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Published as doi: 10.1096/fj.07-097576.
 (The FASEB Journal. 2008;22:2690-2701.)
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The Wilms’ tumor suppressor Wt1 activates transcription of the erythropoietin receptor in hematopoietic progenitor cells

Karin M. Kirschner*, Patricia Hagen*, Christiane S. Hussels{dagger}, Matthias Ballmaier{ddagger}, Holger Scholz*,1 and Christof Dame{dagger},1,2

* Institut für Vegetative Physiologie and

{dagger} Department of Neonatology, Campus Virchow-Klinikum, Charité—Universitätsmedizin Berlin, Berlin, Germany; and

{ddagger} Department of Pediatric Hematology and Oncology, Medizinische Hochschule, Hannover, Germany

2Correspondence: Department of Neonatology, Campus Virchow-Klinikum, Charité–Universitätsmedizin Berlin, Augustenburger Platz 1, D-13353 Berlin, Germany. E-mail: christof.dame{at}charite.de


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The Wilms’ tumor protein Wt1 is required for embryonic development and has been implicated in hematologic disorders. Since Wt1 deficiency may compromise the proliferation and differentiation of erythroid progenitor cells, we analyzed the possible role of the transcriptionally active Wt1 isoform, Wt1(–KTS), in regulating the expression of the erythropoietin receptor (EpoR). Wt1 and EpoR were coexpressed in CD117+ hematopoietic progenitor cells and in several hematopoietic cell lines. CD117+ cells of Wt1-deficient murine embryos (Wt1–/–) exhibited a significantly lower proliferation response to recombinant erythropoietin than CD117+ cells of heterozygous (Wt1+/–) and wild-type littermates (Wt1+/+). EpoR expression was significantly diminished in hematopoietic progenitors (CD117+) that lacked Wt1, and the erythroid colony-forming capacity was reduced by more than 50% in fetal liver cells of Wt1-deficient embryonic mice. Wt1(–KTS) significantly increased endogenous EpoR transcripts in transfected cells. The proximal EpoR promoter of human and mouse was stimulated more than 10-fold by Wt1(–KTS) in transiently cotransfeced K562 erythroleukemia cells. A responsible cis-element, which is highly conserved in the EpoR promoter of human and mouse, was identified by mutation analysis, electrophoretic mobility shift assay, and chromatin immunoprecipitation assay. In conclusion, activation of the EpoR gene by Wt1 may represent an important mechanism in normal hematopoiesis.—Kirschner, K. M., Hagen, P., Hussels, C. S., Ballmaier, M., Scholz, H., Dame, C. The Wilms’ tumor suppressor Wt1 activates transcription of the erythropoietin receptor in hematopoietic progenitor cells.


Key Words: erythropoiesis • colony-forming unit-erythroid • gene expression • development • oncogene


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
IN ADDITION TO ITS FUNCTION as a suppressor of pediatric kidney cancer (Wilms’ tumor, nephroblastoma) (1 , 2) the Wt1 protein also plays a role in normal human hematopoiesis and in leukemia. During hematopoiesis, Wt1 is expressed in a small subset of dormant cells, as well as in lineage-committed progenitor cells, but it is not detectable in mature blood cells (3 4 5 6) . Fetal liver cells from embryonic mice lacking Wt1 (C57/B6 Wt1–/–) have an ~75% reduction in erythroid blast-forming units (BFU-Es), erythroid colony-forming units (CFU-Es), and colony-forming unit-granulocyte macrophage-erythroid-megakaryocyte (CFU-GEMM) (7) . Furthermore, forced expression of Wt1 by retroviral infection induced myelomonocytic differentiation of CD34+ human hematopoietic progenitors and quiescence of a subset of immature CD38–/CD34+ cells (8) . In murine 32D cl3 cells, which differentiate in response to granulocyte colony-stimulating factor (G-CSF), expression of Wt1 also resulted in an accelerated differentiation (9) . The proposed role of Wt1 in murine hematopoiesis was challenged by a recent study detecting no significant differences between the clonogenic potential of hematopoietic progenitor cells from the aorta-gonad-mesonephros (AGM) region of wild-type (Wt1+/+) and Wt1-null (Wt1–/–) embryos (10) . Although the reasons for these partly controversial results remain to be resolved, valuable evidence of the function of Wt1 during hematopoiesis can be obtained from the identification of its downstream target genes in hematopoietic precursor cells.

The Wt1 gene encodes zinc finger proteins, which can function as both transcription factors and post-transcriptional regulators. Among the four alternative Wt1 splice variants, insertion of a lysine-threonine-serine (+KTS) tripeptide between the third and fourth zinc finger reduces the DNA binding affinity of Wt1 protein (11 , 12) . The Wt1(+KTS) forms presumably fulfill a role in RNA processing (13) , whereas Wt1 molecules without the KTS splice insertion, Wt1(–KTS), interact with GC- and TC-rich DNA sequences and function as transcription factors (14) . Expression of Wt1 is highest in the developing embryo, where it is detected in a limited set of tissues, predominantly in progenitor cells residing within the genitourinary system (15) . Direct proof that Wt1 is necessary for embryonic development came from gene-targeting experiments in mice. Mouse embryos with inactivation of both Wt1 alleles are usually lethal around midgestation and exhibit a predominant feature with the lack of kidneys and gonads (16) , in addition to defects in the development of the spleen (17) and the nervous system (18 , 19) .

The molecular mechanisms by which the transcription factor Wt1(–KTS) regulates hematopoietic differentiation are not fully understood. Such mechanisms may involve a direct effect on target genes in hematopoietic progenitor cells and/or the interaction with other proteins (20 , 21) . Focusing on the implication of Wt1 in erythropoiesis, we reported recently that Wt1 activates transcription of the erythropoietin (Epo) gene in fetal mouse liver (22) . It is therefore conceivable that impaired expression of the major hematopoietic growth factor Epo may contribute to a compromised hematopoiesis in Wt1 deficiency. However, the report of a reduced in vitro proliferative capacity of Wt1–/– early erythroid progenitors, in particular, of BFU-E and CFU-E (8) , points rather to a cell-autonomous defect. Such defect may include a diminished expression of hematopoietic growth factor receptors in the absence of Wt1.

The Epo receptor (EpoR) is essential for the development of ‘definitive’ hematopoiesis, as indicated by the intrauterine death of mice with homozygous deletion of the EpoR gene (EpoR–/–) between day e10.5 and e13.5. In vitro, Epo binding to the EpoR contributes to the commitment of hematopoietic precursors (CFU-GEMM) to the erythroid lineage and the differentiation of BFU-E to CFU-E. At later stages, the EpoR is crucial for the survival of CFU-E progenitors and their terminal differentiation (23 , 24) . EpoR expression can be regulated at the transcriptional and post-transcriptional level and by post-translational modification (25 26 27) . At the level of transcription, EpoR mRNA expression is controlled by positive and negative cis-regulatory elements that are located upstream of the EpoR gene. A minimal EpoR promoter element has been identified in the 5'-untranslated region (UTR) between the nucleotides (nt) –76 and nt + 33, relative to the transcription start site (28) . As previously shown, the GC-rich EpoR promoter, which does not contain a TATA box, can be trans-activated by the hematopoietic transcription factor GATA-1 and the ubiquitously expressed Sp1 protein (29 30 31) . Interestingly, our sequence inspection revealed several predicted binding motifs for Wt1 (GnGGGnGnG) in highly conserved regions of the EpoR promoter. In view of the above, and to clarify the role of Wt1 in murine hematopoiesis, we examined the hypothesis that the EpoR gene could be a molecular downstream target of the Wt1 transcription factor.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals
Mouse breeding pairs (C57BL/6 strain) with heterozygous Wt1 deletion (Wt1+/–) were originally obtained from the Jackson Laboratory (Bar Harbor, ME, USA) and mated in compliance with legal authorities. The embryos were collected at the indicated time points [morning of vaginal plug was considered embryonic day (e) 0.5] and genotyped by polymerase chain reaction (PCR) (18) .

Cell culture
Human hepatoma-derived HepG2 cells (ATCC HB-8065) and human embryonic kidney (HEK) 293 cells (ACC 305) were purchased from the American Type Culture Collection (Manassas, VA, USA) and grown as described previously (22 , 32) . All other cell lines were obtained from the German Collection of Microorganisms and Cell Cultures (Braunschweig, Germany): human erythroleukemia K562 cells (ACC 10), human acute myeloic leukemia UT-7 cells (ACC137), human erythroleukemia (HEL) cells (ACC 11), human B-cell precursor leukemia Reh cells (ACC 22), human neuroblastoma SH-SY5Y cells (ACC 209), and mouse erythroleukemia MEL-745A cells (ACC 501). The cells were grown according to the supplier’s protocol.

Isolation of hematopoietic progenitor cells from murine fetal liver
Hematopoietic progenitors were isolated from murine fetal liver (e11.5) by magnetic cell sorting using MACS CD117 MicroBeads (Miltenyi Biotec, Bergisch Gladbach, Germany) according to the supplier’s protocol. In brief, time-phased pregnant heterozygous Wt1+/– mice were sacrificed by cervical dislocation. Fetuses with the adhering placentas were removed after laparoscopy and placed in ice-cold Iscove’s medium supplemented with 2% fetal bovine serum (FBS). The livers were carefully isolated, and the tails of the embryos were collected for genotyping. The hepatic tissue was minced in a Petri dish using a razor blade. After addition of 1 ml Iscove’s medium, the cell aggregates were disrupted by 3 or 4 drawings through a 21-gauge needle. The disrupted tissue was transferred into 14-ml tissue culture tubes and incubated for 5 min on ice to allow larger fragments to settle. The remaining cells were centrifuged for 10 min at 400 g and resuspended at 107 cells per 80 µl in phosphate-buffered saline, pH 7.2; 0.2% BSA; and 2 mM EDTA. After addition of 20 µl CD117 microbeads, the cells were incubated for 15 min on ice. CD117+ cells were isolated by magnetic separation and cultured in Iscove’s medium supplemented with 20% BIT 9500 (StemCell Technologies, Vancouver, BC, Canada), 1% glutamate, 40 µg/ml low-density lipoprotein, 100 µmol/l β-mercaptoethanol, and 100 ng/ml stem cell factor. Recombinant human Epo (rhEpo; Epoetin-beta, Roche, Grenzach-Wyhlen, Germany) was added to the culture medium as indicated.

In vitro progenitor cell assays and benzidine staining
Wt1+/+, Wt1+/–, and Wt1–/– fetal livers, taken at day e11.5 of development, were dissociated in Iscove’s medium supplemented with 2% fetal bovine serum as described above. Cell aliquots were counted in a Neubauer chamber, and 1 x 105 cells were seeded per well and cultured for 2 days as duplicates in Methocult GF M3434 medium (StemCell Technologies) in a humidified 5% CO2 atmosphere at 37°C. For the detection of erythroid progenitors, the livers of Wt1+/+, Wt1+/–, and Wt1–/– littermates were dissociated, and hemoglobin-positive cells were detected by benzidine staining after lysis of red blood cells (33) . Hemoglobin-containing cells were counted in a Neubauer chamber.

EMSA
In vitro binding assays were performed with GST-purified Wt1, as described previously (22) . A double-stranded oligonucleotide (oligo –32/–8) was chosen on the basis of predicted Wt1 binding sites in the human EpoR promoter [National Center for Biotechnology Information (NCBI) accession no. M76595.1]: 5'-GAGTGCTGGCCCCGCCCCCTCGGGG-3' and end-labeled with 32P. Two different base pair mutations were introduced into oligo –32/–8 either alone (mutant A, 5'-GAGTGCTGGCAACGCCCCCTCGGGG-3'; mutant B, 5'-GAGTGCTGGCCCCGCAAACTCGGGG-3') or in combination (mutant A/B, 5'-GAGTGCTGGCAACGCAAACTCGGGG-3'). An unrelated oligonucleotide corresponding to nt –116/–87 relative to the transcription start site of the human Epo gene (NCBI M11319.1) served as a negative control (5'-TGCTCTGACCCCGGGTGGCCCCTACCCCTG-3').

Chromatin immunoprecipitation (ChIP) assay
ChIP assay was performed in duplicate with freshly prepared cells from murine fetal livers at e11.5 (n=8 per experiment) by using a ChIP Assay Kit (17-295; Upstate, Lake Placid, NY, USA). Proteins were cross-linked to the DNA by incubation of the cells with formaldehyde at a final concentration of 0.4%. For immunoprecipitation, antibodies (4 µg each) against acetylated histone 3 (06-599; Upstate), Wt1 (C-19), Sp1 (PEP-2), GATA-1 (N-6), STAT5 (H-134), and RNA polymerase II (N-20) were used (all from Santa Cruz Biotechnologies, Santa Cruz, CA, USA). Incubation with no antibody served as a negative control, the "input" sample as a positive control. For analysis of ChIP experiments, the following primer pairs were used to amplify a 330-bp sequence within the murine EpoR promoter (NCBI X53081): 5'-TGACCACATTAGCAAAGCCA-3' (forward primer), 5'-CCCTGAGTTTGTCCATGATG-3' (reverse primer). A 253-bp sequence within the murine thrombopoietin receptor (TpoR) promoter (NCBI Z22657.1) was amplified using the following oligonucleotides: 5'-AGAGACAGGGACAGGACGT-3' (forward primer), 5'-AGGGTCTCCCTTTGCCTGT-3' (reverse primer).

Plasmids
A 309-bp sequence of the human EpoR promoter (nt –309 to nt+4 relative to the transcription start site; NCBI M76595) was cloned by PCR from genomic DNA using the PCR primers shown in Table 1 . The amplified DNA was ligated into the SacI and HindIII restriction sites of the pGL2 reporter vector (Promega, Mannheim, Germany). Likewise, a 180-bp sequence of the proximal murine EpoR promoter (nt –180 to nt+17 relative to the transcription start site; NCBI M38133) was cloned and ligated into the pGL2 reporter vector. The Wt1 expression constructs (murine Wt1 cDNA +/–KTS in pCB6+) were kindly provided by Daniel A. Haber (Massachusetts General Hospital Cancer Center and Harvard Medical School, Charlestown, MA, USA) (34) . A PCR-based protocol was applied to introduce site-directed mutations into the predicted Wt1 binding elements within the human and mouse EpoR promoter. The mutant EpoR promoter sequences were ligated into the pGL2 basic reporter vector and verified by automated sequencing (MWG Biotech, Ebersberg, Germany).


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  		Table 1. Oligonucleotides used for PCR amplification of the human and murine wild-type EpoR promoter and for introduction of site-directed mutations

Cell transfections and reporter gene assays
K562 and HEK293 cells were transiently cotransfected with the following plasmids using Geneporter2, according to the supplier’s protocol (Gene Therapy Systems, PEQLAB, Erlangen, Germany): 100 ng promoter-firefly luciferase plasmid, 125 ng Wt1(–KTS), Wt1(–KTS) or empty pCB6+ expression constructs, and a Renilla luciferase plasmid for normalization of the transfection efficiencies. The cells were lysed 36 h after transfection, and luciferase activities were measured in a luminometer (Microlite TLX1, MGM Instruments, Hamden, CT, USA) with luciferin (Promega) as a substrate. Values are presented as relative light units, and the results are averages of 5 transfection experiments, each performed in duplicate.

Quantitative RT-PCR analysis of gene expression
Total RNA preparation and first-strand cDNA synthesis were carried out as described previously (35) . A 1:200 volume aliquot of the reaction was taken for PCR quantification with the GeneAmp 5700 Sequence Detection System (Perkin-Elmer, Applied Biosystems, Foster City, CA, USA) (22 , 36) . Primer sequences that were used for PCR amplification are shown in Table 2 . Transcript levels were compared on the basis of differences in the threshold cycles (Ct) values. Only samples with equal levels of GAPDH mRNA (± 0.5 Ct) were taken into account. Transcript levels of the gene of interest are indicated as the difference between the Ct value of H2O (background) and the Ct value of the respective gene.


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  		Table 2. Primer sequences used for RT-PCR of indicated genes

Immunoblot
Western blot analysis of Wt1 protein expression in HepG2 cells that had been transfected with Wt1 expression constructs or with "empty" vector was performed as described elsewhere (22) . A polyclonal anti-Wt1 antibody from rabbit (C-19; Santa Cruz Biotechnology; 1:100 dilution in PBS, 5% Blotto, 0.05% Tween-20) and a goat polyclonal anti-β-actin antibody (C-11; Santa Cruz Biotechnology; 1:500 dilution) were used for protein detection.

Statistics
ANOVA with Bonferroni test as post hoc test calculations and Student’s t test were performed as indicated to reveal statistical significances. Values of P < 0.05 were considered statistically significant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
EpoR mRNA is reduced in CD117+ cells from Wt1-deficient murine fetal liver
We first examined whether EpoR expression levels differ in wild-type and Wt1-mutant hematopoietic progenitor cells. For this purpose, CD117+ cells were isolated by magnetic cell sorting from the livers of Wt1+/+, Wt1+/–, and Wt1–/– murine embryos (e11.5). EpoR transcripts were significantly reduced by 36 ± 16% in hematopoietic progenitor cells of homozygous Wt1-deficient vs. normal and heterozygous fetal liver (Fig. 1 ). No significant differences in EpoR mRNA levels were found between Wt1+/+ and Wt1+/– CD117+ cells. To exclude that the lower EpoR mRNA levels resulted from a general down-regulation of gene expression in Wt1–/– cells, we analyzed two other hematopoietic growth factor receptors that are normally expressed during the early stages of murine hematopoiesis. As shown in Fig. 1 , no significant differences in the expression of thrombopoietin receptor (TpoR [c-mpl]), granulocyte-colony stimulating factor (G-CSF) receptor, and the selected stem cell receptor (c-kit) were detected between Wt1+/+, Wt1+/–, and Wt1–/– progenitor cells. Likewise, mRNA levels of Gata-1 and Sp1, two established transcriptional activators of the EpoR gene (29 30 31) , were similar in normal and Wt1-mutant (Wt1+/–, Wt1–/–) CD117+ cells (Fig. 1) . These findings indicate that Wt1 is required for normal EpoR transcripts in CD117+ hematopoietic precursors from fetal mouse liver. Furthermore, Gata-1 and Sp1 are unlikely to account for the reduced EpoR expression in Wt1-deficient cells. Quantitative analysis of EpoR protein expression could not be performed by Western blot or immunohistochemistry due to nonspecific cross-reactivity with several other proteins, i.e., heat shock protein (HSP) 70 (37 38 39) . Flow cytometric analysis using biotinylated rhEpo (Epoetin-beta) failed because of limited sensitivity of the method (data not shown).


Figure 1
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  		Figure 1. Gene expression in hematopoietic progenitor cells from normal wild-type (Wt1+/+) and Wt1-mutant (Wt1+/–, Wt1–/–) mouse embryos. Real-time RT-PCR analysis of gene expression in freshly isolated CD117+ cells from the livers of Wt1+/+, Wt1+/–, and Wt1–/– littermates at e11.5. Transcripts were normalized to Gapdh mRNA levels and defined as 1 in Wt1+/+ embryos. Values are means ± SD. *P < 0.01; Student’s t test. EpoR transcripts were reduced by ~40% in CD117+ cells of Wt1–/– vs. Wt1+/+ and Wt1+/– embryos. In contrast, receptors of thrombopoietin (c-mpl), granulocyte colony-stimulating factor (G-CSFR), and the selected stem cell factor (c-kit) were expressed at a similar level in all groups. Similarly, no differences were detected in the expression of Gata-1 and Sp1, two previously identified activators of the EpoR gene (29 30 31) , between Wt1+/+ vs. Wt1+/– and Wt1–/– hematopoietic progenitor cells.

Wt1-deficient hematopoietic progenitor cells display a reduced proliferation response to recombinant human erythropoietin
We investigated next whether the lower EpoR levels in Wt1–/– hematopoietic progenitor cells correlate with a reduced susceptibility to recombinant human erythropoietin (rhEpo). For this purpose, CD117+ cells were freshly isolated from the livers of Wt1+/+, Wt1+/–, and Wt1–/– murine embryos (C57/B6) at e11.5. In each experiment, progenitors from two Wt1+/+, Wt1+/–, and Wt1–/– littermates were seeded at 1 x 105 viable cells per well and grown for 48 h in the presence of rhEpo at concentrations between 10–5 and 6 U/ml. Figure 2 shows the representative results of one of three independent experiments. The characteristic S-shaped dose–response curve was shifted to higher rhEpo concentrations in Wt1-deficient compared to normal and heterozygous progenitor cell assays. The EC50 values were 38 mU/ml (Wt1+/+), 43 mU/ml (Wt1+/–), and 122 mU/ml (Wt1–/–) of rhEpo. Cell growth was maximal at rhEpo concentrations above 1 U/ml medium but remained significantly lower in Wt1-deficient than in wild-type and heterozygous hematopoietic progenitors even with 6 U rhEpo/ml medium (Fig. 2) . CD117+ cells failed to grow in the absence of rhEpo, no matter whether they had been obtained from wild-type or Wt1-mutant fetal liver (data not shown). These findings indicate that Wt1 is required for the normal proliferation of murine hematopoietic progenitor cells in response to rhEpo.


Figure 2
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  		Figure 2. Effect of rhEpo on the growth of murine hematopoietic progenitor cells. CD117+ cells were freshly isolated from the livers of Wt1+/+, Wt1+/–, and Wt1–/– embryos at day e11.5. Cells were seeded at a density of 1 x 105 per well and cultured in the presence of rhEpo (10–5 to 6.0 IU/ml medium). Cells were counted after 2 days. Data points represent means ± SD of 4 independent cultures at each rhEpo concentration. Dose-response curves represent a nonlinear least-square fit to the data points. A shift of the curve to higher rhEpo concentrations is seen in Wt1–/– CD117+ cells, indicating a reduced proliferative capacity of hematopoietic progenitor cells in the absence of Wt1.

In vitro erythroid differentiation is compromised in Wt1–/– fetal liver cells
In vitro hematopoietic progenitor assays were performed with freshly prepared cells from murine fetal liver (e11.5) to explore whether the reduced proliferation response of Wt1-deficient CD117+ cells to rhEpo would correlate with impaired erythroid differentiation. Because previous studies yielded conflicting results with regard to the colonogenic potential of Wt1–/– vs. Wt1+/+ hematopoietic progenitor cells (7 , 10) , we applied the same protocol using nonselected murine fetal liver cells. The freshly isolated cells were grown in MethoCult medium supplemented with 3 U rhEpo/ml medium to stimulate differentiation along the erythroid lineage. Cultures derived from Wt1–/– embryos gave less than 50% the number of colonies as those obtained from wild-type littermates. No differences in the colony-forming potential were seen between progenitor cells of Wt1+/+ and Wt1+/– embryos (Fig. 3 A). In another set of experiments, hemoglobin staining with benzidine was used as a measure of erythroid differentiation. As shown in Fig. 3B , the fraction of benzidine-reactive cells was significantly smaller in freshly prepared Wt1–/– vs. Wt1+/+ and Wt1+/– fetal liver cells.


Figure 3
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  		Figure 3. Hematopoietic potential of liver cells from Wt1+/+, Wt1+/–, and Wt1–/– embryonic mice, and EpoR mRNA expression in the liver and heart. A) Colony-forming capacity of murine fetal liver cells. Freshly prepared e11.5 fetal liver cells were grown at 37°C and 5% CO2 in Methocult GF M3434 medium containing rhEpo (3 IU/ml). Notably, erythroid colony formation was reduced significantly in Wt1–/– vs. Wt1+/– and Wt1+/+ cells. *P < 0.01; Student’s t test. B) Hemoglobinization of freshly isolated liver cells from murine embryos at day e11.5. The cells were stained with benzidine to visualize intracellular hemoglobin. Values are means ± SD. *P < 0.01; Student’s t test. C) EpoR mRNA expression in liver and heart of mouse embryos. Ten livers and 7 hearts were collected at day e11.5. EpoR transcripts were measured by real-time RT-PCR and normalized to Gapdh mRNA levels. Values are means ± SD. *P < 0.005; Student’s t test.

Moreover, we investigated whether the differences in EpoR mRNA expression between normal and Wt1-deficient CD117+ cells (Fig. 1) could be recapitulated in intact hematopoietic tissue. For this purpose, fetal livers were collected from Wt1+/+, Wt1+/–, and Wt1–/– embryos at day e11.5, and gene expression patterns were analyzed by real-time RT-PCR. EpoR transcript levels were reduced by ~50 ± 25% (P<0.005, n=10 each) in Wt1–/– vs. Wt1+/+ livers (Fig. 3C ). No significant decrease in EpoR mRNA was seen in the livers of Wt1+/– embryos. Hearts were analyzed as a nonhematopoietic tissue that expresses both EpoR und Wt1. Remarkably, EpoR mRNA expression, which was normalized to Gapdh transcripts, was similar in the hearts of Wt1+/+, Wt1+/–, and Wt1–/– embryos at e11.5 (Fig. 3C ).

Wt1 and EpoR are coexpressed in CD117+ fetal liver cells and hematopoietic cell lines
As a prerequisite for EpoR regulation by Wt1, one would expect that both genes are coexpressed in a population of hematopoietic progenitor cells. To determine the distribution of Wt1 and EpoR in hematopoietic and nonhematopoietic precursors, we isolated CD117+ and CD117– cells from the liver of normal mouse embryos (e11.5). The selection procedure was highly reliable, as indicated by the expression of stem cell factor receptor, c-kit, exclusively in CD117+ but not in CD117– cells (Fig. 4 A). Importantly, EpoR and Wt1 were coexpressed in CD117+ cells, which represent a population of predominantly hematopoietic progenitors. Their hematopoietic origin is reflected by the significant enrichment of transferrin receptor (Trf-R) mRNA compared to CD117– cells (P<0.01; Fig. 4A ). In contrast, expression levels of the ubiquitous Sp1 molecule, an established transcriptional activator of the EpoR gene (31 , 40) , were significantly higher (P<0.05) in CD117– vs. CD117+ cells. Wt1 transcripts could not be detected in CD117– cells, which expressed the EpoR gene at a similar level as CD117+ progenitors (Fig. 4A ).


Figure 4
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  		Figure 4. Analysis of gene expression in progenitor cells from murine fetal liver. CD117+ and CD117– cells were isolated by magnetic cell sorting from the livers of 3 wild-type murine embryos at e11.5. A) Notably, Wt1 transcripts were detected in CD117+ cells, but not in CD117– progenitors. Conversely, Sp1 mRNA levels were significantly higher in CD117– cells than in CD117+ cells. Efficacy in selecting hematopoietic progenitors is indicated by the higher levels of transferrin receptor (Trf-R) mRNA in CD117+ compared to CD117– cells. B) The relative gene expression levels of EpoR and Wt1 in various hematopoietic and nonhematopoietic cell lines. Although Wt1 and EpoR mRNA were detected in all hematopoietic cell lines, including human erythroleukemia (K562, HEL, UT-7) and leukemia (Reh) cells and murine erythroleukemia (MEL) cells, among nonhematopoietic cells investigated, only HepG2 cells expressed also both genes.

CD117 (c-kit) is not only present in hematopoietic precursors but also in mesenchymal cells and immature hepatocytes in the fetal liver (41 , 42) . To address the relationship between Wt1 and EpoR in a complementary approach, we determined the expression levels of both genes in different hematopoietic and nonhematopoietic cell lines. Remarkably, Wt1 and EpoR transcripts were detected in all hematopoietic cell lines examined, including human erythroleukemia (K562, HEL, UT-7) and leukemia (Reh) cells, as well as murine erythroleukemia (MEL) cells (Fig. 4B ). The only nonhematopoietic cell line that coexpressed both genes at a detectable level were human hepatoma-derived HepG2 cells. All other tested cell lines contained either Wt1 (HEK293 cells) or EpoR mRNA (SH-SY5Y human neuroblastoma cells) only (Fig. 4B ). These results indicate that Wt1 and its proposed molecular target gene, EpoR, are coexpressed in CD117+ cells from murine fetal liver and in different hematopoietic cell lines.

EpoR mRNA is increased by transient expression of the Wt1(–KTS) protein in HepG2 cells
To examine whether Wt1 activates the expression of the EpoR gene in vitro, HepG2 cells were transiently transfected with different Wt1 expression constructs (4 independent experiments). Human HepG2 cells were chosen because of their higher transfection efficiencies compared to other cell lines. Using a variety of different protocols for the transfection of hematopoietic cells, Wt1 protein could not be increased above the endogenous level in these cells. However, HepG2 cells, which served also as a model for hepatic Epo production (43) , have a low basal expression of both, EpoR and Wt1. EpoR transcripts increased approximately 2-fold within 48 h after transient transfection with the Wt1(–KTS) construct compared to cells that had been transfected with "empty" pCB6+ plasmid. In contrast, transfection with the Wt1(+KTS) form did not significantly change the endogenous EpoR mRNA content (Fig. 5 A). Transient transfection with Wt1(–KTS) and Wt1(+KTS) expression constructs resulted in a moderate but significant increase of Wt1 protein in HepG2 cells (Fig. 5B ). These observations indicate that the Wt1(–KTS) isoform increases EpoR mRNA in vitro either directly or through indirect mechanisms.


Figure 5
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  		Figure 5. Stimulation of EpoR expression by the Wt1(–KTS) protein in human HepG2 cells. A) EpoR and GAPDH transcripts were determined by real-time RT-PCR 48 h after transient transfection of HepG2 cells with expression vectors encoding for Wt1(–KTS), Wt1(+KTS), or the empty pCB6+ vector, respectively. Values are means ± SD of 4 independent experiments. *P < 0.005; ANOVA. B) Western blot analysis confirms expression of Wt1(–KTS).

The Wt1(–KTS) protein binds to the EpoR promoter
EMSAs were performed to explore whether Wt1 protein can bind to the proximal EpoR promoter. On the basis of its sequence similarity with known Wt1-binding sites of other genes, we selected a double-stranded DNA oligonucleotide located between the nucleotides nt –32 and nt –8 relative to the transcription start site in the human EpoR promoter, which is also highly conserved in the mouse. As shown in Fig. 6 A (left panel), the Wt1(–KTS) protein bound to the oligonucleotide –32/–8 with higher affinity than the Wt1(+KTS) molecule. The two Wt1 proteins did not interact with an oligonucleotide derived from the promoter of the human Epo gene (22) , which served as a negative control. Two potential Wt1 motifs within the oligonucleotide –32/–8 were analyzed in more detail by introducing mutations to abrogate binding of the Wt1(–KTS) protein. As shown in Fig. 6A (right panel), a double base-pair mutation at positions nt –22 to nt –21 in the oligonucleotide (mutant A) did not diminish Wt1(–KTS) binding. However, mutations at positions nt –17 to nt –15 (mutant B) clearly abolished the DNA binding activity of Wt1(–KTS). Consistently, Wt1(–KTS) did not bind to an oligonucleotide that contained both mutations in combination (mutant A/B).


Figure 6
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  		Figure 6. Interaction of Wt1 with the EpoR promoter in vitro and in vivo. A) EMSA demonstrates binding of the recombinant Wt1(–KTS) protein to a 25-bp oligonucleotide (oligo –32/–8) of the human EpoR promoter (lane 2). Arrowhead (>) indicates the specific Wt1-mediated shift. Wt1(+KTS) protein, which plays a role in RNA processing rather than in transcriptional regulation (13) , shows strongly reduced interaction with oligo –32/–8 (lane 3). Incubation of Wt1(–KTS) or Wt1(+KTS) protein with an unrelated oligonucleotide derived from the human Epo gene promoter (22) confirms the specificity of Wt1 binding to the human EpoR promoter (lanes 5 and 6). Introduced mutations (mutant B and mutant A/B) into oligo –32/–8 abrogate binding of the Wt1(–KTS) protein (lanes 10 and 12), whereas the affinity for Wt1(–KTS) is maintained in mutant A (lane 8). B) Scheme of the murine EpoR 5'-untranslated region. Arrow indicates the transcription start site (+1), and the start codon (ATG) is indicated. The position of nucleotides is given relative to the transcription start according to the reference sequence (NCBI X53081). ChIP analysis of protein interaction with the EpoR gene was performed in freshly prepared murine fetal liver cells (e11.5; bottom panel). Control experiments were performed by amplification of a sequence within the murine thrombopoietin receptor (mTpoR) promoter. Panel shows the PCR-amplified products of the immunoprecipitates, which were electrophoresed in a 1.5% agarose gel and stained with ethidium bromide; for better visualization, the gel photograph is presented as a negative of the original. In addition to Wt1, Sp1, GATA-1, and STAT5 did also interact with the EpoR promoter in vivo.

ChIP assay experiments were performed to investigate whether Wt1 interacts with the promoter of the EpoR gene in its natural chromosomal context. The experiments were carried out with freshly isolated cells from dissected murine fetal liver (e11.5), which is the primary site of hematopoiesis at this stage of development. As shown in a representative agarose gel (Fig. 6B ), Wt1 bound to the proximal murine EpoR promoter. Likewise, Sp1, Gata-1, and STAT5 also interacted with the EpoR promoter in murine fetal liver cells. In contrast, Wt1, like Gata-1 and STAT5, did not bind to the murine TpoR promoter (Fig. 6B ), which is consistent with our finding of a normal Tpo receptor mRNA expression in Wt1-deficient hematopoietic progenitors (Fig. 1) .

Wt1(–KTS) protein activates the EpoR promoter
Luciferase-reporter gene assays were performed in HEK293 and K562 erythroleukemia cells to examine whether binding of the Wt1 proteins can activate the EpoR promoter. Therefore, reporter gene constructs, which contained the human and mouse EpoR promoter sequence in the pGL2basic vector, were transiently cotransfected with Wt1(–KTS), Wt1(+KTS), or empty (pCB6+) expression plasmids, respectively. Compared to cotransfection with the pCB6+ plasmid, the Wt1(–KTS) protein enhanced EpoR promoter activity usually more than 10-fold in both cell lines, as indicated by the significant increase of normalized luciferase activity (P<0.001; 5 independent experiments; Fig. 7 ). For comparison, the Wt1(+KTS) protein had a minor, though significant, stimulatory effect on EpoR promoter activity in K562, but not in HEK293 cells. Activation of a GC-rich promoter by the Wt1(+KTS) protein, which fulfils a presumed role in RNA processing (13) , has been also reported in our earlier study (44) and may reflect functional redundancy of the different Wt1 molecules. To confirm the cis-elements required for stimulation of the EpoR promoter by the Wt1 protein, we introduced the indicated mutations (mutant A/B; Fig. 7A ), which resulted in a loss of Wt1(–KTS) binding activity in the EMSA experiments (Fig. 6A ). The mutant human EpoR promoter-reporter construct could not be activated by Wt1(–KTS) in HEK293 cells. However, the activity of the mutant reporter construct in K562 cells was slightly increased by Wt1(–KTS). Likewise, the mutant murine EpoR promoter-reporter construct was activated by cotransfection of Wt1(–KTS) in K562 and HEK293 cells (Fig. 7B, C ). Whether these differences in the susceptibility of the mutant promoter-reporter constructs to stimulation by the Wt1 protein represent cell-type-specific phenomena or point to the existence of yet other, unidentified Wt1 binding cis-elements in the GC-rich EpoR promoter, is currently unknown. In either case, stimulation by Wt1(–KTS) was significantly weaker with the mutant vs. wild-type reporter constructs in both cell lines.


Figure 7
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  		Figure 7. Activation of the EpoR promoter by Wt1(–KTS). A) Sequence alignment of the human and murine EpoR promoter fragments, containing the CG-rich Wt1 binding site identified in EMSA and ChIP experiments (see Fig. 6 ). The mutation A/B, resulting in a loss of Wt1(–KTS) binding, is indicated. B, C) Reporter constructs containing the human or murine EpoR promoter sequence were transiently transfected into erythroid K562 cells (B) and HEK293 cells (C) along with expression plasmids for Wt1(–KTS), Wt1(+KTS), and empty vector (pCB6+). A Renilla luciferase vector was used for the normalization of transfection efficiencies. Values are means ± SD of 5 independent experiments, each performed in duplicate. *P < 0.05, **P < 0.01, ***P < 0.001; ANOVA.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Herein, we describe the Wilms’ tumor suppressor, Wt1, as a novel transcriptional activator of the EpoR gene. This conclusion is supported by the fact that the transcriptionally active Wt1(–KTS) isoform stimulates the human and murine EpoR promoter by binding to a GC-rich element in proximity to the transcription start site (Figs. 6 and 7) . Both genes are coexpressed in CD117+ hematopoietic progenitors from fetal mouse liver and in various erythroid cell lines (Fig. 4) . In homozygous Wt1-deficient embryonic mice (e11.5), EpoR expression is significantly reduced in the liver as the primary hematopoietic organ at this stage of development (Fig. 3) . The proliferation of CD117+ progenitors in response to rhEpo and the in vitro erythroid colony-forming potential of murine fetal liver cells (e11.5) are compromised in the absence of Wt1 (Figs. 2 and 3) . Our results suggest that such a defect could be at least in part due to the lower expression level of EpoR in Wt1-deficient CD117+ cells (Fig. 1) . By demonstrating that Wt1 is required not only for the normal expression of Epo in murine fetal liver (22) but also synergistically activates EpoR expression in hematopoietic progenitor cells, our findings establish the Wt1(–KTS) molecule as a transcriptional regulator during hematopoiesis.

Previous work suggests that Wt1 may exert distinct effects on the lineage-specific differentiation of hematopoietic progenitor cells. Thus, Alberta et al. (7) described that murine fetal liver cells lacking Wt1 exhibit an ~75% reduction in the proliferation of BFU-E, CFU-E, and CFU-GEMM progenitors (7) . In contrast, King-Underwood et al. (10) reported that Wt1-deficient murine fetal liver cells can reconstitute hematopoiesis following their transplantation into irradiated recipients. However, Wt1-null embryonic stem cells, although contributing to other organ systems, competed poorly with wild-type stem cells in the formation of the hematopoietic system (7) . These inconsistent findings could be due to differences in the study design and genetic heterogeneity of the animals used. Because both investigations were performed in different mouse strains (C57/B6 and CBA, respectively), this could indicate that the function of Wt1 during murine hematopoiesis is controlled by additional modifier genes. In support of such a possibility, the phenotype of the developmental abnormalities, i.e., malformation of the spleen, seen in Wt1–/– mice is also dependent on the genetic background of the mutant animals (17) .

Unlike other investigators, who isolated nonselected cells from murine fetal liver (7 , 10) , we initially selected for CD117+ progenitors. The hematopoietic source of these cells is revealed by their more than 50-fold higher expression level of the Trf-R compared to CD117– cells (Fig. 4) . As one major novel finding of this study, we show that the reduced proliferation response of Wt1–/– hematopoietic progenitors to rhEpo is associated with a lower expression rate of EpoR (Fig. 1) . Obviously, the decrease of EpoR mRNA was not due to a general inhibition of gene expression in Wt1-deficient CD117+ cells as the transcript levels of several other hematopoiesis-associated genes, such as TpoR (c-mpl) and G-CSFR, were similar to those in wild-type progenitor cells (Fig. 1) . Interestingly, even in the absence of Wt1, EpoR expression is not completely abrogated, but EpoR transcripts are reduced by ~40% in Wt1-deficient hematopoietic progenitor cells. Consistently, murine embryos with targeted inactivation of the Wt1 gene usually develop less severe defects of their hematopoietic system than mice with homozygous lack of EpoR (23 , 24 , 45) . These results suggest that the transcriptional control of EpoR expression is established with some functional redundancy, thereby allowing other transcription factors to compensate to some extent for the absence of Wt1. Importantly, Sp1 and Gata-1, two previously recognized transcriptional activators of the EpoR gene (29 30 31) , were expressed at similar levels in Wt1+/+, Wt1+/–, and Wt1–/– CD117+ cells (Fig. 1) . It appears therefore possible that the two transcription factors maintain a reduced expression rate of EpoR in Wt1–/– progenitor cells. Unfortunately, EpoR expression could not be analyzed at the protein level because of the lack of specific anti-EpoR antibodies (37 38 39) and technical limitations of flow cytometry to detect the expected relatively small number of EpoR on the cell surface of immature CD117+ cells isolated from the murine fetal liver (46) .

To confirm the findings obtained with CD117+ progenitor cells, we performed in vitro progenitor assays with fetal liver cells of wild-type and Wt1-deficient mice (C57/B6 strain). Consistently, Wt1–/– fetal liver cells had a significantly reduced colony-forming potential and a lower number of hemoglobin-positive cells than Wt1+/+ or Wt1+/– cells (Fig. 3) . Indeed, we did not detect differences in the erythroid colony-forming potential between Wt1+/– and Wt1+/+ fetal liver cells. While this observation is in conflict with previous data from another group reporting a reduced colony-forming capacity of Wt1+/– murine fetal liver cells (7) , it is in accordance with the lack of an obvious hematopoietic disorder in neonatal and adult Wt1+/– mice. However, even a small percentage of surface EpoR may suffice to maintain basal proliferative activity (47 , 48) . Together, with the finding that forced Wt1 expression is associated with impaired hematopoiesis (8 , 49) , our results suggest that a balanced level of Wt1 may be required for the normal differentiation of lineage-committed hematopoietic progenitor cells. Such dose-dependent effect of a transcriptional regulator is also known for other factors involved in hematopoiesis, e.g., GATA-3 (50 , 51) .

In addition to validating the role of Wt1 in murine hematopoiesis, our study extends the understanding of the transcriptional regulation of the EpoR gene. So far, GATA-1 and Sp1 have been identified as the major transcriptional regulators of the EpoR gene (29 , 31) . During the differentiation of erythroid progenitor cells, GATA-1 is thought to act on EpoR expression in a cell cycle-dependent manner. As previously shown in logarithmically growing Epo-dependent leukemia cells (UT-7 cell line), both EpoR and GATA-1, but not GATA-2 mRNA levels, concomitantly decrease in the G0/G1 phase and later increase in the S und G2/M phase. These dynamic changes in EpoR mRNA go in parallel with changes in the binding activity of GATA-1 to the EpoR promoter. It has been speculated that the increase of EpoR mRNA during the resting phase, if induced by growth factor starvation, is dependent on other transcription factors than GATA-1 (52) . In accordance, increased levels of GATA-1 alone are not sufficient to maintain or stabilize the erythroid phenotype during differentiation (53) . Since high levels of EpoR expression in hematopoietic cells apparently do not depend on the level of GATA-1 alone, Wt1 could be the previously postulated additional factor required for activation of the TATA-less EpoR promoter in these cells (31) .

The stage-specific expression of transcription factors during erythroid differentiation needs to be precisely fine-tuned. Interestingly, Wt1 in hematopoietic progenitors itself seems to be regulated by GATA-1 and c-Myb (54 , 55) . GATA-1 can transactivate a specific enhancer region 3' of the Wt1 gene, and another GATA-1 sensitive enhancer element is located within the third exon of Wt1 (54 , 55) . This suggests that the level of Wt1 in hematopoietic progenitor cells is tightly controlled by a number of other transcription factors, some of which may activate the EpoR gene also through a direct mechanism. So far, our data do not support a role for Wt1 in EpoR expression in nonerythroid tissues, such as the heart (Fig. 3) . Consistently, while Wt1 and EpoR are coexpressed in all hematopoietic cell lines examined in this study, regulation of both genes does not seem to be tightly linked in nonhematopoietic cells (Fig. 4B ). However, given the importance of EpoR in mediating the neuro- and cardioprotective effects of Epo (56 , 57) , additional elaborate studies will be necessary to address this issue more in detail. With the present study, we identified the EpoR gene as the first downstream target of Wt1 within the hematopoietic cytokine receptor superfamily. Activation of the EpoR gene by Wt1 may represent an important mechanism in normal murine hematopoiesis. Furthermore, other Wt1 gene targets may additionally contribute to the decreased Epo-mediated proliferation of Wt1-deficient hematopoietic progenitors.


   ACKNOWLEDGMENTS
 
The expert technical assistance of Inge Grätsch and Angelika Richter is gratefully acknowledged. The Wt1 expression construct was kindly provided by Daniel A. Haber (Massachusetts General Hospital Cancer Center and Harvard Medical School, Charlestown, MA, USA). The authors also thank Hannes Sallmon and Martin Klar (Department of Neonatology, Campus Virchow-Klinikum, Charité- Universitätsmedizin Berlin), Joachim Fandrey (Department of Physiology, University of Duisburg-Essen, Germany), and Jörg Bungert (Department of Biochemistry and Molecular Biology, University of Florida, Gainesville, FL, USA) for the critical reading and discussion of the manuscript. This study was supported by the Deutsche Forschungsgemeinschaft (Da 484/2–1 and Scho 634/5–1) and the Bundesministerium für Bildung und Forschung (grants NGFN, KGCV1, and 01GS0416).


   FOOTNOTES
 
1 These authors contributed equally to this work.

Received for publication October 10, 2007. Accepted for publication March 27, 2008.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Hastie, N. D. (1994) The genetics of Wilms’ tumor—a case of disrupted development. Annu. Rev. Genet. 28,523-558[Medline]
  2. Hohenstein, P., Hastie, N. D. (2006) The many facets of the Wilms’ tumour gene, WT1. Hum. Mol. Genet. 15(Spec. 2),R196-R201[Abstract/Free Full Text]
  3. Baird, P. N., Simmons, P. J. (1997) Expression of the Wilms’ tumor gene (WT1) in normal hemopoiesis. Exp. Hematol. 25,312-320[Medline]
  4. Hosen, N., Sonoda, Y., Oji, Y., Kimura, T., Minamiguchi, H., Tamaki, H., Kawakami, M., Asada, M., Kanato, K., Motomura, M., Murakami, M., Fujioka, T., Masuda, T., Kim, E. H., Tsuboi, A., Oka, Y., Soma, T., Ogawa, H., Sugiyama, H. (2002) Very low frequencies of human normal CD34+ haematopoietic progenitor cells express the Wilms’ tumour gene WT1 at levels similar to those in leukaemia cells. Br. J. Haematol. 116,409-420[CrossRef][Medline]
  5. Fraizer, G. C., Patmasiriwat, P., Zhang, X., Saunders, G. F. (1995) Expression of the tumor suppressor gene WT1 in both human and mouse bone marrow. Blood 86,4704-4706[Free Full Text]
  6. Maurer, U., Brieger, J., Weidmann, E., Mitrou, P. S., Hoelzer, D., Bergmann, L. (1997) The Wilms’ tumor gene is expressed in a subset of CD34+ progenitors and downregulated early in the course of differentiation in vitro. Exp. Hematol. 25,945-950[Medline]
  7. Alberta, J. A., Springett, G. M., Rayburn, H., Natoli, T. A., Loring, J., Kreidberg, J. A., Housman, D. (2003) Role of the WT1 tumor suppressor in murine hematopoiesis. Blood 101,2570-2574[Abstract/Free Full Text]
  8. Ellisen, L. W., Carlesso, N., Cheng, T., Scadden, D. T., Haber, D. A. (2001) The Wilms tumor suppressor WT1 directs stage-specific quiescence and differentiation of human hematopoietic progenitor cells. EMBO J. 20,1897-1909[CrossRef][Medline]
  9. Loeb, D. M., Summers, J. L., Burwell, E. A., Korz, D., Friedman, A. D., Sukumar, S. (2003) An isoform of the Wilms’ tumor suppressor gene potentiates granulocytic differentiation. Leukemia 17,965-971[CrossRef][Medline]
  10. King-Underwood, L., Little, S., Baker, M., Clutterbuck, R., Delassus, S., Enver, T., Lebozer, C., Min, T., Moore, A., Schedl, A., Pritchard-Jones, K. (2005) Wt1 is not essential for hematopoiesis in the mouse. Leuk. Res. 29,803-812[CrossRef][Medline]
  11. Haber, D. A., Sohn, R. L., Buckler, A. J., Pelletier, J., Call, K. M., Housman, D. E. (1991) Alternative splicing and genomic structure of the Wilms tumor gene WT1. Proc. Natl. Acad. Sci. U. S. A. 88,9618-9622[Abstract/Free Full Text]
  12. Englert, C., Vidal, M., Maheswaran, S., Ge, Y., Ezzell, R. M., Isselbacher, K. J., Haber, D. A. (1995) Truncated WT1 mutants alter the subnuclear localization of the wild-type protein. Proc. Natl. Acad. Sci. U. S. A. 92,11960-11964[Abstract/Free Full Text]
  13. Larsson, S. H., Charlieu, J. P., Miyagawa, K., Engelkamp, D., Rassoulzadegan, M., Ross, A., Cuzin, F., van Heyningen, V., Hastie, N. D. (1995) Subnuclear localization of WT1 in splicing or transcription factor domains is regulated by alternative splicing. Cell 81,391-401[CrossRef][Medline]
  14. Lee, S. B., Haber, D. A. (2001) Wilms tumor and the WT1 gene. Exp. Cell. Res. 264,74-99[CrossRef][Medline]
  15. Pritchard-Jones, K., Fleming, S., Davidson, D., Bickmore, W., Porteous, D., Gosden, C., Bard, J., Buckler, A., Pelletier, J., Housman, D., Van Heyningen, V., Hastie, N. (1990) The candidate Wilms’ tumour gene is involved in genitourinary development. Nature 346,194-197[CrossRef][Medline]
  16. Kreidberg, J. A., Sariola, H., Loring, J. M., Maeda, M., Pelletier, J., Housman, D., Jaenisch, R. (1993) WT-1 is required for early kidney development. Cell 74,679-691[CrossRef][Medline]
  17. Herzer, U., Crocoll, A., Barton, D., Howells, N., Englert, C. (1999) The Wilms tumor suppressor gene wt1 is required for development of the spleen. Curr. Biol. 9,837-840[CrossRef][Medline]
  18. Wagner, K. D., Wagner, N., Vidal, V. P., Schley, G., Wilhelm, D., Schedl, A., Englert, C., Scholz, H. (2002) The Wilms’ tumor gene Wt1 is required for normal development of the retina. EMBO J. 21,1398-1405[CrossRef][Medline]
  19. Wagner, N., Wagner, K. D., Hammes, A., Kirschner, K. M., Vidal, V. P., Schedl, A., Scholz, H. (2005) A splice variant of the Wilms’ tumour suppressor Wt1 is required for normal development of the olfactory system. Development 132,1327-1336[Abstract/Free Full Text]
  20. Maheswaran, S., Park, S., Bernard, A., Morris, J. F., Rauscher, F. J., 3rd, Hill, D. E., Haber, D. A. (1993) Physical and functional interaction between WT1 and p53 proteins. Proc. Natl. Acad. Sci. U. S. A. 90,5100-5104[Abstract/Free Full Text]
  21. Wang, W., Lee, S. B., Palmer, R., Ellisen, L. W., Haber, D. A. (2001) A functional interaction with CBP contributes to transcriptional activation by the Wilms tumor suppressor WT1. J. Biol. Chem. 276,16810-16816[Abstract/Free Full Text]
  22. Dame, C., Kirschner, K. M., Bartz, K. V., Wallach, T., Hussels, C. S., Scholz, H. (2006) Wilms tumor suppressor, Wt1, is a transcriptional activator of the erythropoietin gene. Blood 107,4282-4290[Abstract/Free Full Text]
  23. Wu, H., Liu, X., Jaenisch, R., Lodish, H. F. (1995) Generation of committed erythroid BFU-E and CFU-E progenitors does not require erythropoietin or the erythropoietin receptor. Cell 83,59-67[CrossRef][Medline]
  24. Lin, C. S., Lim, S. K., D'Agati, V., Costantini, F. (1996) Differential effects of an erythropoietin receptor gene disruption on primitive and definitive erythropoiesis. Genes Dev. 10,154-164[Abstract/Free Full Text]
  25. Komatsu, N., Fujita, H. (1993) Induced megakaryocytic maturation of the human leukemia cell line UT-7 results in down-modulation of erythropoietin receptor gene expression. Cancer Res. 53,1156-1161[Abstract/Free Full Text]
  26. Migliaccio, A. R., Jiang, Y., Migliaccio, G., Nicolis, S., Crotta, S., Ronchi, A., Ottolenghi, S., Adamson, J. W. (1993) Transcriptional and posttranscriptional regulation of the expression of the erythropoietin receptor gene in human erythropoietin-responsive cell lines. Blood 82,3760-3769[Abstract/Free Full Text]
  27. Migliaccio, A. R., Migliaccio, G., D'Andrea, A., Baiocchi, M., Crotta, S., Nicolis, S., Ottolenghi, S., Adamson, J. W. (1991) Response to erythropoietin in erythroid subclones of the factor-dependent cell line 32D is determined by translocation of the erythropoietin receptor to the cell surface. Proc. Natl. Acad. Sci. U. S. A. 88,11086-11090[Abstract/Free Full Text]
  28. Maouche, L., Cartron, J. P., Chretien, S. (1994) Different domains regulate the human erythropoietin receptor gene transcription. Nucleic Acids Res. 22,338-346[Abstract/Free Full Text]
  29. Zon, L. I., Youssoufian, H., Mather, C., Lodish, H. F., Orkin, S. H. (1991) Activation of the erythropoietin receptor promoter by transcription factor GATA-1. Proc. Natl. Acad. Sci. U. S. A. 88,10638-10641[Abstract/Free Full Text]
  30. Chiba, T., Ikawa, Y., Todokoro, K. (1991) GATA-1 transactivates erythropoietin receptor gene, and erythropoietin receptor-mediated signals enhance GATA-1 gene expression. Nucleic Acids Res. 19,3843-3848[Abstract/Free Full Text]
  31. Chin, K., Oda, N., Shen, K., Noguchi, C. T. (1995) Regulation of transcription of the human erythropoietin receptor gene by proteins binding to GATA-1 and Sp1 motifs. Nucleic Acids Res. 23,3041-3049[Abstract/Free Full Text]
  32. Dame, C., Sola, M. C., Fandrey, J., Rimsza, L. M., Freitag, P., Knopfle, G., Christensen, R. D., Bungert, J. (2002) Developmental changes in the expression of transcription factors GATA-1, -2 and -3 during the onset of human medullary haematopoiesis. Br. J. Haematol. 119,510-515[CrossRef][Medline]
  33. Orkin, S. H., Harosi, F. I., Leder, P. (1975) Differentiation in erythroleukemic cells and their somatic hybrids. Proc. Natl. Acad. Sci. U. S. A. 72,98-102[Abstract/Free Full Text]
  34. Haber, D. A., Park, S., Maheswaran, S., Englert, C., Re, G. G., Hazen-Martin, D. J., Sens, D. A., Garvin, A. J. (1993) WT1-mediated growth suppression of Wilms tumor cells expressing a WT1 splicing variant. Science 262,2057-2059[Abstract/Free Full Text]
  35. Dame, C., Fahnenstich, H., Freitag, P., Hofmann, D., Abdul-Nour, T., Bartmann, P., Fandrey, J. (1998) Erythropoietin mRNA expression in human fetal and neonatal tissue. Blood 92,3218-3225[Abstract/Free Full Text]
  36. Wagner, K. D., Wagner, N., Wellmann, S., Schley, G., Bondke, A., Theres, H., Scholz, H. (2003) Oxygen-regulated expression of the Wilms’ tumor suppressor Wt1 involves hypoxia-inducible factor-1 (HIF-1). FASEB J. 17,1364-1366[Abstract/Free Full Text]
  37. Elliott, S., Busse, L., Bass, M. B., Lu, H., Sarosi, I., Sinclair, A. M., Spahr, C., Um, M., Van, G., Begley, C. G. (2006) Anti-Epo receptor antibodies do not predict Epo receptor expression. Blood 107,1892-1895[Abstract/Free Full Text]
  38. Brown, W. M., Maxwell, P., Graham, A. N., Yakkundi, A., Dunlop, E. A., Shi, Z., Johnston, P. G., Lappin, T. R. (2007) Erythropoietin receptor expression in non-small cell lung carcinoma: a question of antibody specificity. Stem Cells 25,718-722[CrossRef][Medline]
  39. Kirkeby, A., van Beek, J., Nielsen, J., Leist, M., Helboe, L. (2007) Functional and immunochemical characterisation of different antibodies against the erythropoietin receptor. J. Neurosci. Methods 164,50-58[CrossRef][Medline]
  40. Dame, C. (2003) Molecular biology of the erythropoietin receptor in hematopoietic and non-hematopoietic tissues. Molineux, G. Foote, M. Elliott, S. G. eds. Erythropoietin and Erythropoiesis ,35-64 Birkhäuser Basel, Switzerland.
  41. Minguet, S., Cortegano, I., Gonzalo, P., Martinez-Marin, J. A., de Andres, B., Salas, C., Melero, D., Gaspar, M. L., Marcos, M. A. (2003) A population of c-Kit(low)(CD45/TER119)- hepatic cell progenitors of 11-day postcoitus mouse embryo liver reconstitutes cell-depleted liver organoids. J. Clin. Invest. 112,1152-1163[CrossRef][Medline]
  42. Dan, Y. Y., Riehle, K. J., Lazaro, C., Teoh, N., Haque, J., Campbell, J. S., Fausto, N. (2006) Isolation of multipotent progenitor cells from human fetal liver capable of differentiating into liver and mesenchymal lineages. Proc. Natl. Acad. Sci. U. S. A. 103,9912-9917[Abstract/Free Full Text]
  43. Goldberg, M. A., Glass, G. A., Cunningham, J. M., Bunn, H. F. (1987) The regulated expression of erythropoietin by two human hepatoma cell lines. Proc. Natl. Acad. Sci. U. S. A. 84,7972-7976[Abstract/Free Full Text]
  44. Wagner, K. D., Wagner, N., Sukhatme, V. P., Scholz, H. (2001) Activation of vitamin D receptor by the Wilms’ tumor gene product mediates apoptosis of renal cells. J. Am. Soc. Nephrol. 12,1188-1196[Abstract/Free Full Text]
  45. Kieran, M. W., Perkins, A. C., Orkin, S. H., Zon, L. I. (1996) Thrombopoietin rescues in vitro erythroid colony formation from mouse embryos lacking the erythropoietin receptor. Proc. Natl. Acad. Sci. U. S. A. 93,9126-9131[Abstract/Free Full Text]
  46. Broudy, V. C., Lin, N., Brice, M., Nakamoto, B., Papayannopoulou, T. (1991) Erythropoietin receptor characteristics on primary human erythroid cells. Blood 77,2583-2590[Abstract/Free Full Text]
  47. Sawyer, S. T., Krantz, S. B., Goldwasser, E. (1987) Binding and receptor-mediated endocytosis of erythropoietin in Friend virus-infected erythroid cells. J. Biol. Chem. 262,5554-5562[Abstract/Free Full Text]
  48. Philo, J. S., Aoki, K. H., Arakawa, T., Narhi, L. O., Wen, J. (1996) Dimerization of the extracellular domain of the erythropoietin (EPO) receptor by EPO: one high-affinity and one low-affinity interaction. Biochemistry 35,1681-1691[CrossRef][Medline]
  49. Svedberg, H., Richter, J., Gullberg, U. (2001) Forced expression of the Wilms tumor 1 (WT1) gene inhibits proliferation of human hematopoietic CD34(+) progenitor cells. Leukemia 15,1914-1922[Medline]
  50. Chen, D., Zhang, G. (2001) Enforced expression of the GATA-3 transcription factor affects cell fate decisions in hematopoiesis. Exp. Hematol. 29,971-980[CrossRef][Medline]
  51. Pandolfi, P. P., Roth, M. E., Karis, A., Leonard, M. W., Dzierzak, E., Grosveld, F. G., Engel, J. D., Lindenbaum, M. H. (1995) Targeted disruption of the GATA3 gene causes severe abnormalities in the nervous system and in fetal liver haematopoiesis. Nat. Genet. 11,40-44[CrossRef][Medline]
  52. Komatsu, N., Kirito, K., Kashii, Y., Furukawa, Y., Kikuchi, J., Suwabe, N., Yamamoto, M., Miura, Y. (1997) Cell-cycle-dependent regulation of erythropoietin receptor gene. Blood 89,1182-1188[Abstract/Free Full Text]
  53. Leonard, M., Brice, M., Engel, J. D., Papayannopoulou, T. (1993) Dynamics of GATA transcription factor expression during erythroid differentiation. Blood 82,1071-1079[Abstract/Free Full Text]
  54. Wu, Y., Fraizer, G. C., Saunders, G. F. (1995) GATA-1 transactivates the WT1 hematopoietic specific enhancer. J. Biol. Chem. 270,5944-5949[Abstract/Free Full Text]
  55. Zhang, X., Xing, G., Fraizer, G. C., Saunders, G. F. (1997) Transactivation of an intronic hematopoietic-specific enhancer of the human Wilms’ tumor 1 gene by GATA-1 and c-Myb. J. Biol. Chem. 272,29272-29280[Abstract/Free Full Text]
  56. Noguchi, C. T., Asavaritikrai, P., Teng, R., Jia, Y. (2007) Role of erythropoietin in the brain. Crit. Rev. Oncol. Hematol. 64,159-171[CrossRef][Medline]
  57. Fandrey, J. (2006) A cordial affair-erythropoietin and cardioprotection. Cardiovasc. Res. 72,1-2[Free Full Text]




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