines could influence Delta4 expression on embryonic stroma cells. A 2-hour exposure of high concentrations of EPO to adherent embryonic liver cells resulted in a rapid upregulation of Delta4 protein expression, which quickly returned to baseline values when EPO was removed. Our data suggest that at least with specific regard to Delta4 protein expression, regulation appears more likely at the translational or post-translational level.

The rare expression of Delta4 in human embryos, with an apparent restriction to the liver, and the significant induction of erythropoiesis with the maintenance of the primitive potential upon activation of HSCs by it, suggest an interesting model for adaptation to oxygen tension. In specific locations, particularly at times of rapid growth like the embryo, which has been argued to be under an almost constant state of hypoxia [43, 44], standard responses to lowering oxygen tension may not be sufficient. Furthermore, the hypoxic state of the embryo has been associated with augmented hematopoiesis in both murine and human systems [45–47]. With specific regard to Delta4, if an acute hypoxic state persists, this could result in elevated local levels of EPO, which would directly induce the surrounding cells to express Delta4. In turn, HSCs, once activated by Delta4, would be programmed toward the erythroblast lineage to alleviate the hypoxia but also to maintain their primitive potential, ensuring, at least in the short term, that the HSC pool not be exhausted. Upon return to a more amenable oxygen tension, Delta4 expression would no longer be induced.

In conclusion, this study supports previous demonstrations that Notch-mediated control of cell fate is an induced and, similar to many developmental signals, transient signaling event [48], which is sufficient to address the immediate needs of the growing organs. In turn, this may offer new approaches for tissue engineering when attempting to induce multicellular differentiation from a single stem cell.

	ACKNOWLEDGMENTS

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We are indebted to Amgen for providing us with rhu-SCF and Flt3-ligand; Kirin for providing us with rhu-PEG-MDGF; Novartis for providing us with rhu-IL-3; Cilag AG (Schaffhausen, Switzerland, http://www.cilag.ch) for providing us with rHuEPO; and Dr. K. Mori (Niigata University, Niigata, Japan) for providing us with the MS-5 cell line. We thank Y. Lécluse and F. Larbret for performing the cell sorting. This work was supported by Inserm ("Poste Vert") and grants from CRC (Contrat de Recherche Clinique, no. 2000.10, Institut Gustave Roussy), ARC (Association de Recherche contre le Cancer, no. 4300), and HFSP (Human Frontier Science Program, no. RG0345/1999-M).

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Top Abstract Introduction Materials and Methods Results Discussion References

 

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