Loss of Hmga1 gene function affects embryonic stem cell ... - CiteSeerX

The FASEB Journal express article 10.1096/fj.02-0977fje. Published online June 17, 2003.

Loss of Hmga1 gene function affects embryonic stem cell lymphohematopoietic differentiation Sabrina Battista,*,† Francesca Pentimalli,* Gustavo Baldassarre,† Monica Fedele,* Vincenzo Fidanza,† Carlo M. Croce,† and Alfredo Fusco* *Dipartimento di Biologia e Patologia Cellulare e Molecolare, c/o Centro di Endocrinologia ed Oncologia Sperimentale del CNR, Facoltà di Medicina e Chirurgia, Università degli Studi di Napoli “Federico II,” Naples, Italy; and †Kimmel Cancer Institute, Jefferson Medical College, Philadelphia 19107, Pennsylvania, USA Corresponding author: Alfredo Fusco, Dipartimento di Biologia e Patologia Cellulare e Molecolare, c/o Centro di Endocrinologia ed Oncologia Sperimentale del CNR, Facoltà di Medicina e Chirurgia, Università degli Studi di Napoli “Federico II,” via Pansini 5, 80131 Naples, Italy. E-mail: [email protected] ABSTRACT By interacting with transcription machinery, high-mobility group A 1 (HMGA1) proteins alter the chromatin structure and thereby regulate the transcriptional activity of several genes. To assess their role in development, we studied the in vitro differentiation of embryonic stem (ES) cells that bear one or both disrupted Hmga1 alleles. Here, we report that Hmga1 null ES cells generate fewer T-cell precursors than do wild-type ES cells. Indeed, they preferentially differentiate to B cells, probably consequent to decreased interleukin 2 expression and increased interleukin 6 expression. Moreover, a lack of HMGA1 expression induces changes in hemopoietic differentiation, i.e., a reduced monocyte/macrophage population and an increase in megakaryocyte precursor numbers, erythropoiesis, and globin gene expression. Re-expression of the Hmga1 gene in Hmga1 null ES cells restores the wild-type phenotype. The effect on megakaryocyte/erythrocyte lineages seems, at least in part, mediated by the GATA-1 transcription factor, a key regulator of red blood cell differentiation. In fact, we found that Hmga1−/− ES cells overexpress GATA-1 and that HMGA1 proteins directly control GATA-1 transcription. Taken together, these data indicate that HMGA1 proteins play a prime role in lymphohematopoietic differentiation. Key words: HMGA1 • homologous recombination • T lymphocytes • interleukin • ES cells The mammalian high-mobility group A (HMGA) family of chromosomal proteins includes HMGA1a and HMGA1b, which are encoded by Hmga1 through alternative splicing (1), and the closely related HMGA2 protein (2). These proteins bind the minor groove of AT-rich DNA sequences. Their DNA binding domain is located in the N-terminal region of the protein and contains three short basic repeats, the so-called AT hooks. HMGA proteins are involved in the regulation of chromatin structure and function (3). Although they do not have transcriptional activity per se, they regulate the transcriptional activity of an increasing number of genes, both

positively and negatively, by interacting with the basal transcription machinery. Hence, the name architectural transcription factors (4). Rearrangements of the Hmga1 gene have been frequently described in human benign tumors of mesenchymal origin (5), and Hmga1 gene expression is a constant feature of human tumors (see ref 6 for review). However, HMGA proteins seem to exert their major physiological role during embryonic development (7, 8). In fact, Hmga1 gene expression is quite high during embryogenesis, whereas it is negligible in normal adult tissues. To identify the differentiative pathways in which HMGA1 is involved, we produced embryonic stem (ES) cells carrying a disrupted Hmga1 gene. ES cells can be kept in an undifferentiated state by treatment with leukemia inhibiting factor (LIF) or can be induced to differentiate into specific cell types via specific factors (9, 10). In this system, gene expression can be studied throughout differentiation, which in addition reflects the in vivo temporal pattern. Differentiation of ES cells in semisolid methylcellulose-based medium has been used to evaluate the ability of these cells to produce lymphohematopoietic lineages (10). During differentiation of the hematopoietic system, progeny of undifferentiated pluripotent stem cells become committed to one of several developmental pathways and thereby give rise to precursors and mature cell types of a particular hemopoietic lineage (11). Here, we compared the in vitro differentiation of wild-type ES cells with Hmga1+/− and Hmga1−/− ES cells: knockout of the Hmga1 gene drastically impaired monocyte/macrophage lineages and differentiation of multilineage hemopoietic progenitors. In addition, Hmga1 null ES cells showed a reduced ability to generate T-cell precursors compared with wild-type ES cells, whereas they preferentially generated B-lymphocyte precursors. Moreover, Hmga1−/− ES cells showed increased megakaryocyte/erythrocyte differentiation as well as globin gene expression. We showed that HMGA1 proteins directly bound the erythroid-specific transcription factor GATA-1 gene regulatory region and down-regulated its expression, which provides a possible mechanism of action on hematopoietic differentiation. MATERIALS AND METHODS Vectors Several overlapping clones were isolated from a λφXII phage library of a 129/SVJ mouse strain (Stratagene, La Jolla, CA) by using standard procedures. Restriction mapping and partial sequence analyses were used to identify exon-intron boundaries. Genomic sequences encompassing exons V, VI, VII, VIII, and IX were subcloned. The fifth (containing ATG) and sixth exons were substituted by an HindIII-EcoRI fragment containing a neocassette derived from the PGK-neoplasmid. To rescue the Hmga1 gene, the murine full-length Hmga1b cDNA was subcloned into pcDNA3.1/Hygro(+) expression vector (Invitrogen, Carlsbad, CA). The same construct was also used for transient transfections. The reporter constructs, containing the GATA-1 upstream regulating sequence IE2.6intLUC and IE3.9intLUC, have been described (12) and were a gift from Dr. M. Yamamoto.

ES cell culture and transfections ES cells AB2.1 are described elsewhere (13). Twenty micrograms of the targeting construct were linearized with PacI and electroporated into ES AB2.1 cells (1 × 107) at 0.23 kV, 500 µF. ES cells were selected 24 h later with Geneticin G418 (180 µg/ml). Nine hundred clones were screened by Southern blotting as described below, and three heterozygous clones were identified and expanded. The heterozygous clones were plated at a concentration of 1 × 106 cells/plate and selected with a high concentration (6 mg/ml) of Geneticin G418 to obtain double-knockout cells. About 20 clones were obtained and screened by Southern blotting. For the rescue, 20 µg of plasmids (pc-Hmga1/Hygro or the empty vector) were electroporated into two Hmga1−/− ES cell clones. Cells were selected 24 h later with 0.25 mg/ml hygromycin. Transgene expression was detected by Northern blotting and reverse transcriptase-polymerase chain reaction (RTPCR). Southern blot analyses Genomic DNA from Geneticin G418-resistant ES cells was digested with EcoRI and underwent electrophoresis on 0.7% agarose gel. The probe used for Southern blotting hybridization was a 550-bp fragment obtained by PstI digestion of the Xba-EcoRI fragment and 3' to the stretch included in the recombination construct. Differentiation of ES cells by retinoic acid ES cells were differentiated as described elsewhere (9). Before differentiation, feeder fibroblasts were eliminated by three passages on gelatin-coated plates, and LIF (103 U/ml) was added to the medium to keep the cells undifferentiated. Cells (1 × 106) were plated on bacterial-grade plates and kept for 4 days in suspension cultures. During the next 4 days, 5 × 10−7 M retinoic acid was added to the medium, after which embryoid bodies (EBs) were plated on cell culture dishes and incubated without retinoic acid for 19 days. Generation of EBs from ES cells in methylcellulose-based medium ES cells were differentiated essentially as described before (10). Forty-eight hours before differentiation, 2 × 105 ES cells were plated on gelatin-coated plates in Iscove’s Modified Dulbecco’s Medium (IMDM) medium supplemented with 15% FBS, sodium pyruvate (1 mM), L-glutamine (2 mM), and nonessential amino acids (0.1 mM) (GIBCO BRL Life Technologies, Gaithersburg, MD); monothioglycerol (MTG; 100 µM, Sigma-Aldrich, St. Louis, MO); LIF (10 ng/ml, Chemicon, Temecula, CA); penicillin G and streptomycin. Materials for differentiation were purchased from Stem Cell Technologies (Vancouver, BC, Canada) unless specified otherwise. To obtain EBs, 2 × 103 ES cells were plated on low-adherence 35-mm Petri dishes as a single-cell suspension in primary differentiation medium, constituted by 0.9% methylcellulose in IMDM medium, 15% FBS (Stem Cell Technologies), L-glutamine (2 mM), MTG (150 µM), and murine stem cell factor (mSCF: 40 ng/µl). Feeding medium (0.5% primary differentiation medium, 15% FBS, 150 µM MTG, mSCF 160 ng/ml) was added after 7 days of culture, and then every 3–4 days. Hematopoietic differentiation

within EBs was assessed according to the presence of hemoglobinized cells and/or the appearance of evading macrophages. Generation of hematopoietic colonies by a “two-step” differentiation method For hematopoietic differentiation, 11-day-old EBs were disrupted by collagenase (Stem Cell Technologies) according to the manufacturer's instructions. Cells (5 × 104) were plated in methylcellulose cultures (in 35-mm low-adherence Petri dishes), as described above, except that additional growth factors (1% bovine serum albumin, 10 µg/ml insulin, 200 µg/ml transferrin, 160 ng/ml mSCF, 30 ng/ml interleukin [IL]-3, 30 ng/ml human IL-6, and 3 U/ml erythropoietin) were added. Hematopoietic colonies were scored microscopically after 11 and 19 days, and colony types were determined by assessment of morphology. Megakaryocyte differentiation and identification ES-derived megakaryocyte differentiation was induced with the MegaCult-C kit (Stem Cell Technologies), according to the manufacturer's instructions. EBs grown in methylcellulose-based medium for 11 days were disrupted by collagenase and replated in collagen-based medium with recombinant human thrombopoietin (50 ng/ml), IL-3 (10 ng/ml), IL-6 (20 ng/ml), and IL-11 (50 ng/ml). Colony-forming unit megakaryocytes (CFU-Mks) were detected by staining the dehydrated and acetone-fixed colonies with a 0.1 M sodium phosphate buffer solution of acetylthiocholiniodide, sodium citrate, copper sulfate, and potassium ferricyanide to detect acetylcholinesterase activity. FACS analyses FACS analyses were performed as described previously (14). EBs were generated in methylcellulose-based medium, as described above. After 25 days, EBs were disrupted by collagenase, according to the manufacturer’s instructions. After washing once in PBS, aliquots of 1 × 106 cells were placed in 96-well, round bottom plates, incubated for 30 min at room temperature with normal rat and goat IgG (Santa Cruz Biotechnology, Santa Cruz, CA), and then incubated for 1 h at 4°C with conjugated monoclonal antibodies: anti-mouse CD44 (Ly-1) (phycoerythrin [PE] conjugated, clone IM7), anti-mouse CD117 (c-kit) (PE conjugated, clone ACK45), anti-mouse CD90.2 (Thy-1.2) (fluorescein isothiocyanate [FITC] conjugated, clone 532.1), and anti-mouse B220 (CD45R) (FITC conjugated, clone 145-2c11). All monoclonal antibodies were from PharMingen (San Diego, CA). After two washes in PBS, the cells were fixed in 1% paraformaldehyde, washed in PBS, and analyzed by flow cytometry (Coulter Counter, Becton, Dickinson, San Jose, CA) with CellQuest software (Becton, Dickinson). Viable cells were gated on the basis of forward and side scatter characteristics. A total of 40,000 gated events were analyzed for each sample. Generation of single- and double-knockout mice Several different Hmga1+/− ES cell clones were microinjected into C57BL/6J blastocysts at 3.5 days postcoitus (dpc) and were reimplanted into foster mothers (Animal Facility at Thomas Jefferson University). Chimeric mice were crossed to wild-type mice, and some of them gave germ line transmission. Single-knockout mice were then intercrossed to obtain double-knockout

mice. Pregnant mothers were killed at 9.5 and 14.5 dpc, and embryo genotype and viability were evaluated. RT-PCR analyses of embryos and ES cell cultures Tissues from mouse embryos were rapidly dissected, frozen on dry ice, and stored at –80°C. Total RNA from embryos and cell culture was extracted with TRI-reagent solution (Molecular Research Center, Cincinnati, OH), according to the manufacturer’s protocol, and treated with DNase I (GenHunter, Nashville, TN). One microgram of RNA was reverse-transcribed by using a mix of poly(dT) and random exonucleotides as primers and MuLV reverse transcriptase (PerkinElmer, Boston, MA). PCR amplification was performed as described before (15) in a GeneAmp PCR System 9600, using the primers listed in Table 1. Also non-reverse-transcribed RNA was amplified (data not shown). PCR products were separated on 2% agarose gel and, if necessary, blotted and hybridized with specific probes. Transient transfections Wild-type or double-knockout ES cells (4 × 105) were plated in six-well plates and were transfected at 48 h after plating with 1 µg of luciferase (LUC) reporter plasmid (IE2.6intLUC, IE3.9intLUC, or pGL3), by FuGene6 (Roche, Indianapolis, IN). When indicated, 3 µg of pcDNA3.1/hygro(+)-Hmga1b was cotransfected. Cells were harvested 48 h after transfection and lysates were analyzed for luciferase activity. Transfection efficiency was normalized by using the β-galactosidase activity. All assays were performed in triplicate and were repeated in three independent experiments. Formaldehyde cross-linking and chromatin immunoprecipitation Subconfluent plates of wild-type and Hmga1−/− ES cells were cross-linked via formaldehyde, and chromatin immunoprecipitation experiments were carried out as described previously (21). For immunoprecipitation, chromatin was mixed with anti-HMGA1, anti-HA, or normal rabbit IgG and incubated with rotation for 2 h at 4°C. After incubation with protein A/G-Sepharose beads (Santa Cruz Biotechnology), the beads were harvested by centrifugation and then washed sequentially as described (21). Formaldehyde cross-links were reversed, and the DNA was precipitated with ethanol, resuspended in water, and treated with RNase A (50 µg/ml) followed by proteinase K treatment (100 µg/ml). DNAs were extracted with phenol-chloroform and chloroform, precipitated with ethanol, and dissolved in water. Input DNA and immunoprecipitated DNAs were analyzed by PCR for the presence of Gata-1 promoter sequences. PCR analysis of immunoprecipitated DNA PCR reactions were performed with AmpliTaq Gold DNA Polymerase (PerkinElmer). The primers used to amplify the sequence of the GATA-1 upstream activating element, spanning nucleotides from −3230 to −2969 and indicated as region A (GenBank accession number AF136573), were (forward) 5'-AGTGGCAACAGATCACAAAG-3' and (reverse) 5'AACCACCTCCAATTCTCACC-3'. The primers amplifying the element between the upstream

activating element and the erythroid promoter (from −2001 to −1719, indicated as region B) were (forward) 5'TGAGAAGTCGCTGTTCACAG-3' and (reverse) 5'-GAGGCTTGTCCACTTACTGA-3'. PCR products were resolved on 2% agarose gels and transferred to nylon membrane. Regions A and B were amplified by PCR from the 3.9IEint-luc plasmid (12) and were used as probes for the blotted PCR products. Band shift assays Production of the recombinant HMGA1-His protein is described elsewhere (14). Undifferentiated wild-type and Hmga1−/− ES cells were lysed in NP-40 lysis buffer, supplemented with protease-inhibitor cocktail according to standard procedures. DNA binding assays were performed as reported by Thanos and Maniatis (4). The oligonucleotides used were GATA-1970 and mutant oligonucleotides (M1–M6), from −1970 to −1924 in the 5' GATA-1 upstream sequence, depicted in Fig. 8B; IL2CD28RE (22), and the unrelated sequence Pm (5'TGTCTTCCTTCCTATTCCTAAGAAATA). Poly(dGdC) was used as a specific competitor. A 100-fold excess of specific or nonspecific unlabeled competitor oligonucleotides was added in some experiments. For supershift, protein extracts were incubated with 1 µl of anti-HMGA1 or antiPit1 antibodies (Santa Cruz Biotechnology) for 4 h on ice. The DNA-protein complexes were resolved on 6% nondenaturing acrylamide gels and visualized by exposure to autoradiographic film. RESULTS Generation of Hmga1−/− ES cells To inactivate the Hmga1 gene, we cloned it from the 129/SVJ mouse strain and constructed a targeting vector in which exons V and VI were replaced with a neocassette. This vector was transfected into AB.2.1 ES cells, and two clones with a single-knockout Hmga1 allele were obtained and expanded. Several Hmga1−/− clones were generated by selecting the Hmga1 +/− ES cells with higher doses of Geneticin G418 (see Materials and Methods). The targeting of both Hmga1 alleles was demonstrated by Southern blot analysis (not shown). Northern blot analysis confirmed the reduction and absence of Hmga1-specific transcripts in Hmga1+/− and Hmga1−/− ES cells, respectively (Fig. 1A). The same results were obtained when the HMGA1 protein levels were evaluated by Western blot analysis (data not shown). To exclude the possibility that changes in Hmga1−/− cell phenotype might be a consequence of genome manipulation, we transfected the Hmga1 null ES cells with the pcDNA3.1Hmga1b/Hygro construct, carrying the Hmga1b cDNA and the gene for resistance to hygromicin under the transcriptional control of the cytomegalovirus promoter. Several hygromicin-resistant clones were picked up, and transgene expression was detected by RT-PCR (Fig. 1B) and Northern blotting (not shown). The Hmga1-expressing clones, R1 and R2, were used in the following experiments. Hmga1−/− ES cells had a reduced capacity to generate EBs Wild-type, Hmga1+/−, and Hmga1−/− ES single-cell suspensions were seeded in methylcellulose-based medium, were grown in suspension in bacteria-grade dishes, or were

permitted to grow in medium droplets under a plate lid (23). Under these conditions, they generated tridimensional cystic structures of differentiated cells, known as the EBs. As shown in Table 2, Hmga1−/− ES cells formed a much lower number of EBs compared with wild-type and single-knockout ES cells. EBs derived from Hmga1−/− ES cells were also smaller than wild-type and Hmga1+/− cells (Fig. 2). The number and size of EBs generated by Hmga1−/− cells transfected with an Hmga1b cDNA expression vector (Hmga1−/−R cells), but not with the empty vector, were similar to those of parental EBs (Fig. 2 and Table 2). These data are consistent with the higher growth rate observed in Hmga1+/+ and Hmga1−/−R undifferentiated ES cells compared with Hmga1−/− cells (data not shown). EB morphology revealed a role for HMGA1 in myeloerythroid hematopoiesis Establishment of the hematopoietic system within EBs is perhaps the most thoroughly analyzed developmental program. In fact, a large number of studies have documented the development of various erythroid, myeloid, and lymphoid lineages within EBs (10, 24). We next analyzed the capacity of Hmga1 null ES cells to differentiate into hematopoietic cell lineages. To do this, ES cells were seeded onto a methylcellulose-based medium with FBS and mSCF and incubated for 25 days. Hematopoietic EBs were identified by the appearance of macrophages and granulocytic cells at the edges of EBs and by hemoglobinization of erythroid cells, identifiable as blood islands in the cystic structures. As shown in Fig. 2 and summarized in Table 2, wild-type ES cells gave rise to hematopoietic EBs with a high frequency (66%). Conversely, only 9.4% of Hmga1−/− EBs showed a hematopoietic phenotype. It is noteworthy that the major loss was in the monocyte-macrophage (myeloid) component of the EBs. Indeed, Hmga1−/− EBs were almost completely devoid of a myeloid halo, whereas blood islands were readily detectable. The Hmga1+/− ES cells showed intermediate values. The deficiency observed in the Hmga1−/− macrophage-monocyte component was efficiently rescued by the expression of the transfected Hmga1b cDNA, but not by transfection of the backbone vector (Fig. 2 and Table 2). In the great majority of Hmga1−/−R EBs, the myeloid halo prevailed on the central body, constituting the largest part of the EBs, and the number of blood islands was greatly reduced. The early developmental stage of the EBs generated by the Hmga1 null ES cells was confirmed by c-Kit expression. In fact, the c-Kit receptor is expressed on hematopoietic progenitor cells. It is highly expressed in the lineage marker-negative (lin−) immature cell population, it is downregulated on maturation in lin+ cells (25), and it has been demonstrated that ES cells can generate c-Kit+ hematopoietic precursors (26). FACS analysis of 25-day-old EBs showed c-Kit expression by 57.8% of Hmga1−/− cells vs. a mere 23.1% of the wild-type ES cells (not shown). RT-PCR analysis confirmed that Hmga1−/− EBs express higher c-Kit levels than do wild-type cells (not shown). Conversely, wild-type and Hmga1−/− EBs did not differ with regard to expression of other hematopoietic markers such as EpoR, c-Myb, and CD45 (not shown). T-cell differentiation pathway was impaired in Hmga1−/− ES cells and embryos FACS analyses were carried out with EBs-derived cells, by using PE- and/or FITC-labeled monoclonal antibodies against T- and B-lymphoid surface markers (Thy-1.2 and B220). These experiments showed that the Thy-1.2 T-cell-specific marker was expressed at higher levels in the wild-type ES cell population (30.3%) than in Hmga1 null ES cells (9.3%) (Fig. 3A). Conversely,

with antibodies against the pan-B-cell marker B220 (CD45R) (19), B220+ cells were higher in Hmga1 null cells than in wild-type cells (58.7% vs. 29.0%). There was no difference in the expression of the surface marker CD44 between Hmga1+/+ and Hmga1−/− EBs (Fig. 3A). Because these results suggested that B- and T-cell precursor development was altered after Hmga1 gene inactivation, we analyzed expression of pre-B- and pre-T-cell markers by RT-PCR. As shown in Fig. 3B, Thy-1.2, which was undetectable in undifferentiated ES cells and after 11 days of culture in methylcellulose medium, was expressed after 20 days. According to FACS data, Thy-1.2 expression level was almost fivefold lower (as evaluated by densitometry) in Hmga1−/− cells than in wild-type ES cells. The reintroduction of Hmga1b in ES cells rescued Thy-1.2 gene expression to levels comparable to those of wild-type ES cells. This finding indicates that HMGA1 is involved, directly or indirectly, in determining the differentiation of Tcell precursors. GATA-3, another marker of T-cell differentiation (27), slightly decreased in double-knockout cells after 20 days in methylcellulose-based medium (Fig. 3B) and, similarly to Thy-1.2, increased in rescued EBs. Conversely, expression of B-cell lymphoid markers VpreB and B29 was more abundant in Hmga1 null ES cells than in wild-type cells (Fig. 3B). Lack of HMGA1 proteins compromised cytokine expression both in vitro and in vivo HMGA1 proteins are modulators of the function of many of the transcriptional factors that control cytokine gene transcription, in which they play either a positive or a negative role depending on the cytokine promoter and its ratio to other transcription factors (28, 29). Because recent findings have shown that expression of IL-2 and IL-2Rα is essential for the autocrine loop that drives T-cell proliferation and clonal expansion after an immune stimulus, we analyzed the IL-2/IL-2Rα pathway in these cells compared with wild-type ES cells to define the mechanisms underlying the reduced capacity of Hmga1−/− ES cells to differentiate along the T-cell lymphocytic pathway. RT-PCR analyses showed that IL-2 was expressed at a higher level in 25-day-old wild-type EBs than in age-matched double-knockout EBs (Fig. 3C). Conversely, expression levels of IL-6, a cytokine that induces terminal maturation of B cells into antibody-producing cells, were higher in Hmga1−/− EBs in comparison to the wild-type EBs (Fig. 3C). There was no difference in the types of EBs in IL-7 expression levels (Fig. 3C). RT-PCR analyses performed with fetal livers from embryos at 14.5 dpc showed higher IL-2 expression in wild-type embryos than in single- and double-knockout embryos (Fig. 3D). Mixed hemopoietic colonies derived from Hmga1−/− EBs were reduced in number and size ES cells were differentiated with the two-step differentiation protocol (see Materials and Methods), by which differentiating ES cells can produce most, if not all, the colony-forming cells found in normal bone marrow (18). Eleven-day-old EBs were disrupted, and cells were replated as described under Materials and Methods. Colonies were observed after 11–20 days in differentiating medium (Fig. 4). Wild-type ES cells gave rise to erythroid (Fig. 4a), macrophage (Fig. 4c), and multilineage (Fig. 4e) colonies. In sharp contrast, Hmga1−/− macrophage and

multilineage colonies were very scarce (Fig. 4d and Fig. 4f). Erythroid colonies remained unchanged (Fig. 4b). Hmga1−/− colonies expressed enhanced levels of early megakaryocyte markers To characterize megakaryocyte differentiation, we performed both molecular and cytochemical analyses. RT-PCR analyses of 25-day-old Hmga1−/− EBs showed a substantial increase in the early mekagaryocyte markers PF-4 (Fig. 5A) and GpIIb (not shown). Interestingly, they also expressed higher IL-6 levels (Fig. 3C) than did wild-type ES cells. This result is consistent with the finding that IL-6 stimulates development of megakaryocytes, which brings them to maturity and increases platelet numbers (reviewed in ref 30). When cytochemical analyses of EB-derived megakaryocyte colonies were performed, CFU-Mks were identified by megakaryocyte-specific acetylcholinesterase activity. Hmga1−/− cells generated more abundant and significantly larger CFU-Mk colonies than did wild-type cells (Fig. 5B). Taken together with the molecular data, this observation indicates that loss of Hmga1−/− results in greatly enhanced proliferation of megakaryocite progenitors. Interestingly, PF-4 expression was increased in fetal livers from Hmga1−/− 14.5-dpc mice compared with wild-type and single-knockout mice (not shown). Loss of Hmga1 was associated with increased globin gene expression in vitro and in vivo To characterize erythropoietic differentiation in Hmga1−/− ES cells, globin gene expression was evaluated in EBs by use of RT-PCR, after treatment with retinoic acid, a potent regulator of mesodermal differentiation. There were striking differences in the expression of ζ-, α-, βΗ1, and β-major globin genes between wild-type and double-knockout cells (Fig. 6A). All four globins were expressed at higher levels in Hmga1−/− ES cells than in wild-type ES cells. In the wildtype cells, ζ-globin was expressed 6 days after retinoic acid treatment (not shown) and was undetectable at 19 days. Conversely, in Hmga1−/− ES cells, ζ-globin expression was still detectable after 19 days of differentiation. There was no difference in the levels of the mesodermal gene brachyury and the neural genes MASH1 and Wnt-1 (not shown). Hmga1−/− R ES cell clones showed the same behavior of the wild-type ES cells (not shown). It is noteworthy that the same results were obtained in EBs grown in methylcellulose-based medium (not shown). These data strongly indicate that the Hmga1 gene plays a main role in erythroid differentiation. Because ES cell-derived erythropoiesis recapitulates in vivo erythropoiesis in the yolk sac (10), we addressed the question of whether the phenotype observed in vitro also occurred in vivo. Fetal globin (βH1 and ζ) gene expression was higher in 9.5- and 14.5-dpc Hmga1−/− yolk sacs (Fig. 6B) and in 14.5-dpc fetal livers (not shown) compared with wild-type tissues. Interestingly, Hmga1 expression was detectable in the yolk sacs of wild-type and single-knockout mice at 9.5 dpc (Fig. 6B). Its expression decreased at 14.5 dpc and increased in fetal liver (not shown). Lack of HMGA1 proteins induced up-regulation of GATA-1 in vitro and in vivo GATA-1 is a key regulator of red blood cell differentiation, and its consensus sequence is contained in all erythroid-specific genes including globin genes (31, 32). Therefore, we analyzed GATA1 mRNA levels in differentiated wild-type and knockout ES cells to verify whether HMGA1 proteins regulate expression of GATA-1 transcription factor. At 25 days in

methylcellulose-based medium, Hmga1−/− cells expressed higher levels of GATA-1 than did wild-type cells (Fig. 7A). Restoration of HMGA1 expression in Hmga1 null cells inhibited GATA-1 expression. In undifferentiated ES cells, GATA-1 expression was up-regulated in Hmga1−/− cells compared with wild-type cells, as detected by Western blotting and RT-PCR (not shown). In yolk sacs from Hmga1−/− 9.5-dpc embryos, GATA-1 was up-regulated compared with wild-type and single-knockout embryos (Fig. 7B), which indicates that expression of Hmga1 and GATA-1 is inversely related in vitro and in vivo. Expression of GATA-1 in 14.5-dpc yolk sacs was barely detectable, whereas it was highly expressed in 14.5dpc fetal livers (not shown). HMGA1 proteins negatively regulated GATA-1 transcription The results reported above suggested that HMGA1 proteins may be involved in down-regulation of GATA-1 expression. A region located at kbp −3.9 to −2.6, 5' to the first hematopoietic exon of the Gata-1 gene and extending to the first intron (int) of the Gata-1 gene, is sufficient to recapitulate Gata-1 gene expression (12). To investigate the effect of HMGA1 on GATA-1 transcription, we transiently transfected wild-type and Hmga1−/− ES cells with reporter constructs in which the luciferase gene was under the control of the GATA-1 proximal promoter (IE) and the first intron regions, with or without the upstream enhancer element (IE3.9intLUC and IE2.6intLUC, respectively). As shown in Fig. 7C, activity of both constructs was higher in Hmga1−/− ES cells than in wild-type cells. In particular, with IE3.9intLUC, luciferase activity was twofold higher in Hmga1−/− ES cells than in wild-type cells. Consistent with our RT-PCR data, when the Hmga1-expressing construct was cotransfected with either of the reporter constructs, a sixfold reduction of luciferase activity was observed, both in wild-type and in double-knockout cells. Conversely, no difference in luciferase activity was observed between wild-type and double-knockout cells when pGL3 was transfected (Fig. 7C) either in presence or absence of the Hmga1 expressing construct. The ability of HMGA1 to repress both IE3.9 and IE2.6 suggests that HMGA1 may bind both the upstream activating element (−3.9/−2.6) and the sequence downstream –2.6. We identified two regions, from −3230 to −2969 (12) (here referred as region A) and from −2001 to −1719 (here referred as region B) particularly rich in AT, and hence potential HMGA1 binding sites. We used chromatin immunoprecipitation to test this hypothesis (see Materials and Methods). Chromatin from wild-type and double-knockout ES cells, immunoprecipitated with antiHMGA1 antibodies or unbound, was amplified by PCR by using specific primers. Both band A (Fig. 8A) and band B (not shown) were enriched in immunoprecipitated chromatin from wildtype ES cells compared with chromatin that was not bound, but not in chromatin immunoprecipitated from Hmga1−/− ES cells. To further investigate the binding of HMGA1 proteins to the GATA-1 upstream regulating sequence, we performed EMSA with the 46-bp AT-rich oligodeoxynucleotide (GATA-1970) (Fig. 8B) included in region B. As shown in Fig. 8C, the purified synthetic protein His-HMGA1 protein bound directly to this DNA region. HMGA1b recombinant protein had a very high affinity for GATA-1970 oligonucleotide; indeed, binding was detected with only 5 ng of protein (lane 2) and generated two specific bands, C1 and C2, that were competed by a 100-fold molar excess of both GATA-1970 oligonucleotide (lane 3) and IL-2RE oligonucleotide (22) (lane 4)

but not by unrelated unlabeled oligonucleotide at the same concentration (lane 5). Because GATA-1970 has multiple AT-rich tracts, the C1 band could result from the cooperative binding of multiple HMGA1b molecules. The specific bands were supershifted by an anti-HMGA1 antibody (lane 6) but not by an unrelated antibody (anti-Pit-1 protein antibody) (lane 7). Six different mutant GATA-1970 oligonucleotides (Fig. 8B) were less able to compete for binding to the probe compared with the wild-type sequence (Fig. 8C, lanes 8–13). The least effective competitor was mutant M6, which lacks 11 T residues; consequently, this stretch represents the strongest HMGA1 binding site. EMSA analysis of protein extracts from undifferentiated wild-type and Hmga1−/− ES cells resulted in two closely migrating bands, B1 and B2, in wild-type ES cells (Fig. 8D, lane 2). Remarkably, band B2 comigrated with band C2 and Hmga1−/− ES cells lacked band B2 (lane 3), which suggests that it resulted from HMGA1 binding alone. However, band B1 was very faint in Hmga1−/− cells compared with wild-type cells, probably because HMGA1 proteins allow the assembly of factors included in complex B1. A 100-fold molar excess of the unlabeled GATA1970 oligonucleotide or specific competitor IL2RE, but not the same amount of an unrelated sequence, competed for binding with the probe (lanes 4–9). Band B2 was considerably reduced when protein extracts from wild-type ES cells were incubated with anti-HMGA1 antibodies (lanes 10 and 11) but not with unrelated antibodies (lanes 12 and 13). The facts that Hmga1−/− ES cells do not bind strongly to this oligonucleotide and that they expressed higher GATA-1 protein levels (not shown) suggest that this region has a negative, HMGA1-mediated effect on GATA-1 transcription, at least in this system. These results indicate that HMGA1 proteins bound the GATA-1 upstream control element and region B, not previously described as a transcription control element. DISCUSSION We used ES cells carrying disruption of one or both alleles of the Hmga1 gene to try to clarify the role of HMGA1 proteins in embryonic development, particularly in lymphohematopoietic differentiation. ES cells spontaneously differentiate and generate various cell types under appropriate culture conditions (9, 10, 23). Therefore, differentiation of ES cells in vitro is a powerful model system for addressing questions related to lineage commitment and offers several advantages over comparable approaches in which the whole embryo is used. Lymphoid differentiation Here, we showed that HMGA1 proteins play a role in determining the fate of the lymphoid cell common precursor both in vitro and in vivo. In fact, the number T-cell precursors was greatly reduced in Hmga1−/− ES cells, whereas the number of B-cell precursors increased. The reintroduction of the cDNA coding for HMGA1b was able to rescue the wild-type phenotype. Because IL-2 is down-regulated in both double-knockout EBs and fetal livers compared with age-matched wild-type counterparts, we suggest that the impairment in T-cell development could be due to a reduction in IL-2 expression. Our results are consistent with the finding that HMGA1 proteins regulate IL-2 gene promoter activity (14, 33) and hence T-cell proliferation (34). HMGA1 expression levels were consistently high in 17.5 dpc thymus of wild-type murine

fetuses (7). In contrast to reported findings of Wiles and Keller (10), other authors have shown that T cells develop in EBs (26). The results of RT-PCR analyses concerning T-cell differentiation are supported by RT-PCR analyses of fetal livers from 14.5 dpc embryos. A common lymphoid precursor has been postulated for B and T lymphocytes (35). Our results from FACS and RT-PCR analyses suggest that HMGA1 expression would force the common lymphoid precursor to differentiate to T rather than B lymphocytes, probably by fine tuning the expression levels of cytokines regulating B- and T-cell proliferation and differentiation. As inferred from our results, expression of HMGA1 would stimulate IL-2 (a T-cell inducer) and down-regulate IL-6 (a B-cell inducer; reviewed in ref 30). Data from our laboratory showed that HMGA1 up-regulated leptin gene expression in functional assays (36); moreover, leptin expression was consistently reduced in Hmga1 null ES cells (data not shown). Because leptin enhances the potential of the fetal liver-derived myeloid/lymphoid colonies (37), it is conceivable that loss of Hmga1 can impair lymphohemopoietic differentiation also by acting on leptin gene expression, and perhaps through other yet to be defined mechanisms. Myeloid differentiation The observation that myeloid differentiation was impaired in Hmga1−/− EBs but not in Hmga1−/−R EBs suggests that HMGA1 proteins are crucial for myeloid lineage commitment. This observation coincides with the finding that HMGA1 physically interacts with the Ets myeloid transcription factor PU.1 and enhances its transcriptional activity (38). Hematopoietic colonies We report here also that Hmga1 null ES cells generated smaller and a greatly reduced number of hematopoietic colonies compared with wild-type cells and that, in contrast to wild-type cells, Hmga1−/− cells did not form multilineage colonies. In fact, the only colonies detected were hemoglobinized erythroid CFUs, which suggests that erythropoiesis is unaffected in Hmga1−/− EBs. These results suggest that HMGA1 is required for normal proliferation and/or survival of hematopoietic multipotential progenitors. A loss in the macrophage population is expected, because HMGA1 regulates the expression of granulocyte-macrophage colony-stimulating factor (22). Expression of the hematopoietic progenitor c-Kit receptor was higher both in Hmga1−/− EBs and ES cells compared with wild-type counterparts. Because c-Kit is expressed in hematopoietic progenitors and is down-regulated during their differentiation (25), c-kit could be important for establishing the hematopoietic phenotype of Hmga1−/− ES cells. Megakaryocyte and erythrocyte differentiation Here, we also showed that megakaryocte and erythroid differentiation processes are increased in Hmga1−/− EBs. It has yet to be established whether or not these megakaryocyte undergo terminal maturation. Moreover, at 25 days in differentiation medium, in Hmga1−/− EBs, globin and Gata-1 gene expression was higher compared with that in wild-type EBs. The increased expression of GATA-1 in Hmga1 null ES cells may account, at least in part, for these modifications. In fact, GATA-1 transcription factor is critical for megakaryocyte growth regulation and platelet biogenesis (reviewed in ref 39), regulates erythroid-specific gene

expression, and affects erythroid differentiation in a dose-dependent manner (31, 32). Our data consistently indicated that GATA-1 expression is negatively regulated by HMGA1. Indeed, we showed that HMGA1 proteins can specifically bind two AT-rich sequences in the GATA-1 upstream activating element and down-regulate GATA-1 promoter activity in functional assays. Our results also agree with previous data (12) showing the presence of a repressive transcriptional activity in the region of the GATA-1 upstream regulating sequence, indicated as 651–1174, and named region A in our study (Fig. 8). Furthermore, negative control regions have been described in the zebrafish GATA-1 promoter region (40). Besides a GATA-1-mediated effect of HMGA1 on globin gene expression, HMGA1 could exert a repressive role by directly binding to silencer sequences. HMGI binds the GATA-1 site in the γ-globin promoter (41). This binding is lost consequent to a point mutation in the HMGI binding site. Interestingly, this mutation has been found in γ-globin promoters found in patients with hereditary persistence of fetal hemoglobin (HPFH) and correlates with adult expression of the fetal γ-globin gene. Similarly, Chase and colleagues (42) suggested that the relative amount of HMGA1 helps determine the repressive state of the β-globin gene. In agreement with these observations, HMGA1 is down-regulated during erythroid differentiation in chick embryos (43) and in primary peripheral blood cells (42). A silencer at the 3'-end of the human ζ-globin gene, bound by nuclear factor-κB (NF-κB), has been described (44). HMGA1 is known to enhance NF-κB binding to this sequence (4), and we found that the homologous murine sequence is bound by HMGA1b recombinant protein (unpublished results). Because different hematopoietic lineages are differently affected in Hmga1−/− ES cells (some of them are compromised, but others are favored), we can argue that what we observe with regard to hematopoietic differentiation of Hmga1−/− ES cells is not just a consequence of poor EB formation. Moreover, all data concerning hematopoietic differentiation in Hmga1−/− embryos confirmed what may be inferred from ES cell differentiation. In conclusion, our results may represent advancement in the understanding of the regulation of the lineage commitment in lymphohematopoietic differentiation. ACKNOWLEDGMENTS This study was supported by the Associazione Italiana per la Ricerca sul Cancro (AIRC), Progetto Finalizzato “Biotecnologie” of Consiglio Nazionale delle Ricerche, and the MURST Project “Terapie Antineoplastiche Innovative.” We thank Dr. Yamamoto and Dr. Onodera for providing the plasmids IE2.6intLUC and IE3.9LUC. We are indebted to Ms. Jean Gilder for editing the text. REFERENCES 1.

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Table 1 Primers for RT-PCR analyses Gene

Size

5'-Sequence

3'-Sequence

Reference

c-Kit

765

5'-TGTCTCTCCAGTTTCCCTGC

5'-TTCAGGGACTCATGGGCTCA

11

IL-7

355

5'-ACATCATCTGAGTGCCACA

5'-CTCTCAGTAGTCTCTTTAG

11

ζ-globin

370

5'-ACCACCATGTCTCTGATG

5'-GTCAGGATAGAAGACAGG

—

Brachiury

230

5'-GCTGTGACTGCCTACCAGAATG

5'-GAGAGAGAGCGAGCCTCCAAAC

16

Wnt-1

266

5'-TCCTCCACGAACCTGTTGACGG

5'-GATTGCGAAGATGAACGCTGTTTC

16

MASH1

301

5'-CTCGTCCTCTCCGGAACTGATG

5'-CGACAGGACGCCCGCCTGAAAG

16

α-globin

350

5'-CTCTCTGGGGAAGACAAAAGCAAC

5'-GGTGGCTAGCCAAGGTCACCAGCA

17

βH1 (fetal)

415

5'-ACCCTCATCAATGGCCTGTGG

5'-TCAGTGGTACTTGTGGGACAGC

18

βminor (adult) 444

5'-ATGGTGCACCTGACTGATGCTG

5'-GGTTTAGTGGTACTTGTGAGCC

18

Thy-1

423

5'-ACAGCCTGCCTGGTGAACCAAA

5'-GGCCCAACCAGTCACAGAGAAA

18

IL-6

585

5'-AACCACGGCCTTCCCTACTTCA

5'-GCATAACGCACTAGGTTTGCCG

18

CD45(Ly-5)

456

5'-CCTGAGTCTGCATCTAAACCCC

5'-CTGAATCTCCCTCGTACACTCC

—

VpreB

400

5'-GTCTGAATTCCTCCAGAGCCTAAGATCCC

5'-CAGGTCTAGAGCCATGGCCTGGACGTCTG

19

B29

322

5'-CACTGAATTCCCAAGGAAGCCCTTGTTCCC

5'-TAAGTCTAGAAGTTCCGTGCCACAGCTGTC

19

Gata-1

409

5'-TGGATTTTCCTGGTCTAGGG

5'-CTTCAAGGTGTCCAAGAACG

17

Gata-3

566

5'-ACGTCTCACTCTCGAGGCAGCATG

5'-GAAGTCCTCCAGCGCGTCATGCAC

17

IL-2

245

5'-TCGCATCCTGTGTCACATTG

5'- TCAATTCTGTGGCCTGCTTG

14

Pf-4

250

5'-CTCTTGACATGAGCGTCGCTGCGG

5'-CTTGATCACCTCCAGGCAGGTGAA

20

GpIIB

352

5'-AGGCAGAGAAGACTCCGGTA

5'-TACCGAATATCCCCGGTAAC

20

c-Myb

475

5'-GAGCTTGTCCAGAAATATGGTCCGAAG

5'-GGCTGCCGCAGCCGGCTGAGGGAC

17

Hmga1

759

5'-AGAGAAGGAGAATGAGCGAG

5'-TACTTTGGTGGGGACATGCT

—

Gapdh

161

5'-ACATGTTCCAATATGATTCC

5'-TGGACTCCACGACGTACTCAG

—

Table 2 Scoring of EBs after 20 days in methylcellulose ES cell line

No. of EBs/plate

Hmga1+/+ Hmga1+/− Hmga1−/− Hmga1−/−CMV Hmga1−/−R1 Hmga1−/−R2

116 ± 12 102 ± 10 69 ± 9 100 ± 12 270 ± 23 95 ± 10

% hematopoietic EBs 66.0 20.0 9.4 5.3 65.0 45.6

Fig. 1

Figure 1. Identification of Hmga1 double-knockout clones and their rescued derivatives. A) Northern blot analysis of Hmga1 expression in wild-type, Hmga1+/+, Hmga+/−, and Hmga−/− ES cells. Ethidium bromide staining of the same gel is shown as control for RNA loading. B) RT-PCR analyses for exogenous Hmga1 expression in Hmga1−/− ES cell clones. RT-PCR was performed on RNA from Hmga1−/−, Hmga1−/−CMV, and two Hmga1−/−R ES cell clones. C stands for the negative control. GAPDH mRNA was amplified as an internal control.

Fig. 2

Figure 2. Phenotypic appearance of ES-derived EBs in methylcellulose-based medium.

Fig. 3

Figure 3. Lack of HMGA1 reduced T-cell and elicited B-cell differentiation both in vitro and in vivo. A) FACS analyses. Black: negative staining; red: PE- or FITC-positive staining. CD44, B220, and Thy-1.2 reactivity in Hmga1+/+ and Hmga1−/− EBs. B) RT-PCR analyses of markers specific for T cells (Thy-1.2 and GATA-3) and B cells (B29 and VpreB) in wild-type, Hmga1−/−, Hmga1−/−CMV, and Hmga1−/−R EBs. GAPDH was included as an internal control. C) IL-2, IL-6, and IL-7 expression in Hmga1+/+ and Hmga1−/− EBs. D) IL-2 expression in fetal livers from Hmga1+/+, Hmga1+/−, and Hmga1−/− 14.5-dpc embryos. GAPDH was included as an internal control.

Fig. 4

Figure 4. Hematopoietic colonies from Hmga1+/+ and Hmga1−/− EBs. Hematopoietic colonies were generated by disruption of 11-day-old EBs and replating them in methylcellulose-based medium. Wild-type cells gave rise to erythroid (a), macrophage (c), and mixed (e) colonies. The Hmga1−/− colonies were smaller and poorly formed (d and f), and only erythroid colonies were detected (b).

Fig. 5

Figure 5. Megakaryocytic differentiation in wild-type and Hmga1−/− ES cells. A) RT-PCR for the megakaryocyte marker PF-4 in Hmga1+/+, Hmga1+/−, Hmga1−/−, Hmga1−/−CMV, and Hmga1−/−R1 colonies. GAPDH expression was included as an internal control. B) Megakaryocyte colonies from Hmga1+/+ and Hmga1−/− ES cells identified by acetylcholinesterase activity.

Fig. 6

Figure 6. Globin gene expression in Hmga1+/+ and Hmga1−/− retinoic acid-differentiated EBs and embryos. A) RT-PCR analyses of ζ-, βH1, α-, and β-major globin in Hmga1+/+ and Hmga1−/− EBs, induced by retinoic acid. B) Globin and Hmga1 gene expression in yolk sacs (YS) from 9.5- and 14.5-dpc Hmga1+/+, Hmga1+/−, and Hmga1−/− embryos. GAPDH expression was evaluated as an internal control.

Fig. 7

Figure 7. HMGA1 proteins regulated GATA-1 expression. RT-PCR analyses of GATA-1 in retinoic acid-treated Hmga1+/+, Hmga1+/−, Hmga1−/−, Hmga1−/−CMV, and Hmga1−/−R1 EBs (A), undifferentiated wild-type and Hmga1−/− ES cells (B, lanes 2 and 3) and yolk sacs (YS) from 9.5- and 14.5-dpc Hmga1+/+, Hmga1+/−, and Hmga1−/− embryos (B, lanes 4–9). GAPDH expression was evaluated as an internal control. C) Luciferase activity of GATA-1 upstream regulating sequences in Hmga1+/+ and Hmga1−/− ES cells. When indicated, 3 µg of Hmga1-expressing vector was cotransfected.

Fig. 8

Figure 8. HMGA1 binding to GATA-1 upstream regulating regions. A) In vivo binding site detection by PCR of immunoprecipitated chromatin DNA. Negative control (lane 1); PCR products of input (lanes 2 and 3) and anti-HMGA1 immunoprecipitated (IP; lanes 4 and 5) DNA from wild-type (lanes 2 and 4) and Hmga1−/− (lanes 3 and 5) ES cells. B) AT-rich, 46-bp oligodeoxynucleotide sequence (from −1970 to −1924) in GATA-1 upstream regulating region, used for EMSA. WT: wild-type sequence; M1–M6: mutant sequences. C) EMSA of recombinant HMGA1b-His with GATA-1 oligonucleotide. Lane 1, probe only; lanes 2–7, HMGA1b-His (5 ng); lane 3, with 100 × unlabeled GATA-1 oligonucleotide; lane 4, with 100 × unlabeled IL2RE oligonucleotide; lane 5, with 100 × unlabeled unrelated oligonucleotide; lane 6, with anti-HMGA1 antibody; lane 7, with anti-Pit-1 antibody; lanes 8–13: with 100 × unlabeled mutant GATA-1 oligonucleotides M1–M6. D) EMSA of wild-type and Hmga1−/− ES cell protein extracts with GATA1970 oligonucleotide probe; lanes 4 and 5, with 100 × unlabeled GATA-1 oligonucleotide; lanes 6 and 7: with IL-2RE competitor; lanes 8 and 9, with a specific competitor; lanes 10 and 11, with anti-HMGA1 antibody; lanes 12 and 13, with anti-Pit-1 antibody. G indicates wild-type GATA-1970 oligonucleotide; IL, IL2CD28RE; Pm, Pm oligonucleotide; M1– M6, mutant GATA-1970 oligonucleotides; αY, α-HMGA1 antibody; αP, α-Pit protein antibody.