Activation of Vitamin D Receptor by the Wilms' Tumor GeneProduct Mediates Apoptosis of Renal Cells : Journal of the American Society of Nephrology

Journal Logo

Molecular Medicine, Genetics, and Development

Activation of Vitamin D Receptor by the Wilms' Tumor GeneProduct Mediates Apoptosis of Renal Cells


Author Information
Journal of the American Society of Nephrology 12(6):p 1188-1196, June 2001. | DOI: 10.1681/ASN.V1261188
  • Free


The Wilms' tumor gene product, WT1, first was identified as a tumor suppressor by its mutational inactivation in a subset of Wilms' tumors (reviewed in reference 1). Wilms' tumor (nephroblastoma) is a childhood malignancy of the kidney caused by a failure of the metanephric mesenchyme to differentiate into glomeruli and tubules (2). WT1 gene expression is critical for genitourinary development, and homozygous disruption of wt1 in mice caused agenesis of the kidneys, likely as a result of a loss of metanephric blastemal cells (3).

WT1 is related structurally and functionally to the early growth response family of zinc finger transcription factors and originally was characterized as a transcriptional repressor (reviewed in reference 4). More recent studies indicate that WT1 also can stimulate gene transcription (5,6,7) and possibly in involved in RNA processing (8). Putative WT1 downstream targets include growth- and differentiation-promoting genes, such as insulin-like growth factor 2 (9), Pax2 (10), amphiregulin (11), and E-cadherin (12), among others (reviewed in reference 13). However, the endogenous transcript levels of most candidate genes with WT1-sensitive promoters failed to correlate with WT1 expression (14), suggesting that not all of the putative targets are normally controlled by WT1.

Expression of the WT1 gene is developmentally regulated in embryonic kidneys. WT1 mRNA and protein levels in the metanephric mesenchyme increase upon induction by the ureteric bud (15). Levels remain high in the renal vesicles and the comma- and S-shaped bodies, where WT1 expression is restricted to the podocyte layer of the differentiating glomeruli (15). Little is known about the physiologic function of WT1 during development. WT1 induced apoptosis in osteosarcoma cells by suppressing the synthesis of epidermal growth factor receptor (16). Because programmed cell death occurs at a large scale in embryonic kidneys (17) and because epidermal growth factor rescued the cultured renal mesenchyme from apoptosis (18), WT1 has been proposed to act as a proapoptotic signal in the developing kidney as well (16). However, direct experimental evidence that WT1 can indeed mediate programmed death of renal embryonic cells is still missing.

To characterize further the function of the WT1 transcription factor, we aimed to identify novel WT1 downstream target genes. Using the approach of educated guessing, we tested the hypothesis that vitamin D receptor (VDR) expression is regulated by WT1. VDR belongs to the steroid superfamily of nuclear receptors and mediates the genomic actions of the active metabolite 1,25-dihydroxyvitamin D3 (1,25-(OH)2D3). Upon binding to its ligand, VDR heterodimerizes with the retinoid X receptor (RXR), and this complex physically interacts with specific DNA recognition sites that are present in the regulatory regions of putative downstream target genes (reviewed in reference 19).

The following lines of evidence suggested to us that VDR might be a potential target for WT1. First, the promoters of the VDR genes from human and mouse contain several predicted WT1 consensus-binding sites (20,21). Second, a number of studies have shown that 1,25-(OH)2D3 can act as a signal for cell differentiation and also may induce apoptosis (reviewed in reference 22). Finally, VDR mRNA and protein have been detected in embryonic kidneys, thus pointing to a role for the vitamin D system in renal development (23).

Materials and Methods

Cell Culture and Transfections

Human embryonic kidney (HEK) 293 cells (ATCC CRL-1573) and HeLa cells (ATCC CRL-7923) were obtained from the American Type Culture Collection (ATCC). The cells were grown in Dulbecco's modified Eagle's medium (DMEM; Life Technologies, Eggenstein, Germany) supplemented with 10% fetal calf serum (FCS; Biochrom KG, Berlin, Germany), 100 IU/ml penicillin (Life Technologies), and 100 μg/ml streptomycin (Life Technologies). The cells were seeded at a density of 3 × 106 cells/100-mm dish on the day before the transfection. Mouse wt1 expression constructs with no genomic mutations (wt1 cDNA in pCB6+ (24)) were transfected into HEK 293 cells at 70% confluence with the use of the calcium phosphate precipitation technique (25). Stable clones were selected with 300 μg/ml G418 (Life Technologies) and expanded separately. Transient co-transfections were performed with the indicated amounts of plasmids and 1 μg of a cytomegalovirus promoter-driven β-galactosidase expression construct as an internal control for transfection efficiency.

Luciferase and β-Galactosidase Assays

The cells were lysed 48 h after the transfection, and luciferase activities were measured in a luminometer (Lumat LB 9501, Bertholdt, Germany) with the use of beetle luciferin as a substrate (Promega, Mannheim, Germany). β-galactosidase activities were determined spectrophotometrically (Beckman DU 540 spectrophotometer) with the use of a commercial kit according to the manufacturer's instructions (Promega). Results shown are averages of five transfection experiments, each performed in duplicate. P < 0.05 was considered significant (ANOVA).

Determination of Cell Proliferation

Cell proliferation was estimated by counting aliquots of the trypsinized cells in a Neubauer chamber and by measuring 5-bromo-2′-deoxy-uridine incorporation into genomic DNA followed by an enzyme-linked immunosorbent assay detection (Roche Diagnostics, Mannheim, Germany). For this purpose, HEK 293 cells stably transfected either with a wt1 expression construct or with the empty pCB6+ vector were seeded into 96-well plates at a density of 104 cells per cm2 with variable concentrations of 1,25-(OH)2D3 in the medium. The cultures were incubated for 4 d, and the medium containing 1,25-(OH)2D3 was renewed daily. Data presented are means ± SEM of 5 experiments performed as duplicates.

Measurement of 1,25-(OH)2D3 Concentrations

Concentrations of 1,25-(OH)2D3 were measured in tissue culture supernatants at different time points (4, 8, 12, 16, and 24 h) after incubation. The supernatants were delipidated by treatment with a dextran sulfate/magnesium chloride reagent, followed by immunoextraction with a monoclonal anti-1,25-(OH)2D3 antibody (Immunodiagnostics Ltd., Hamburg, Germany). 1,25-(OH)2D3 concentrations in the extracts were measured by RIA as described by the manufacturer (Immunodiagnostics Ltd.).

RNA Preparation and Northern Blot Hybridization

Isolation of total RNA and Northern blot hybridization were performed as described elsewhere (9). Equal amounts of total RNA were pooled from stable clones grown separately.

Sodium Dodecyl Sulfate—Polyacrylamide Gel Electrophoresis

Total cell lysates from subconfluent cultures were prepared by heating the samples in TBS/1% sodium dodecyl sulfate (SDS) buffer to 95°C for 3 min. Twenty μg of protein were loaded per lane and transferred after separation on a 10% SDS—polyacrylamide gel electrophoresis onto polyvinylidenedifluoride membranes (Amersham Pharmacia Biotech, Freiburg, Germany) with the use of a semidry blotting apparatus (BioRad, Mu[Combining Diaeresis]nchen, Germany). A polyclonal anti-WT1 antibody from rabbit (WT 180; Santa Cruz Biotechnology, Heidelberg, Germany) was used at a 1:1000 dilution for immunoblotting. After incubation with a goat anti-rabbit secondary antibody (1:5000), the reaction products were detected by enhanced chemoluminescence system (Amersham Pharmacia Biotech). A rat monoclonal anti-VDR antibody (1:500 dilution; Affinity Bioreagents, Hamburg, Germany) and a biotinylated goat anti-rat IgG secondary antibody (1:1000 dilution; Dianova, Hamburg, Germany) were used for immunodetection of VDR protein with the biotin-streptavidin technique.

WT1 Immunostaining of Cultured Mouse Renal Embryonic Cells

After fixation with 3% paraformaldehyde in phosphate-buffered saline, the cells were permeabilized with 0.1% Triton X-100 and endogenous peroxidase activities were blocked for 5 min in a solution of 3% H2O2 in methanol (1:4). After washing in TBS, the cells were incubated for 16 h at 4°C with a rabbit polyclonal anti-WT1 antibody (Santa Cruz Biotechnology) diluted 1:150 in TBS with 5% normal goat serum. This incubation was followed by a 2-h treatment with biotinylated secondary antibody (goat anti-rabbit, 1:100 in TBS with 1% bovine serum albumin; Vector Laboratories Inc.) and the streptavidin-peroxidase complex (Sigma, Deisenhofen, Germany). WT1-positive cells were identified by their brown color after visualization with diaminobenzidine and hydrogen peroxide (Sigma).

Construction of Reporter Plasmids

A 1451-bp fragment from the mouse vdr promoter (from 70 to 1521 bp of the published sequence (20)) was cloned by PCR with the use of genomic DNA from mouse liver as a template and the following primers: 5′-TGCCCTAAGGTGTTGGCT-3′ (forward primer), 5′-TGGACACACAGCTCGGCG-3′ (reverse primer). The PCR product was ligated into the SmaI/BglII restriction sites of the pGL2basic reporter plasmid and confirmed by dideoxy sequencing of both strands. This construct was designated pVDR1451. 5′-deletion mutants were generated from pVDR1451 by PCR with the use of the following forward primers: 5′-CGGATCATCACAGGCAGA-3′, 5′-TTGAGGGGCAGGGCGGTC-3′, 5′-CCAGGTGCTGAGCAGTCT-3′. The PCR products were cloned into the KpnI/BglII sites of pGL2basic and designated pVDR306, pVDR105, and pVDR60, respectively.

Electrophoretic Mobility Shift Assays

Mouse recombinant WT1 protein was generated in the presence of [35S]methionine with the use of the TNT quick coupled transcription-translation system (Promega). An aliquot of the translation product was analyzed on a 10% SDS-polyacrylamide gel, which showed a single band of the size of WT1 protein (approximately 52 kD). DNA binding reactions were performed for 30 min at room temperature with 20 ng of the unlabeled recombinant WT1 protein (+ and - KTS isoforms) in 10 μl of a 1 × reaction buffer (10 mM Tris-HCl [pH 7.5], 50 mM NaCl, 1 mM MgCl2, 0.5 mM ethylenediaminetetraacetate, 0.5 mM DTT, 14% glycerol, 0.05 mg/ml poly [dI-dC]). For the supershift experiments, 1 μg of a polyclonal rabbit anti-WT1 antibody (C19; Santa Cruz Biotechnology) was added to the reaction mixture. The end-labeled 21-bp double-stranded oligonucleotide (5′-TGAACTTAGTGGGCGTGGTTG-3′) contained the predicted WT1 element from the proximal mouse vdr promoter. A 28-bp DNA fragment including the WT1 consensus binding site from the platelet-derived growth factor A-chain promoter served as a specific competitor (5′-GGGGCGGGGGCGGGGGCGGGGGAGGGG-3′) (26).

Induction and Analysis of Apoptosis in Cultured Mouse Renal Embryonic Cells

Primary cultures of kidney cortical cells were prepared from mouse embryos on gestational day 12 (E12) by an enzymatic digestion procedure as described previously (27) and grown for 1 d in DMEM-10% FCS. The viable cells were split into 96-well plates at a density of 103 cells/cm2 and cultured for 72 h in DMEM-10% FCS supplemented with variable concentrations (1 nM to 1 μM nominally) of 1,25-(OH)2D3. The active vitamin D compound (Leo Pharmaceutical, Ballerup, Denmark), which was obtained as a 4 × 10-3 M stock solution, was diluted to 10-6 M in ethanol. The final ethanol content in the tissue culture medium was ≤0.1 vol %. Control experiments were performed by incubation of the cells for 72 h with the appropriate amounts of ethanol (in DMEM-10% FCS). The following techniques were applied to identify apoptotic cells: (1) the characteristic signs of apoptosis, including membrane blebbing, cytoplasmic shrinkage, and nuclear condensation, were observed at 400× magnification after hematoxylin and eosin staining of the cultures (Axioplan 2; Zeiss, Go[Combining Diaeresis]ttingen, Germany); (2) free 3′-OH termini in apoptotic cells were fluorescence-labeled with the terminal deoxynucleotidyl transferase (TdT) technique with the use of a commercial kit according to the manufacturer's instructions (Roche Diagnostics); (3) apoptotic cells were stained with a FITC-conjugated anti-annexin V antibody (Roche Diagnostics) and visualized under a fluorescence microscope (400× magnification, Axioplan, Zeiss). The annexin V immunopositive cells were counted in 10 optical fields from two dishes each. Only those annexin V—positive cells that did not stain with propidium iodide were considered apoptotic. The fraction of cells that exhibited annexin V immunoreactivity was calculated in five independent experiments and taken as an estimate for the proapoptotic action of 1,25-(OH)2D3.


Constitutive WT1 Expression in HEK 293 Cells

To identify novel putative downstream target genes of the WT1 transcription factor, we established a human embryonic kidney cell line (HEK 293) with constitutive wt1 expression. The transfected cDNA corresponded to the most abundant wt1 isoform containing a 17 amino acid insertion from alternatively spliced exon 5 and three additional amino acids (KTS) between zinc fingers 3 and 4 of the WT1 molecule (28). Renal cells were chosen because the fetal kidney is a physiologic site of WT1 expression and because low levels of the endogenous WT1 mRNA were transcribed in HEK 293 cells (29), suggesting that the transacting factors required for normal WT1 expression are operating in this cell line. No WT1 protein was detectable by immunoblotting in HEK 293 cells that were transfected stably with the empty pCB6+ expression vector (Figure 1A). WT1 protein levels in pooled samples (n = 5 different clones) of the wt1 transfectants were approximately fourfold higher (by densitometry) than in embryonic rat kidney on gestational day 18 (E18). Taking into account that wt1 is expressed only in a small percentage of cells in E18 kidneys (15), the average amount of wt1 protein per cell therefore may be approximately the same in our wt1-HEK 293 transfectants and in a restricted cell population in the developing kidney in vivo.

Figure 1:
. Expression of Wilms' tumor transcription factor WT1 and vitamin D receptor (VDR) in human embryonic kidney (HEK) 293 cells. (A) Immunoblot demonstrating WT1 protein in HEK 293 cells that were transfected stably either with a mouse wt1 expression construct (WT1) or with the empty pCB6+ expression vector (pCB6+), respectively. WT1 protein levels (approximately 52 kD-band) were only slightly higher in pooled samples (n = 5) of wt1-transfected HEK 293 cells than in kidneys from rat embryos on gestational day 18 (E18). VDR mRNA (B) and protein (D) levels in pools (n = 5) of wt1- and pCB6+-transfected HEK 293 cells. Note the different intensities (fivefold by densitometry) of the specific Northern blot hybridization signals between the wt1-expressing and the pCB6+-transfected cells (B). Equal amounts of β-actin transcripts were detected in the two cell populations (C). The increase of VDR mRNA in wt1-expressing HEK 293 cells was mirrored by increased VDR protein levels (D). HeLa cells, which also expressed the approximately 50-kD VDR protein, served as positive controls (D).

VDR Expression in Transfected HEK 293 Cells

We performed Northern blot hybridization using a full-length human cDNA probe to compare VDR mRNA levels in wt1- and pCB6+-transfected clones. As shown in Figure 1B, VDR mRNA content was significantly higher (fivefold by densitometry) in wt1-expressing than in pCB6+-transfected cells. The increase of VDR mRNA in wt1-HEK 293 was associated with upregulation of VDR protein as indicated by immunoblot analysis with a rat monoclonal anti-VDR antibody (Figure 1D). β-Actin transcript levels were approximately the same in the wt1- and pCB6+-transfected clones (Figure 1C). Because HEK 293 cells that were transfected stably either with wt1 cDNA (in pCB6+ plasmid) or with pCB6+ empty vector shared a similar genetic background, the observed differences in VDR gene expression likely were due to the different WT1 levels.

Activation of the VDR Gene Promoter by WT1

To test whether transcription from the vdr promoter could be activated by wt1, we transiently co-transfected HEK 293 cells with wt1 expression vectors (four different splicing variants) and a construct containing approximately 1.5 kb of mouse vdr promoter sequence (20) upstream of the firefly luciferase reporter gene (pVDR1451). Transient co-transfections were performed because the permanent lines were extremely difficult to transfect, yielding transfection efficiencies of less than 5%. Normalized luciferase activities that were taken as a measure for the transcriptional activity of the vdr promoter are indicated in Figure 2. Co-transfection of pVDR1451 (2 μg) along with the different wt1 expression constructs (15 μg each) stimulated vdr promoter activity more than fourfold. Luciferase activity of pVDR1451 increased in proportion with the amount (2 to 15 μg) of co-transfected wt1 expression plasmid (data not shown). In contrast, transcription from the vdr promoter was not changed significantly by co-transfection of a pCB6+ construct that contained the full-length wt1 cDNA in antisense (WT1 rev.) orientation (Figure 2A). Serial 5′-deletion analysis detected a 201-bp fragment in the proximal vdr promoter that was required for transcriptional activation by wt1 (Figure 2B). This sequence contained a predicted WT1 consensus site (-GNGGGNGNG-) 115 bp upstream of exon 1 (20). Electrophoretic mobility shift assays were performed to examine whether recombinant wt1 protein (+ and - KTS isoforms) could physically interact with the putative WT1 element. Using the 21-bp oligonucleotide from the mouse vdr promoter as a probe (5′-TGAACTTAGTGGGCGTGGTTG-3′), we obtained a single retardation band with both wt1 isoforms (Figure 3). A similar retardation signal was seen with nuclear extracts prepared from the permanent wt1-expressing clones (data not shown). The shifted band could be competed with excess amounts of a 28-bp unlabeled oligonucleotide including a WT1 binding site from the platelet-derived growth factor A-chain promoter (26). The retardation band was supershifted upon incubation of the reaction mixture with a polyclonal rabbit anti-WT1 antibody (Figure 3). No band shift was obtained with an oligonucleotide containing an Sp1 instead of a WT1 consensus binding site (data not shown).

Figure 2:
. WT1 activates the vdr promoter in HEK 293 cells. (A) Effect of transient co-transfection of expression constructs encoding four different wt1 splicing variants (±17 amino acids/±KTS) on the transcriptional activity of an approximately 1.5-kb mouse vdr promoter construct (pVDR1451). Shown are the relative luciferase activities normalized to β-galactosidase activity in each sample. Values are means ± SEM of five experiments in duplicate. *, statistical significances (P < 0.05) versus transfection of the vdr promoter construct (pVDR1451) alone; WT1 rev., a pCB6+ construct containing the full-length wt1 cDNA in reverse orientation. (B) Serial 5′-deletion analysis of the mouse vdr promoter. Promoter constructs of various lengths were co-transfected transiently into HEK 293 cells either along with empty pCB6+ expression plasmid (□) or with the wt1 cDNA (+KTS isoform) in pCB6+ (▪). The relative lengths of the vdr promoter constructs and the locations of the putative WT1 consensus binding sites are indicated in the schematic drawing. Normalized luciferase activities are expressed as relative light units (RLU). Values shown are averages ± SEM of five experiments performed in duplicate. *, statistical significance (P < 0.05) between wt1- and pCB6+-transfected cells. Note that deletion of a 201-bp sequence (pVDR306 versus pVDR105) abolished the wt1 responsiveness of the vdr promoter.
Figure 3:
. Electrophoretic mobility shift assay performed with 20 ng of recombinant wt1 protein (+ and - KTS isoforms) and a 21-bp double-stranded sequence containing the predicted WT1 consensus site from the proximal mouse vdr promoter. A 28-bp unlabeled oligonucleotide including a WT1 binding site in the platelet-derived growth factor A-chain promoter (27) was used at the indicated molar excess in competition experiments. Supershift assays were performed by incubating the binding reactions with a polyclonal anti-WT1 antibody (pAb).

1,25-(OH)2D3 Inhibits the Proliferation of HEK 293 Cells

To test whether renal cell growth might be controlled by a vitamin D—dependent signaling mechanism, we studied the effect of 1,25-(OH)2D3 on the proliferation of pCB6+- and wt1-transfected HEK 293 cells. As shown in Figure 4A, the basal proliferation rates were significantly lower in the wt1-expressing lines as compared with the pCB6+-transfected cells. Incubation with the active metabolite 1,25-(OH)2D3 dose-dependently inhibited 5-bromo-2′-deoxy-uridine incorporation into chromosomal DNA. A comparison between the slopes of the growth curves in Figure 4B reveals that the wt1-expressing HEK 293 cells were by far more sensitive to the antiproliferative action of 1,25-(OH)2D3 than the pCB6+ transfectants. Notably, the 1,25-(OH)2D3 concentrations measured in the tissue culture supernatants after 24 h of incubation were approximately 15-fold lower than those nominally adjusted in the medium. This finding is consistent with an estimated in vitro half-life of 1,25-(OH)2D3 of approximately 6 h (30).

Figure 4:
. (A) Proliferation of HEK 293 cells that were transfected stably either with a wt1 expression construct (WT1) or with the empty pCB6+ expression plasmid. The cells were seeded into 96-well plates at an initial density of 104 cells/cm2 and grown for 3 d in Dulbecco's modified Eagle's medium (DMEM)-10% fetal calf serum (FCS). Cells from two wells were harvested each day and counted in a Neubauer chamber to estimate cell proliferation. Values are means ± SEM of five experiments. *, statistically significant differences (P < 0.05) between wt1- and pCB6+-transfected cells. (B) 1,25-dihydroxyvitamin D3 (1,25-(OH)2D3) inhibits the proliferation of HEK 293 cells. The stable cell lines were grown for 4 d with the indicated concentrations of 1,25-(OH)2D3 nominally adjusted in the tissue culture medium. Cell proliferation was assayed by measurement of 5-bromo-2′-deoxy-uridine incorporation with an enzyme-linked immunosorbent assay technique. Values presented are means ± SEM of five experiments performed in duplicate. Symbols indicate statistical significance (P < 0.05) within (*) and between (+) the wt1- and pCB6+-transfected lines, respectively. Incubation of stable HEK 293 cells with vehicle alone (0.1% ethanol in DMEM-10% FCS) had no significant effect on proliferation rates (data not shown).

1,25-(OH)2D3 Induces Apoptosis of Wt1-Expressing Renal Embryonic Cells

Because the active vitamin D metabolite can induce apoptosis in a variety of cells (reviewed in reference 22), we examined whether programmed cell death also was involved in the action of 1,25-(OH)2D3 on embryonic renal cells. As a more physiologic model than the permanent HEK 293 cell lines, we used renal cells that were freshly isolated from mouse embryonic kidney cortex. These primary cultures consisted of more than 80% wt1-immunopositive cells, a significant fraction of which exhibited the signs of apoptosis (e.g., membrane blebbing, cytoplasmic shrinkage, chromatin condensation) after a 72-h incubation with 1 nM 1,25-(OH)2D3 (Figure 5DversusFigure 5C). Programmed cell death was confirmed by the demonstration of nuclear DNA fragmentation with the use of in situ DNA nick end labeling assay (Figure 5FversusFigure 5E) and by annexin V immunocytochemistry. The effect of the active vitamin D metabolite was dose-dependent, and the fraction of annexin V—positive cells was 45 ± 6%, 62 ± 5%, 73 ± 6%, and 82 ± 4% in the presence of 1 nM, 10 nM, 100 nM, and 1 μM 1,25-(OH)2D3, respectively. Of note, the active vitamin D metabolite caused apoptosis preferentially of the wt1-expressing renal embryonic cells, and—by consequence—the fraction of wt1-immunonegative cells increased from less than 20% to more than 90% after a 72-h exposure to 1,25-(OH)2D3 (Figure 5BversusFigure 5A).

Figure 5:
. 1,25-(OH)2D3 induces apoptosis in primary cultures of mouse renal embryonic cells. Isolated cells were grown for 3 d in DMEM-10% FCS containing either 1 nM 1,25-(OH)2D3 or 0.1% ethanol as a vehicle (five experiments each performed in duplicate). Representative microscopy of cells from mouse embryonic kidney cortex after a 72-h incubation period either with 1,25-(OH)2D3 (B, D, F) or with 0.1% ethanol as a vehicle (A, C, E), respectively. WT1-positive cells (brown color) were detected with the use of a polyclonal anti-WT1 antibody followed by immunoperoxidase labeling with an indirect biotin-streptavidin technique. Note that the fraction of wt1-immunopositive cells decreased dramatically (B versus A) after 1,25-(OH)2D3 treatment. The characteristic signs of apoptosis including DNA condensation became visible after hematoxylin and eosin staining of the 1,25-(OH)2D3-treated cultures (arrows in D). Apoptosis was confirmed by end-labeling of free 3′-OH termini in cultures incubated with the active vitamin D3 metabolite (F). Significantly fewer apoptotic cells were seen in the controls (E). Magnifications: ×200 in A and B; ×400 in C and D; ×100 in E and F.

VDR and WT1 Expression in the Developing Kidney

Time courses of wt1 and vdr expression were studied by Northern blot analysis in the developing rat kidney in vivo. Intrarenal wt1 and vdr transcript levels were closely related between days 15 and 21 of embryonic development (Figure 6). Whereas vdr mRNA levels remained high throughout adulthood, wt1 transcripts were barely detectable in kidneys from adult rats Figure 6).

Figure 6:
. WT1 and vdr mRNA levels at different time points in developing and adult rat kidneys. Total kidney RNA was prepared from 13 animals each, pooled at equal aliquots, and loaded (20 μg/lane) on a 1.2% agarose-formaldehyde gel. Note the striking correlation of wt1 and vdr expression between days 15 (E15) and 21 (E21) of embryonic development. The same membrane first was hybridized with a full-length wt1 cDNA probe and then used for rehybridization with a 32P-labeled rat vdr cDNA. Ethidium bromide staining indicates roughly equal loading of the RNA gel (bottom).


The purpose of this study was to analyze the downstream signaling mechanisms of the Wilms' tumor transcription factor WT1 in embryonic renal cells. We report the following novel observations:

  1. VDR mRNA and protein levels were upregulated in HEK 293 cells with constitutive WT1 expression.
  2. WT1 transactivated the mouse vdr promoter probably through direct interaction with a predicted consensus binding site.
  3. WT1 significantly increased the susceptibility of HEK 293 cells to the antiproliferative action of 1,25-(OH)2D3.
  4. The active vitamin D metabolite induced apoptosis in primary cultures of wt1-expressing renal embryonic cells.
  5. The temporal expression patterns of wt1 and vdr genes correlated closely in the developing kidney. These findings suggest that transcriptional activation of the VDR by the WT1 zinc finger protein can mediate growth inhibition and apoptosis of renal embryonic cells.

Recent studies have shown that the active metabolite 1,25-(OH)2D3 acting through the intracellular VDR can promote cell differentiation in addition to its effects on calcium and phosphate metabolism (reviewed in references 19 and 22). Interestingly, some of the molecular targets of VDR, e.g., epidermal growth factor receptor (31) and cyclin-dependent kinase inhibitor p21 (32), also are regulated by the WT1 transcription factor (6,16), indicating that WT1 and VDR act through common intracellular signal transduction pathways. Functional synergism between WT1 and VDR is supported by our observation that WT1 stimulated VDR expression in human embryonic kidney cells. Upregulation of VDR by WT1 most likely occurred at the transcriptional level, because co-transfection of WT1 expression constructs stimulated vdr promoter activity probably through direct interaction of WT1 protein with a predicted consensus sequence in the proximal vdr promoter.

The transcriptional effect of WT1 may depend on the type of expression vector used for transfection (33). Because we transfected cytomegalovirus promoter—based constructs that have been found to repress rather than activate gene transcription (33), it might even underestimate the stimulatory potency of wt1 in our experimental setting. For comparison, a threefold induction of the human forkhead gene promoter (34) and a fivefold activation of the syndecan-1 promoter (5) by WT1 have been reported. The exact mechanism for the dual regulatory functions of WT1 that can either repress or enhance gene transcription is still unclear but may involve specific protein—protein interactions (35,36).

The metanephric blastema and the podocyte precursors of the immature glomeruli are major sites of wt1 expression in the developing kidney (15). Vdr immunopositive cells also have been detected recently in the renal mesenchyme and the visceral and parietal glomerular epithelium of embryonic rat kidney (23). Our findings demonstrate that the temporal expression patterns of wt1 and vdr correlate closely during renal development. It seems likely, therefore, that VDR is a physiologic target gene for WT1 not only in vitro but also in the developing kidney in vivo.

The functional significance of our findings is supported by the observation that the growth inhibitory effect of 1,25-(OH)2D3 was enhanced dramatically in wt1-expressing HEK 293 cells. Accordingly, less than nanomolar concentrations of the active vitamin D metabolite 1,25-(OH)2D3 induced programmed death predominantly of the wt1-immunopositive cells from mouse embryonic kidney cortex. These observations suggest that the sensitivity to the proapoptotic action of 1,25-(OH)2D3 was enhanced in the wt1-expressing cells. Apoptosis seems to be particularly important in renal development as the kidneys were the most severely affected organ in Bcl2 knockout mice (37), and approximately 3% of all cells within the nephrogenic renal cortex are apoptotic at any given time during development (17). Our results suggest that the vitamin D endocrine system contributes to the high rate of apoptosis in the developing kidney. Because 1,25-(OH)2D3 may induce programmed cell death through a VDR-dependent signaling mechanism (38), upregulation of VDR by the WT1 transcription factor therefore would increase the susceptibility of renal embryonic cells to the proapoptotic action of vitamin D metabolite. Similar results have found recently with breast cancer cells, which exhibited increased rates of apoptosis in response to chemotherapeutic agents after pretreatment with 1,25-(OH)2D3 (39).

A role of the vitamin D endocrine system in renal development is seemingly in conflict with the normal embryonic development of vdr-/- mice (40,41). Surprisingly, the vdr null mutant animals were phenotypically normal at birth and did not exhibit the signs of vitamin D deficiency until weaning. After weaning, however, vitamin D—dependent rickets type II and growth retardation developed in homozygous mice, leading to death within 15 wk after birth (40,41). The normal phenotype of the vdr null mice does not exclude a role of the vitamin D system in renal cell differentiation but rather suggests a redundancy in vitamin D—dependent signaling throughout embryonic and early postnatal development. Similar findings were made recently with amphiregulin, a member of the epidermal growth factor family, that has been identified as a transcriptional target of WT1 (11). Thus, amphiregulin stimulated the branching morphogenesis of metanephric kidney explants (11), but no gross developmental defects were observed in kidneys from amphiregulin knockout mice (42).

Taken together, our findings demonstrate that the WT1 gene product transcriptionally activates VDR expression in human embryonic kidney cells. Upregulation of VDR by the WT1 transcription factor may mediate apoptosis of renal embryonic cells in response to 1,25-(OH)2D3. These findings suggest a role for the vitamin D endocrine system in the regulation of renal cell growth and differentiation during development.

The authors appreciate the expert technical assistance of A. Richter and I. Gra[Combining Diaeresis]tsch. We thank Dr. F. Priem for the measurement of 1,25-(OH)2D3 and Dr. G. Walz for critical reading of the manuscript. The wt1 expression constructs were a gift from Dr. D. Haber, and 1α,25(OH)2D3 was kindly provided by Dr. L. Binderup (Leo Pharmaceutical Products). This study was financially supported by a grant from the Deutsche Forschungsgemeinschaft (Scho 634/3-1).

1. Hastie ND: The genetics of Wilms' tumor—A case of disrupted development. Annu Rev Genet 28:523 -558, 1994
2. Beckwith JB, Kiviat NB, Bonadio JF: Nephrogenic rests, nephroblastomatosis, and the pathogenesis of Wilms' tumor. Pediatr Pathol 10: 1-36,1990
3. Kreidberg JA, Sariola H, Loring JM, Maeda M, Pelletier J, Housman D, Jaenisch R: WT-1 is required for early kidney development. Cell 74:679 -691, 1993
4. Rauscher FJ III: The WT1 Wilms tumor gene product: A developmentally regulated transcription factor in the kidney that functions as a tumor suppressor. FASEB J 7:896 -903, 1993
5. Cook DM, Hinkes MT, Bernfield M, Rauscher FJ III: Transcriptional activation of the syndecan-1 promoter by the Wilms' tumor protein WT1. Oncogene 13:1789 -1799, 1996
6. Englert C, Maheswaran S, Garvin AJ, Kreidberg J, Haber DA: Induction of p21 by the Wilms' tumor suppressor gene WT1. Cancer Res 57:1429 -1434, 1997
7. Guan LS, Rauchman M, Wang ZY: Induction of Rb-associated protein (RbAp46) by Wilms' tumor suppressor WT1 mediates growth inhibition. J Biol Chem 273:27047 -27050, 1998
8. Larsson SH, Charlieu JP, Miyagawa K, Engelkamp D, Rassoulzadegan M, Ross A, Cuzin F, van Heyningen V, Hastie ND: Subnuclear localization of WT1 in splicing or transcription factor domains is regulated by alternative splicing. Cell 81:391 -401, 1995
9. Drummond IA, Madden SL, Rohwer-Nutter P, Bell GI, Sukhatme VP, Rauscher FJ III: Repression of the insulin-like growth factor gene by the Wilms tumor suppressor WT1. Science257 : 674-678,1992
10. Ryan G, Steele-Perkins V, Morris JF, Rauscher FJ III, Dressler G: Repression of Pax-2 by WT1 during normal kidney development. Development 121:867 -875, 1995
11. Lee SB, Huang K, Palmer R, Truong VB, Herzlinger D, Kolquist KA, Wong J, Paulding C, Yoon SK, Gerald W, Oliner JD, Haber DA: The Wilms tumor suppressor WT1 encodes a transcriptional activator of amphiregulin. Cell 98:663 -673, 1999
12. Hosono S, Gross I, English MA, Hajra KM, Fearon ER, Licht JD: E-cadherin is a WT1 target gene. J Biol Chem275 : 10943-10953,2000
13. Reddy JC, Licht JD: The WT1 Wilms' tumor suppressor gene: How much do we really know? Biochim Biophys Acta1287 : 1-28,1996
14. Tha[Combining Diaeresis]te C, Englert C, Gessler M: Analysis of WT1 target gene expression in stably transfected cell lines. Oncogene 17:1287 -1294, 1998
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 ND: The candidate Wilms' tumour gene is involved in genitourinary development. Nature 346:194 -197, 1990
16. Englert C, Hou X, Maheswaran S, Bennett P, Ngwu C, Re GG, Garvin AJ, Rosner MR, Haber DA: WT1 suppresses synthesis of the epidermal growth factor receptor and induces apoptosis. EMBO J14 : 4662-4675,1995
17. Coles HSR, Burne JF, Raff MC: Large scale normal cell death in the developing rat kidney and its reduction by epidermal growth factor. Development 118:777 -784, 1993
18. Koseki C, Herzlinger D, Al-Awqati Q: Apoptosis in metanephric development. J Cell Biol 119:1327 -1333, 1992
19. Haussler MR, Whitfield GK, Haussler CA, Hsieh JC, Thompson PD, Selznick SH, Dominguez CE, Jurutka PW: The nuclear vitamin D receptor: Biological and molecular regulatory properties revealed. J Bone Miner Res 13:325 -349, 1998
20. Jehan F, DeLuca HF: Cloning and characterization of the mouse vitamin D receptor promoter. Proc Natl Acad Sci USA94 : 10138-10143,1997
21. Miyamoto KI, Kesterson RA, Yamamoto H, Taketani Y, Nishiwaki E, Sawako T, Inoue Y, Morita K, Takeda E, Pike JW: Structural organization of the human vitamin D receptor chromosomal gene and its promoter. Mol Endocrinol 11:1165 -1179, 1997
22. Carlberg C, Polly P: Gene regulation by vitamin D3. Crit Rev Eukaryot Gene Expr 8:19 -42, 1998
23. Johnson JA, Grande JP, Roche PC, Sweeney WE Jr, Avner ED, Kumar R: 1α,25-dihydroxyvitamin D3: Receptor ontogenesis in fetal renal development. Am J Physiol38 : F419-F428,1995
24. Buckler AJ, Pelletier J, Haber DA, Glaser T, Housman DE: The murine Wilms' tumor gene (WT1): Isolation, characterization, and expression during kidney development. Mol Cell Biol11 : 1707-1712,1991
25. Gorman C, Moffat LF, Howard BH: Recombinant genomes which express chloramphenicol acetyltransferase in mammalian cells. Mol Cell Biol 2: 1044-105,1982
26. Wang ZY, Qiu QQ, Huang J, Gurrieri M, Deuel TF: WT1, the Wilms' tumor suppressor gene product, represses transcription through an interactive nuclear protein. Oncogene 10:415 -422, 1995
27. Drummond IA, Mukhopadhyay D, Sukhatme VP: Expression of fetal kidney growth factors in a kidney tumor line: Role of FGF2 in kidney development. Exp Nephrol 6:522 -533, 1998
28. Haber DA, Sohn PL, Buckler AJ, Pelletier J, Call KM, Housman DE: Alternative splicing and genomic structure of the Wilms tumor gene WT1. Proc Natl Acad Sci USA 88:9618 -9622, 1991
29. Scholz H, Bossone SA, Cohen HT, Akella U, Strauss WM, Sukhatme VP: A far upstream cis-element is required for Wilms' tumor-1 (WT1) gene expression in renal cell culture. J Biol Chem272 : 32836-32846,1997
30. Satchell DP, Norman AW: Metabolism of the cell differentiating agent 1alpha,25(OH)2-16-ene-23-yne vitamin D3 by leukemic cells. J Steroid Biochem Mol Biol57 : 117-124,1996
31. Petkovich PM, Wrana JL, Grigoriadis AE, Heersche JN, Sodek J: 1,25-dihydroxyvitamin D3: increases epidermal growth factor receptors and transforming growth factor beta-like activity in a bone-derived cell line. J Biol Chem 262:13424 -13428, 1987
32. Liu M, Lee MM, Cohen M, Bommakanti M, Freedman LP: Transcriptional activation of the Cdk inhibitor p21 by vitamin D3 leads to the induced differentiation of the myelomonocytic cell line U937. Genes Dev 10: 142-153,1996
33. Reddy JC, Hosono S, Licht JD: The transcriptional effect of WT1 is modulated by choice of expression vector. J Biol Chem270 : 29976-29982,1995
34. Ernstsson S, Pierrou S, Hulander M, Cederberg A, Hellqvist M, Carlsson P, Enerba[Combining Diaeresis]ck S: Characterization of the human forkhead gene FREAC-4. J Biol Chem271 : 21094-21099,1996
35. Johnstone RW, See RH, Sells SF, Wang J, Muthukkumar S, Englert C, Haber DA, Licht JD, Sugrue SP, Roberts T, Rangnekar VM, Shi Y: A novel repressor, par-4, modulates transcription and growth suppression functions of the Wilms' tumor suppressor WT1. Mol Cell Biol16 : 6945-6956,1996
36. Maheswaran S, Park S, Bernard A, Morris JF, Rauscher FJ, Hill DE, Haber DA: Physical and functional interaction between WT1 and p53 proteins. Proc Natl Acad Sci USA 90:5100 -5104, 1993
37. Veis D, Sorenson C, Shutter J, Korsmeyer S: Bcl-2-deficient mice demonstrate fulminant lymphoid apoptosis, polycystic kidneys, and hypopigmented hair. Cell 75:229 -240, 1993
38. Davoust N, Wion D, Chevalier G, Garabedian M, Brachet P, Couez D: Vitamin D receptor stable transfection restores the susceptibility to 1,25-dihydroxyvitamin D3 cytotoxicity in a rat glioma resistant clone. J Neurosci Res 52:210 -219, 1998
39. Wang Q, Yang W, Uytingco MS, Christakos S, Wieder R: 1,25-Dihydroxyvitamin D3: and all-trans-retinoic acid sensitize breast cancer cells to chemotherapy-induced cell death. Cancer Res 60:2040 -2048, 2000
40. Li YC, Pirro AE, Amling M, Delling G, Baron R, Bronsson R, Demay MB: Targeted ablation of the vitamin D receptor: An animal model of vitamin D-dependent rickets type II with alopecia. Proc Natl Acad Sci USA 94:9831 -9835, 1997
41. Yoshizawa T, Handa Y, Uematsu Y, Takeda S, Sekine K, Yoshihara Y, Kawakami T, Arioka K, Sato H, Uchiyama Y, Masushige S, Fukamizu A, Matsumoto T, Kato S: Mice lacking the vitamin D receptor exhibit impaired bone formation, uterine hypoplasia and growth retardation after weaning. Nat Genet 16:391 -396, 1997
42. Luetteke NC, Qiu TH, Fenton SE, Troyer KL, Riedel RF, Chang A, Lee DC: Targeted inactivation of the EGF and amphiregulin genes reveals distinct roles for EGF receptor ligands in mouse mammary gland development. Development 126:2739 -2750, 1999
Copyright © 2001 The Authors. Published by Wolters Kluwer Health, Inc. All rights reserved.