Sp1 Is a Critical Regulator of the Wilms' tumor-1 Gene*

(Received for publication, September 13, 1996, and in revised form, October 25, 1996)

Herbert T. Cohen , Steven A. Bossone , Guoming Zhu , Glenn A. McDonald and Vikas P. Sukhatme Dagger

From the Renal Division, Beth Israel Hospital and Harvard Medical School, Boston, Massachusetts 02215

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
Acknowledgments
REFERENCES


ABSTRACT

We performed deletion analysis of WT1-reporter constructs containing up to 24 kilobases of 5'-flanking and first intron WT1 sequence in stably transfected cultured cells as an unbiased approach to identify cis elements critical for WT1 transcription. Although not a tissue-specific element, a proximate 9-base pair CTC repeat accounted for ~80% of WT1 transcription in this assay. Enhancer activity of the element and mutated versions correlated completely with their ability to form a DNA-protein complex in gel shifts. Antibody supershift, oligonucleotide competition, and Southwestern studies indicated that the CTC-binding factor is the transcriptional activator Sp1. Sp1 binds the CTC repeat with an affinity, KD = 0.37 nM, at least as high as the consensus GC box. Similar CTC repeats are found in promoters of other growth-related genes. Because Sp1 is important for WT1 expression, we examined Sp1 immunohistochemistry in fetal and adult kidney. In a pattern that precedes that of WT1 message, Sp1 immunostaining was highest in uninduced mesenchyme, early tubules, developing podocytes, and mature glomeruli, but was minimal in mature proximal tubules. This work suggests abundant Sp1 may be a prerequisite for WT1 expression, and that Sp1 may have a wider role in nephrogenesis.


INTRODUCTION

The Wilms' tumor-1 (WT1) gene is essential for development of the genitourinary tract. Homozygous germline WT1 disruption in mice causes renal agenesis and gonadal failure (1). The knockout mice also have hypoplastic hearts and lungs, likely due to failed interaction with the mesothelium (1), which normally expresses WT1 (2). In humans, germline single-allele WT1 defects cause urogenital abnormalities and strongly predispose to Wilms' tumor. Male genital ambiguity and a nephropathy that progresses to renal failure are frequent features of the Denys-Drash syndrome (3-5), due to single-allele WT1 point mutations (6-8). Less profound genitourinary malformations, such as cryptorchidism and hypospadias, are associated with aniridia and Wilms' tumor (9-11), which with mental retardation comprise the WAGR syndrome, and result from one WT1 allele loss in a chromosome 11p13 deletion (12). Both syndromes are also characterized by persistent nodules of embryonic kidney tissue, called nephrogenic rests, from which malignant Wilms' tumors may arise (13), and their histopathology is suggestive of aberrant kidney development. The WT1 gene encodes a putative tumor suppressor (14), and biallelic intragenic WT1 defects have been found in a small percentage of Wilms' tumors (15). The WT1 protein is a zinc finger transcription factor (16-18) that in most contexts acts as a transcriptional repressor (Ref. 19 and below). Many candidate WT1 target genes have been identified, such as insulin-like growth factor 2 (igf2) (20), platelet-derived growth factor (pdgf) A-chain (21, 22), igf 1 receptor (23), epidermal growth factor receptor (egfr) (24), and Pax2 (25), among others. An attractive hypothesis for Wilms' tumor pathogenesis is that loss of the negative regulator WT1 results in overexpression of one or more growth-promoting target genes (reviewed in Ref. 26).

Little is known about regulation of the WT1 gene, but part appears to be transcriptional. The WIT1 gene is adjacent to WT1 and transcribed in the opposite direction from a shared ~2-kb1 intergenic region (27). Because WIT1 and WT1 are often coexpressed (27), shared cis elements may be important in their regulation. Furthermore, nuclear run-on studies from our laboratory in WT1-expressing and non-expressing cell lines support that a major component of the gene's regulation is transcriptional.2

WT1 is tightly regulated. WT1 message is tissue-restricted and expressed in a striking pattern in nephrogenesis (29). Little WT1 message is detected in undifferentiated metanephric mesenchyme (30-32), but levels increase prominently in the structures formed as the mesenchyme condenses, the renal vesicle, comma-shaped, and S-shaped body, where expression becomes localized to the developing glomerular podocytes (29). Throughout nephrogenesis, WT1 protein is expressed coordinately with WT1 message, supporting that WT1 regulation is not translational (32, 33). In addition, the WAGR genitourinary defects are most likely due to reduced WT1 expression from haplotype insufficiency. The above observations suggest that WT1 message levels are important in development and that transcriptional control of the WT1 gene is likely.

WT1 is transcribed from a 5'-flanking region that is ~70% GC-rich and lacks a consensus TATA box (34, 35). Multiple WT1 transcription start sites (TSS) have been identified over a ~300-bp promoter region (35-38). These data indicate that the human WT1 major TSS is in the region 393-413 bp upstream of the initial ATG (35, 37) (Fig. 1B, dotted overline), and this TSS is conserved in the mouse gene (36). While no consensus initiator element resides in this region, a candidate TATA box (CTTATTTGA) (Fig. 1, A and B) that resembles the functional SV40 early promoter TATA box (CTTATTTAT) (39) is well positioned upstream of the major TSS and may direct initiation. Other regulatory regions of the WT1 promoter remain unexplored.


Fig. 1. Sequence of the WT1 promoter region. A, computer (MacVector®, Kodak) and hand alignment of murine (36) and human WT1 (GenBankTM accession no. U77682[GenBank]) promoter sequences. A consensus is shown above the compared sequences, with discrepancies marked by black boxes containing IUPAC code: K = T/G, M = A/C, R = A/G, S = C/G, W = A/T, Y = C/T. Because of skips (·), numbering is approximate relative to the major TSS (arrow) designated +1. B, sequence of the human WT1 major transcription initiation region and immediate 5'-flank. The dotted line and arrow overlie the identified major start sites (35-37). The sequence is numbered with respect to the 3'-end of the TSS region (+1). Mutations (to adenine) generated in oligonucleotides and cloned into promoter-reporter constructs are designated by the labeled solid lines, A-E, B1-B3, and C4-C6. The nucleotides important to Sp1 binding and reporter activation are emphasized by bold-face type and increased size.
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We report here the identification and characterization of a critical WT1 promoter enhancer element and the corresponding transcriptional activator.


EXPERIMENTAL PROCEDURES

Isolation of a Human WT1 Genomic Clone and Construction of WT1-Reporter Plasmids

A human lung fibroblast (WI38) genomic library in bacteriophage lambda FIX (Stratagene) was screened using two probes to obtain genomic WT1 sequence. The first probe was 5' human WT1 cDNA sequence (gift of K. Call, Harvard School of Public Health), a 1.8-kb EcoRI-EcoRI fragment, and the second was murine WT1 5'-flanking region (bp -593 to +200, relative to the major TSS; Ref. 36) generated by polymerase chain reaction. One clone screening positive with both probes contained 9.4 kb of upstream and 2.4 kb of downstream sequence (designated here -9.4/+2.4 kb) with respect to the WT1 major TSS. The clone was sequenced over ~1.5 kb of the proximate promoter using a deazaG and deazaA chain-termination sequencing kit (Pharmacia Biotech Inc.). The 11.8-kb WT1 genomic fragment was cloned into pBluescript II SK+ (Stratagene), which was converted into a reporter construct by placing a blunted XbaI-BamHI fragment from pCAT-Basic (Promega), containing the chloramphenicol acetyltransferase (CAT) gene and SV40 t intron and poly(A) addition site, into a unique BbrPI site in the WT1 clone 5'-untranslated region at position +206. A still larger -22/+2.4 kb WT1-reporter construct was made by placing a -2.5/+2.4 kb BamHI fragment from the -9.4/+2.4 kb construct into cosmid pWE15 (Stratagene) and inserting upstream a 19.4-kb WT1 BamHI fragment (-22/-2.5 kb) from WT1 cosmid L156 (16) (gift of D. Haber, Massachusetts General Hospital). Serial deletions were made from the -9.4/+2.4 kb construct using restriction sites SnaBI (-4.3 kb) and BamHI (-2.5 kb). In addition, a WT1 PstI-PstI fragment (-2.4 kb/+191 bp) was cloned into pCAT-Basic to yield a reporter construct with the CAT gene in a location similar to the larger constructs. In the latter case, fine deletions were made using polylinker and WT1 restriction sites, yielding 5' deletions at the following locations: SacI (-1.1 kb), HindIII (-453 bp), BpmI (-319 bp), AvaII (-168 bp), BssHII (-122 bp), EagI (-81 bp), and NaeI (-48 bp). Series of -81/+191 bp linker scanning reporter constructs were made by inserting EagI(-86 bp)-NgoMI(-50 bp) sticky-ended double-stranded oligonucleotides (see below) (Oligos Etc.) into a HindIII- and NgoMI-cut -453/+191 bp vector, blunting with T4 polymerase, and ligating. The -453/+191 bp linker scanning constructs were made by placing the EagI-NgoMI oligonucleotides into an EagI- and NgoMI-cut -453/+191 bp vector. To make the minimal promoter linker scanning constructs, EagI-XbaI sticky-ended double-stranded oligonucleotides (see below) were cloned into the XbaI site in E1b TATA-CAT vector (40) such that the final constructs contained from -81 to -39 of WT1 sequence. The -81/+191 bp, -48/+191 bp, and all linker scanning constructs were sequenced to confirm their authenticity.

Transfections and CAT Assays

293, HeLa, HeLa S3, and Drosophila Schneider S2 cells were obtained from ATCC. JMN cells (41) were the gift of J. Rheinwald, Brigham and Women's Hospital. Human cells were maintained in Dulbecco's modified Eagle's medium with 10% fetal bovine serum (HyClone) and Schneider cells in Schneider medium with 10% heat-inactivated serum (Sigma). Cells were transfected by calcium-phosphate precipitation with a total of 20 µg of plasmid DNA/100-mm dish. Transient transfections included 20-50% reporter plasmid, 1-5% transfection control plasmid (CMV-driven beta -galactosidase), and the remainder pBluescript II SK+, while stable transfections comprised 95% reporter construct and 5% neo resistance gene vector (pSV2-neo). Duplicate tissue culture dishes for each construct were transfected in each experiment. Transiently transfected cells were harvested at 48 h, and stably transfected cells were passaged from 100- to 150-mm dishes into medium containing 100-600 µg/ml G418 (Life Technologies, Inc.) at 48 or 72 h (42). Stably transfected cells were maintained in culture under selection for 2-4 weeks, until colonies were obvious and non-resistant cells had died. Stable transfections were considered valid only if >100 colonies of roughly equal size resulted, to negate integration position and copy number effects. CAT assays were then performed on the entire pool of stable colonies and normalized for protein concentration by Bradford assay (Bio-Rad). Percent chloramphenicol acetylation was determined by thin-layer chromatography and scintillation counting of autoradiographed spots.

Nuclear Extract Preparation, Partial Protein Purification, and Electrophoretic Mobility Shift Assays (EMSAs)

Nuclear extracts were prepared from cultured cells (43) and from tissue, with modifications (44). For further purification, HeLa S3 nuclear extracts were prepared according to modifications (45) of another standard protocol (46). HeLa S3 crude nuclear extract, ~150 mg of protein from 10 liters of culture (Cell Culture Center), was partially purified over a 2-ml wheat germ agglutinin-agarose affinity column (Vector Laboratories) (45).

EMSA reactions (25 µl) contained 20 mM HEPES, pH 8.4, 100 mM KCl, 20% glycerol, 0.1 mM EDTA, 0.25 mM ZnSO4, 0.05% Nonidet P-40. Thirty ng to 2 µg of protein and an equal amount of poly(dA·dT) (dA·dT) were added at room temperature 10 min prior to addition of 0.1-7 ng of radiolabeled oligonucleotide probe, after which room temperature incubation continued for 18 min. One µg of acetylated bovine serum albumin was used as carrier with purified protein. Four or 5% acrylamide gels were run in 1 × TAE buffer (40 mM Tris acetate, pH 8.5, 2 mM EDTA) at room temp at 160 V. For quantitation of the EMSA bands and KD determination, a constant amount of extract was incubated with an increasing amount of probe until saturation of factor binding occurred (42). The fraction of probe in a protein-DNA complex was determined by Cerenkov counting of excised, autoradiographed bound and free EMSA bands, and the amount of protein by assuming a protein:DNA molar ratio of 1. The 31-bp WT1 wild-type enhancer or 26-bp Sp1 consensus oligonucleotides were used to determine KD values, which were calculated as -slope-1 in Rosenthal plots (47). For EMSA supershift studies, 1-2 µg of monoclonal or polyclonal Sp1 antibody (Santa Cruz Biotechnology) was added for 40 min at room temperature following standard binding reaction above.

Synthesized oligonucleotides (Oligos Etc.) are shown below. The two series of 45-bp WT1 6-bp and 2-bp linker scanning oligonucleotides were designed with EagI-XbaI ends. The EagI site is found in human WT1, but the XbaI site was created by changing the first 2 nucleotides of the lower strands. The full-length 45-bp oligonucleotides were used for the E1b TATA constructs only. For the WT1 context constructs and for EMSAs, the 45-bp oligonucleotides were inserted into pBluescript II SK+, excised with EagI and NgoMI, and gel-purified to yield 36-bp WT1 oligonucleotides (see Fig. 1B). The 31-bp WT1 oligonucleotides contain a single base change so they could concatamerize. The 26-bp Sp1 consensus oligonucleotide was designed to contain the same 18-bp Sp1 consensus sequence used by Letovsky and Dynan (48) with EagI-NgoMI ends. Another Sp1 consensus oligonucleotide, also containing the 18-bp sequence, was obtained from Promega. Synthesized oligonucleotides are shown with mutations underlined and regions excised in parentheses: WT1 wild-type enhancer (45 bp), upper strand (US) 5'-GGCCGAGCCTCCTGGCTCCTCCTCTTCCCCGCGCCG(CCGGCCCCT)-3' and lower strand (LS) 5'-(<UNL>CT</UNL>AGAGGGG)CCGGCGGCGCGGGGAAGAGGAGGAGCCAGGAGGCTC-3'; WT1 A enhancer (45 bp), (US) 5'-GGCCG<UNL>AAAAAA</UNL>CTGGCTCCTCCTCTTCCCCGCGCCG(CCGGCCCCT)-3' and (LS) 5'-(<UNL>CT</UNL>AGAGGGG)CCGGCGGCGCGGGGAAGAGGAGGAGCCAG<UNL>TTTTTT</UNL>C-3'; WT1 B enhancer (45 bp), (US) 5'-GGCCGAGCCTC<UNL>AAAAAA</UNL>CCTCCTCTTCCCCGCGCCG(CCGGCCCCT)-3' and (LS) 5'(<UNL>CT</UNL>AGAGGGG)CCGGCGGCGCGGGGAAGAGGAGG<UNL>TTTTTT</UNL>GAGGCTC-3'; WT1 C enhancer (45 bp), (US) 5'-GGCCGAGCCTCCTGGCT<UNL>AAAAAA</UNL>CTTCCCCGCGCCG(CCGGCCCCT)-3' and (LS) 5'-(<UNL>CT</UNL>AGAGGGG)CCGGCGGCGCGGGGAAG<UNL>TTTTTT</UNL>AGCCAGGAGGCTC-3'; WT1 D enhancer (45 bp), (US) 5'-GGCCGAGCCTCCTGGCTCCTCCT<UNL>AAAAAA</UNL>CGCGCCG(CCGGCCCCT)-3' and (LS) 5'-(<UNL>CT</UNL>AGAGGGG)CCGGCGGCGCG<UNL>TTTTTT</UNL>AGGAGGAGCCAGGAGGCTC-3'; WT1 E enhancer (45 bp), (US) 5'-GGCCGAGCCTCCTGGCTCCTCCTCTTCCC<UNL>AAAAAA</UNL>G(CCGGCCCCT)-3' and (LS) 5'-(<UNL>CT</UNL>AGAGGGG)CCGGC<UNL>TTTTTT</UNL>GGGAAGAGGAGGAGCCAGGAGGCTC-3'; WT1 B1 enhancer (45 bp), (US) 5'-GGCCGAGCCTC<UNL>AA</UNL>GGCTCCTCCTCTTCCCCGCGCCG(CCGGCCCCT)-3' and (LS) 5'-(<UNL>CT</UNL>AGAGGGG)CCGGCGGCGCGGGGAAGAGGAGGAGCC<UNL>TT</UNL>GAGGCTC-3'; WT1 B2 enhancer (45 bp), (US) 5'-GGCCGAGCCTCCT<UNL>AA</UNL>CTCCTCCTCTTCCCCGCGCCG(CCGGCCCCT)-3' and (LS) 5'-(<UNL>CT</UNL>AGAGGGG)CCGGCGGCGCGGGGAAGAGGAGGAG<UNL>TT</UNL>AGGAGGCTC-3'; WT1 B3 enhancer (45 bp), (US) 5'-GGCCGAGCCTCCTGG<UNL>AA</UNL>CCTCCTCTTCCCCGCGCCG(CCGGCCCCT)-3' and (LS) 5'-(<UNL>CT</UNL>AGAGGGG)CCGGCGGCGCGGGGAAGAGGAGG<UNL>TT</UNL>CCAGGAGGCTC-3'; WT1 C4 enhancer (45 bp), (US) 5'-GGCCGAGCCTCCTGGCT<UNL>AA</UNL>TCCTCTTCCCCGCGCCG(CCGGCCCCT)-3' and (LS) 5'-(<UNL>CT</UNL>AGAGGGG)CCGGCGGCGCGGGGAAGAGGA<UNL>TT</UNL>AGCCAGGAGGCTC-3'; WT1 C5 enhancer (45 bp), (US) 5'-GGCCGAGCCTCCTGGCTCC<UNL>AA</UNL>CTCTTCCCCGCGCCG(CCGGCCCCT)-3' and (LS) 5'-(<UNL>CT</UNL>AGAGGGG)CCGGCGGCGCGGGGAAGAG<UNL>TT</UNL>GGAGCCAGGAGGCTC-3'; WT1 C6 enhancer (45 bp), (US) 5'-GGCCGAGCCTCCTGGCTCCTC<UNL>AA</UNL>CTTCCCCGCGCCG(CCGGCCCCT)-3' and (LS) 5'-(<UNL>CT</UNL>AGAGGGG)CCGGCGGCGCGGGGAAG<UNL>TT</UNL>GAGGAGCCAGGAGGCTC-3'; WT1 wild-type enhancer (31 bp), (US) 5'-CCGAGCCTCCTGGCTCCTCCTCTTCCCCGCG-3' and (LS) 5'-<UNL>T</UNL>CGGCGCGGGGAAGAGGAGGAGCCAGGAGGC-3'; WT1 C enhancer (31 bp), (US) 5'-CCGAGCCTCCTGGCT<UNL>AAAAAA</UNL>CTTCCCCGCG-3' and (LS) 5'-<UNL>T</UNL>CGGCGCGGGGAAG<UNL>TTTTTT</UNL>AGCCAGGAGGC-3'. Sp1 consensus oligonucleotide (26 bp) was (US) 5'-GGCCGATTCGATCGGGGCGGGGCGAGC-3' and (LS) 5'-CCGGCTCGCCCCGCCCCGATCGAATC (48). egfr sequences were (US) 5'-CGGCCGCCTGGTCCCTCCTCCTCCCGCCCTGCCTGCCGGC-3' and (LS) 5'-GCCGGCAGGCAGGGCGGGAGGAGGAGGGACCAGGCGGCCG-3' (49), and vav was (US) 5'-CGGCCGCCGCCCCATGGCTCCTCCTCCTCCACCCCCTCTAGA-3' and (LS) 5'-TCTAGAGGGGGTGGAGGAGGAGGAGCCATGGGGCGGCGGCCG-3' (50).

Southwestern Analysis of Nuclear Extract Protein

HeLa S3 cell crude nuclear extract (120 µg of protein/lane) was electrophoresed on 6.5% SDS-containing polyacrylamide gels (Bio-Rad). Size-separated proteins were transferred semidry to polyvinylidene difluoride membrane (Millipore). Membrane-bound proteins were denatured in 6 M guanidine in hybridization buffer (20 mM HEPES, pH 7.9, 100 mM KCl, 0.1 mM EDTA, 1 mM dithiothreitol) and stepwise renatured (51). Membrane strips were blocked with 5% nonfat dry milk in hybridization buffer, incubated overnight with monomeric radiolabeled 31-bp WT1 enhancer or 26-bp Sp1 consensus oligonucleotides (2.7 × 106 cpm/ml) in hybridization buffer with 0.25% milk at 4 °C for 15 h, washed six times with hybridization buffer with 0.25% milk at 4 °C, and autoradiographed.

Immunohistochemistry Analysis

Kidneys from adult or fetal (gestation day 18) Wistar rats (Charles River Laboratories) were snap-frozen in OCT compound (Miles, Inc.) by immersion in liquid nitrogen. Frozen sections (8 µm thick) were fixed in -20 °C acetone for 30 min. Endogenous peroxidases were quenched with H2O2, and biotin binding sites blocked according to kit manufacturer (Vector Laboratories). After blocking with normal goat serum, the sections were incubated at 4 °C overnight with rabbit anti-human Sp1 serum AHP312 (Serotec USA) diluted 1:2000. Binding of Sp1 antiserum to kidney tissue was detected with an ABC elite kit and peroxidase substrate kit (Vector Laboratories) using the protocols recommended by the manufacturer. Some Sp1-stained sections were subsequently treated with periodic acid for 10 min and stained with Schiff reagent to visualize basement membranes.


RESULTS

Identification of an Important Cis Element in the Human WT1 Proximate Promoter

To identify key WT1 enhancer sequences, we made -22/+2.4 kb, -9.4/+2.4 kb, and -2.5/+0.2 kb human WT1-CAT reporter constructs and deletions (see "Experimental Procedures") and tested their relative activities in pooled stable transfection assays. We have designated as the WT1 transcription start site (+1) the first G of the GGGG sequence located 393 bp upstream of the WT1 ATG, as determined by primer extension start site mapping in the mouse gene (+1, Figs. 1, A and B) (36), and confirmed for human WT1 as in the region -20 to +1 bp shown in Fig. 1B (35, 37). The CAT gene was placed in the WT1 5'-untranslated region, at position +206 in the constructs derived from the -9.4/+2.4 kb construct and at +191 bp in the -2.5 kb and smaller constructs. Transfections were carried out in human lines, WT1-expressing 293 (embryonic kidney) and JMN (mesothelioma) cells, and non-WT1 expressing HeLa (cervical carcinoma) cells. Relative CAT activities of the reporter constructs were normalized to that of the -48/+191 bp construct.

The intrinsic relative CAT activities of the large WT1-reporter constructs and derived broad deletion constructs were comparable in the three cell lines tested (Fig. 2). In addition, their absolute levels of CAT activity were similar as well (data not shown). No large decrements in relative CAT activity (>3-fold) were observed until the WT1 region from -453 to -48 bp was deleted, whereupon CAT activity fell up to 10.6-fold, with the greatest effects observed in the WT1-expressing 293 and JMN cells (Fig. 2). Although the -22/+2.4 kb construct was less active than the -9.4/+2.4 kb construct, little transcriptional activity was lost with subsequent -4.3/2.4 kb, -2.4/0.2 kb, -1.1/0.2 kb, and -453/+191 bp deletions. In fact, the latter proximate promoter construct retained from 30 to 79% activity of the most active construct (-9.4/+2.4 kb) and was more active than the the largest WT1-reporter construct containing a full 24.4 kb of WT1 locus sequence (Fig. 2). These findings support that over the region tested, the proximate promoter contributes a large fraction of transcriptional activity to the WT1 gene. An important non-tissue-specific WT1 enhancer therefore resides in the proximate promoter, between positions -453 and -48.


Fig. 2. Intrinsic activities of large WT1-reporter constructs and deletions stably transfected into pools of WT1-expressing 293 and JMN cells, and in non-expressing HeLa cells. The constructs are schematized (left) with WT1 sequence as solid black lines and the WT1 enhancer region as a shaded box. Fold activation for each cell line, but not between lines, is normalized to the -48/+191 bp construct. Each reported CAT activity is the mean of one to five experiments. For each experiment, duplicate tissue culture dishes, each containing >100 similar-size G418-resistant colonies, were assayed to minimize copy-number and integration-position effects. Deletion of the region from -453 to -48 bp results in a large decrement in reporter activity in the three cell lines.
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To obtain functional clues about the WT1 proximate promoter, we compared the human (GenBankTM accession no. U77682[GenBank]) and mouse sequences (36) (shown in part in Fig. 1A). This computer (MacVector®, Kodak Corp.) and hand aligned sequence comparison reveals high homology (88.5%) over the -106 to +242 bp promoter region, which falls off to 68% from -107 to -453 bp (data not shown), with islands of conserved sequence. These observations suggest that the highly conserved WT1 immediate 5'-flank may have particular regulatory importance.

We then tested a series of fine deletion constructs spanning from -453/+191 bp to -48/+191 bp to localize the enhancer within this ~400-bp region. Nearly all enhancer activity, 52-220% of the -453/+191 bp construct and 41-66% of the -9.4/+2.4 kb construct, resides in a 33-bp region, from -81 to -48 with respect to the major TSS (Fig. 3), supporting that the 33-bp enhancer is a major contributor to WT1 transcription over the 24.4-kb region examined. This enhancer functioned in all cell lines tested and therefore does not appear to directly contribute to tissue-specific gene expression. It conferred 5.5-12.5-fold activation in the stable transfection studies shown in Fig. 3. In transient transfection studies in the same lines, activation by the enhancer was 2-3-fold less (data not shown), suggesting that the enhancer plays a role in disrupting inhibitory effects of chromatin. The enhancer was active as well in mouse fibroblasts (NIH 3T3 cells) and was equipotent in the reverse orientation (data not shown). The position of the enhancer, centered 45-66 bp upstream of the major transcription start site, indicates that the enhancer likely serves a regulatory function and is not a core promoter element.


Fig. 3. Fine 5' deletions of the -453/+191 bp WT1-CAT construct demonstrate that most enhancer activity resides in a 33-bp element, from -81 to -48 bp (shaded box). Intrinsic CAT activities of the schematized constructs (left) were measured in pools of stably transfected cultured cells and normalized to the the -48/+191 bp construct. One to three experiments in each cell line are represented.
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"Linker Scanning" Analysis of the 33-bp Region

To determine which part of the 33-bp WT1 enhancer was responsible for its activity, a series of -81/+191 bp constructs were made with 6-bp tandem mutations introduced to "scan" the enhancer. Five double-stranded oligonucleotides were placed 5' of the enhancerless -48/+191 bp WT1-reporter constructs to generate a series of -81/+191 bp constructs with conversion of the following nucleotides to 6 adenines, with mutations designated as A (-80 to -75), B (-74 to -69), C (-68 to -63), D (-62 to -57), and E (-56 to -51) (see Fig. 1B). The same pattern of results was seen in each of the cell lines and was observed whether cells were stably or transiently transfected. Mutations in the 6-bp B or C regions completely eliminated enhancer activity, reducing reporter activity to the level of the enhancerless -48/+191 bp construct (Fig. 4). The D mutation allowed partial activation, while the A and E region mutations did not greatly affect enhancer activity (Fig. 4). These studies indicate that enhancer activity depends on the integrity of a small subelement.


Fig. 4. 6-bp tandem "linker scanning" mutations were introduced into the WT1 enhancer in the -81/+191 bp context. The shaded boxes represent the WT1 enhancer, while the hatched regions within represent each mutation's position (see Fig. 1B for designated mutations). Intrinsic CAT activities of the constructs were measured in pools of stably transfected cultured cells and were normalized to the -48/+191 bp construct. In all three lines, the B and C region mutations blocked activation and the D mutation allowed partial activation by the enhancer. Each reported CAT activity is the mean of one to three experiments.
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To determine the contribution of the enhancer to overall transcription of the promoter region, we introduced the B, C, and D linker scanning mutations in context into the active -453/+191 bp proximate promoter construct and tested their effects on reporter activity in pooled stable transfection assays. The -453/+191 bp construct contains all identified forward-direction WT1 transcription start sites (35-37). In both 293 and HeLa cells, mutations in the B or C regions in the context of the active proximate promoter fragment brought reporter activity down about 5-fold (Fig. 5), indicating that the enhancer is responsible for the majority, perhaps 80%, of the transcriptional activity of the promoter. It also suggests that in these larger mutated WT1 promoter constructs, additional GC-rich sequence, to which many transcriptional activators might bind, and a full complement of transcription start sites cannot compensate for loss of the enhancer. As in the -81/+191 bp linker scanning constructs, the -453/+191 bp D mutation still allowed partial activation (Fig. 5).


Fig. 5. The WT1 enhancer tested in a larger WT1 context. The B, C, and D mutations were introduced into the -453/+191 bp WT1-reporter construct (shown schematically). Intrinsic CAT activities of the constructs stably transfected into pools of 293 and HeLa cells are reported. Data represent results of one experiment and are normalized, in each line, to the -48/+191 bp construct. The B and C mutations reduced -453/+191 bp reporter activity at least 5-fold, while the D mutation was partially active.
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To further confirm the importance of the enhancer, single copies of the wild-type and B, C, and D mutated enhancers were tested in front of a heterologous minimal promoter, at a distance from the initiation site comparable to the WT1 context. The base vector was the 13-bp E1b TATA region linked to the CAT gene (40). The same pattern of results was observed in transiently transfected 293 and HeLa cells. As shown in Fig. 6, the enhancerless construct had no CAT activity above background, but the addition of the wild-type enhancer brought reporter activity up remarkably. Fold activation is infinite (Fig. 6), and the level of CAT activity is within 2-fold of the -453/+191 bp construct (data not shown). The D mutated enhancer had about half the activity of the wild-type. These observations indicate that this small WT1 enhancer is itself sufficient to confer a high level of transcriptional activation and, in the WT1 context, likely does so through the major WT1 TSS.


Fig. 6. Single copies of the wild-type and mutated WT1 enhancers were tested in front of a minimal promoter, the E1b TATA box (40), in transiently transfected HeLa cells. A representative experiment with duplicate raw CAT data is shown (right). The wild-type enhancer (and D mutation version) conferred considerable reporter activity, while the B and C mutated enhancers, like the enhancerless base vector, gave no CAT activity above background.
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In sum, we have identified a powerful enhancer in the WT1 promoter that accounts for the majority of its transcriptional activity.

A Prominent Protein-DNA Complex Forms Only in Association with the Functional Enhancer Sequences

The simplest model to explain the linker scanning transfection data is that a transcriptional activator is capable of binding the wild-type and functional mutated enhancers, but not the non-functional mutated enhancers. To test this hypothesis, EMSAs were performed using the 36-bp linker scanning oligonucleotides as probes. In all EMSAs comparing the wild-type and mutated enhancer oligonucleotide probes, the probes were labeled to comparable specific activities and incubated with equivalent amounts of protein. Nuclear extracts were prepared from WT1-expressing human and pig fetal kidney and the above cell lines. As predicted by the model, a prominent protein-DNA complex (upper band) forms in vitro with the wild-type enhancer sequence, but not with the B or C mutated enhancers, which lacked enhancer activity (Fig. 7). In contrast, the A, D, and E mutated enhancers that retained enhancer activity were able to form the same upper band complex as the wild-type (Fig. 8). Interestingly, the D mutated enhancer, which had only partial activating potential (Figs. 4, 5, 6), exhibited reduced affinity for the upper band factor (Fig. 8). An additional double-stranded oligonucleotide, designated F, with bases -50 to -45 changed to adenines, also readily formed the upper band complex (data not shown). These in vitro findings correlate well with the cell culture transfection results and suggest that activity of the WT1 enhancer depends on its ability to form this protein-DNA complex, and that the complex likely contains the relevant transcriptional activator.


Fig. 7. EMSA comparing nuclear extract protein binding to the wild-type WT1 enhancer (-86 to -47 bp) and nonfunctional B and C mutated enhancers. See Fig. 1B for sites of designated mutations. Each probe (labeled above) is incubated with (from left to right) no nuclear extract protein, nuclear extract from pig fetal kidney (PFK), 293 cells, and HeLa cells. A prominent DNA-protein complex (uppermost band) forms with the wild-type enhancer, but not the nonfunctional B and C mutated enhancers.
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Fig. 8. EMSA comparing nuclear extract protein binding to the wild-type WT1 enhancer and the functional A, D, and E mutated enhancers. See Fig. 1B for sites of designated mutations. Each probe (labeled above) is incubated with (from left) no extract, nuclear extract from 293 cells, and from HeLa cells. All these enhancers allow formation of the upper band complex, but the D enhancer, with reduced enhancer activity, exhibits diminished binding.
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To further refine the DNA contacts required for DNA binding of this factor, six additional double-stranded linker scanning oligonucleotides were made containing 2-bp tandem mutations that span the 12 bp of the B and C regions. They are designated B1-3 and C4-6 and have the following 2 bp changed from the wild-type sequence to 2 adenines: B1 (-74, -73), B2 (-72, -71), B3 (-70, -69), C4 (-68, -67), C5 (-66, -65), and C6 (-64, -63) (see Fig. 1B). In EMSA assays, formation of the prominent upper band complex is nearly lost with the B3 mutation, and completely lost with the three C mutations (Fig. 9). Therefore, the wild-type sequence of nucleotides that is essential for protein-DNA complex formation is CTCCTCCT. It is also likely that the first C of the D region contributes to binding as well. The (CTC)3 sequence and position are completely conserved in the mouse WT1 gene (Fig. 1A).


Fig. 9. EMSA comparing 293 cell nuclear extract binding to the A and B mutated enhancers (controls) and 2-bp tandem "linker scanning" oligonucleotides that scan the region from -74 to -63 bp. Each 2-bp mutated oligonucleotide was run without (left of each pair of lanes) and with 293 cell protein. See Fig. 1B for sites of designated mutations. Formation of the upper band protein-DNA complex is greatly decreased with the B3 mutation and essentially eliminated with the C4-C6 mutations.
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Additionally, as a means of further correlating protein-DNA complex formation with enhancer activity, another set of -81/+191 bp WT1-CAT reporter constructs were made using the B1-3 and C4 oligonucleotides and tested in transiently transfected HeLa cells. These 2-bp mutated enhancers conferred transcriptional activation that again correlated with their respective abilities to form an in vitro protein-DNA complex. The B1 and B2 mutated enhancers conferred 8.9- and 7.1-fold activation, respectively, the B3 enhancer (that exhibited much reduced binding) 2-fold activation, while the C4 mutation that showed no binding had no enhancer activity (Fig. 10). These observations underscore how much the conditions of the in vitro binding reactions reflect the activity of the enhancer in vivo and, consequently, provide additional support that the protein-DNA complex formed in vitro may contain the relevant transcriptional activator.


Fig. 10. Activation conferred by the 2-bp mutated WT1 enhancers correlates with their abilities to form the prominent upper band complex, shown in Fig. 9. Data were obtained by transient transfection of HeLa cells (1 experiment).
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The DNA-binding Protein Has Characteristics of Sp1

Formation of the protein-DNA complex requires zinc. When EMSA reaction buffer (containing 0.1 mM EDTA) lacked zinc, no upper band complex formed (data not shown). Addition of excess zinc in the presence of the EDTA or the zinc chelator orthophenanthroline allowed complex formation (data not shown). Sp1, which contains three C2H2 zinc fingers, requires zinc for DNA binding.

Protein-DNA complex formation could be prevented by preincubation of extracts with the lectin wheat germ agglutinin (WGA). This inhibition could be overcome by including in the EMSA reaction N-acetylglucosamine, the ligand recognized by WGA (data not shown). This suggested that the reaction was specific, and that the DNA-binding protein was likely O-glycosylated, as has been described for several transcription factors, most notably Sp1 (52) and HNF1 (53). The protein could also be partially purified using a WGA-agarose affinity resin, as has been shown for both Sp1 (45, 52) and HNF1 (53).

Binding of the factor to DNA was optimized in EMSA studies (data not shown). A KCl concentration of 100 mM was more than 2-fold better than 50 or 150 mM, above or below which binding fell off considerably. Titration of pH showed that pH 8.4 (or even 9.0) was better than pH 7.9, 7.4, 7.1, or 6.5, below which binding was greatly reduced. EDTA concentration was kept at 0.1 mM as zinc concentration was optimized. Binding improved up to 0.25 mM ZnSO4, but was completely abolished with 0.5 mM or greater ZnSO4. Sp1's exquisite sensitivity to zinc concentration for DNA binding is known (54, 55). Addition of magnesium interfered with protein-DNA complex formation. With the exception of magnesium, our optimized conditions are similar to those reported for Sp1 protein-DNA interactions with the GC box: 100 mM KCl, 3-12.5 mM MgCl2, 0-1 mM EDTA, 1 mM dithiothreitol, 0-0.1% Nonidet P-40, HEPES or Tris, pH 7.5 or 7.9 (48, 56-58).

The factor was found in all cell lines tested and in differing amounts, as assessed by EMSA (data not shown). The amount of the factor present in nuclear extracts correlated with the activity of the enhancer in that cell line. The amount of the protein was determined to be 1/5,000 of total nuclear extract protein in HeLa S3 cells, which is comparable to reported Sp1 content (45). In HeLa S3 cells, we found about one-third of the binding activity in the cytoplasmic S100 fraction, but at one-fifth the concentration as in nuclear extract.

The WT1 Enhancer Binding Factor Is Sp1

EMSA studies with Sp1 antibody and competition with a consensus Sp1 oligonucleotide strongly support that the DNA-binding protein is Sp1. An affinity-purified mouse monoclonal antibody directed against Sp1 amino acids 520-538 (Santa Cruz Biotechnology) or an affinity-purified rabbit polyclonal antibody made against Sp1 amino acids 436-454 (Santa Cruz Biotechnology) were added to EMSA reactions containing the wild-type WT1 enhancer sequence or an Sp1 consensus oligonucleotide. These antibodies are directed against sequences that are amino-terminal to the DNA-binding zinc finger domain and would therefore be expected to supershift, not prevent formation of, an Sp1-containing protein-DNA complex. Both antibodies are Sp1-specific, as they do not cross-react with Sp1 family members Sp2, -3, or -4 (Santa Cruz Biotechnology). As shown in Fig. 11, both antibodies were capable of supershifting the prominent upper band complex formed with the WT1 or consensus GC box oligonucleotides, strongly suggesting that the DNA-binding material contains Sp1. Furthermore, the similar appearance of the supershifts from the two different oligonucleotides suggests they are composed of identical material. With the polyclonal antibody, nearly all material in the protein-DNA complex is supershifted, indicating that the major protein component is Sp1, as might be expected for a gel shift with the GC box.


Fig. 11. EMSA supershift study with Sp1 antibodies using the WT1 wild-type enhancer oligonucleotide left eight lanes) and the Sp1 consensus GC box (right seven lanes) as probes. Standard binding reactions (see "Experimental Procedures") were followed by addition of bovine serum albumin (1-2 µg) (control), affinity-purified Sp1 monoclonal antibody (mAb), or affinity-purified Sp1 antiserum (pAb) for 40 min at room temperature. Protein extracts included crude HeLa S3 nuclear extract (lanes 1-4 and 9-12), WGA partially purified Sp1 (lanes 5-8), and affinity-purified Sp1 overexpressed in HeLa S3 cells (Sp1) (Promega) (lanes 13-15). Both the monoclonal antibody and antiserum are able to supershift the upper band complex.
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Oligonucleotide competition studies in EMSAs revealed that the WT1 oligonucleotide could effectively compete for Sp1 binding with a consensus Sp1 site. The WT1 wild-type enhancer oligonucleotide was labeled to the same specific activity as the Sp1 consensus oligonucleotide, and the molar excess of unlabeled competitor is shown (Fig. 12). A 10-fold excess of either unlabeled oligonucleotide (lanes 3 and 8) sustantially reduced binding of Sp1-immunoreactive material to the other binding site (Fig. 12), indicating each site binds Sp1 with comparable affinity. In fact, the WT1 enhancer appears to be the better competitor and hence the higher affinity site (see below) under these assay conditions. The opposite strand of the CTCCTCCTC WT1 sequence (consensus in uppercase: GaGGaGGAG) is consistent in 7 of 9 bases with the Sp1 consensus sequence determined by Kadonaga et al. (T/G, GGGCGG, A/G, G/A, C/T) (59) and later confirmed (60). However, it is still surprising that the 2 nucleotide differences are in the 6-base Sp1 core, GGCGGG, and that the WT1 site was not detected in the DNA target assay (60). Nevertheless, this work indicates that the WT1 (CTC)3 enhancer is a high affinity Sp1 binding site (see below).


Fig. 12. EMSA competition study examining the effect of excess unlabeled Sp1 consensus site oligonucleotide on Sp1 binding to the WT1 enhancer oligonucleotide, and that of excess unlabeled WT1 enhancer oligonucleotide on binding to the Sp1 consensus GC box. Probes were labeled to comparable specific activities. Molar excess of unlabeled competitor oligonucleotide is shown.
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To determine the molecular weight of the WT1 enhancer-binding factor and to confirm it as Sp1, we performed Southwestern analysis (51, 61) of HeLa cell crude nuclear extract using the 31-bp WT1 wild-type, C mutated, and 26-bp Sp1 consensus oligonucleotides. We had also noted a subtle difference in electrophoretic mobility by EMSA of the Sp1-DNA complex using the WT1 versus the Sp1 consensus oligonucleotide and considered that the post-translational modification of the WT1 enhancer-binding material might be different from that which bound the Sp1 consensus site. We therefore attempted to size separate well the p105 and p95 forms of Sp1 using a 6.5% denaturing acrylamide gel before hybridizing with the radiolabeled oligonucleotides. As shown in Fig. 13, the p105 form of Sp1 bound the WT1 wild-type and Sp1 consensus oligonucleotides, while, as expected from the EMSA studies, binding to the C mutated sequence was considerably reduced. The reduced binding to the C mutated enhancer also supports the specificity of the Sp1 protein-WT1 enhancer interaction. In addition, that the assay worked after size separation of proteins and shows a strong interaction between Sp1 and the WT1 enhancer indicates that no cofactor is required for their interaction. These results further support that the WT1 enhancer binds Sp1, and the same form of Sp1 that binds the consensus Sp1 oligonucleotide.


Fig. 13. Southwestern analysis of crude HeLa nuclear extract protein comparing binding to the 31-bp WT1 wild-type enhancer oligonucleotide probe (WT1 probe), the C mutated version, and GC box (26-bp Sp1 consensus probe). Six lanes of HeLa S3 nuclear extract (120 µg/lane) were size separated by SDS-PAGE and transferred to polyvinylidene difluoride membrane (Millipore). Paired lanes were cut from the membrane, hybridized to the different probes (2.7 × 106 cpm/ml) under identical conditions, washed separately, and autoradiographed with membranes re-aligned. A single band of ~105 kDa appears with either the WT1 probe or the Sp1 consensus GC box. Reduced binding to the C mutated probe indicates binding is sequence-specific.
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The WT1 Enhancer Is a High Affinity Sp1 Binding Site

The dissociation constants (KD) of the 31-bp WT1 enhancer and 26-bp Sp1 consensus oligonucleotides for Sp1 protein were determined in EMSAs by titrating an increasing amount of probe against a constant amount of Sp1 in WGA partially purified nuclear extract preparations (Fig. 14A). The amount of bound and free probe were determined by Cerenkov counting of bands excised from the dried, autoradiographed gel. A 140-fold concentration range of probe was tested, including amounts that exceeded by more than 10-fold the observed KD values. Binding was saturable, reaching a plateau of ~0.28 nM with 0.78 nM total probe and beyond (Fig. 14, A and B), and the EC50 was ~0.35 nM. The KD was calculated in Rosenthal (modified Scatchard) plots as the negative reciprocal of the slope (47). The KD of Sp1 for the wild-type WT1 enhancer sequence was 3.7 × 10-10 M (Fig. 14B, inset), and for the Sp1 consensus oligonucleotide was 8.1 × 10-10 M (data not shown). These experiments were performed multiple times, with the WT1 site revealing a consistent 2-fold higher Sp1 affinity than the GC box. Moreover, two different GC box oligonucleotides from different suppliers gave identical results. This observation is also supported by the competition studies (Fig. 12), in which the WT1 enhancer was the better competitor, and Southwestern analysis (Fig. 13), in which equivalent amounts of probe of similar specific activity were used, but in which binding of the labeled WT1 oligonucleotide appears qualitatively better than the consensus oligonucleotide. The published Sp1 KD values for a GC box consensus site (identical to our Sp1 consensus sites) range from 4.1 to 5.3 × 10-10 M using DNase I footprinting and EMSAs (48), which are perhaps no different from what we have observed for the WT1 enhancer site, but which are better than what we have observed for the GC box. It may well be that the relative affinities of Sp1 for different binding sites change under different conditions and, more importantly, that these differences may be biologically relevant. Divalent cation concentrations may be important in this regard, as our studies were performed in the absence of magnesium, which was included in the GC box-Sp1 interaction studies (48, 56), but which we found to interfere with complex formation with the CTC repeat. The correlation we have observed between the function of the wild-type and mutated WT1 enhancers and their ability to bind Sp1 in vitro suggests the conditions we are using for EMSAs provide a good model of the in vivo setting. Other non-consensus sites with Sp1 affinities 10-20-fold lower than the GC box have also been reported (59). The high affinity of Sp1 for the single WT1 enhancer site may help explain its potency, and again underscores the importance of Sp1 to WT1 transcription.


Fig. 14. Determination of dissociation constant (KD) of the WT1 wild-type enhancer oligonucleotide for Sp1 protein. A, EMSA of HeLa S3 cell WGA partially purified Sp1 (100 ng of total protein/lane) incubated with increasing concentrations (shown) of 31-bp WT1 enhancer probe. B, plot of specific probe binding (protein-DNA complex formation) versus total amount of input DNA probe. Probe was quantitated by Cerenkov counting of autoradiographed bands from the dried EMSA gel. Inset, Rosenthal plot of bound/free probe versus bound probe; negative reciprocal of slope = KD.
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We conclude from these experiments that Sp1 binds the WT1 enhancer in vitro and that the enhancer contains a single, near-consensus, high affinity Sp1 site that binds Sp1 at least as well as a consensus GC box. Moreover, in conjunction with the data above, the binding of Sp1 to the wild-type and mutated enhancers in vitro parallels the activation potential of the enhancers, suggesting that the activator is indeed Sp1.

Sp1 Transactivates a Reporter Gene through the WT1 Enhancer

To ascertain the effect of Sp1 on the WT1 enhancer in an Sp1-null setting, we transiently transfected into Drosophila Schneider S2 cells the wild-type and mutated enhancers in the -81/+191 bp context and the -48/+191 bp construct, with and without an Sp1 expression vector. Without Sp1, all the constructs had comparably low absolute reporter activity levels (Fig. 15, black bars). With cotransfection of 100 ng of Sp1 expression vector, reporter activity of the mutated enhancer and enhancerless constructs increased comparably, from ~0.43 to ~2.4%, whereas activity increased from 0.37 to 14.6% (~6-fold more) with the wild-type -81/+191 bp construct (Fig. 15). Therefore, the WT1 enhancer was not functional in the absence of Sp1, and was only active when able to bind Sp1. This result correlates well with the observations in human cells and in EMSAs with human protein, and again supports that the transcriptional activator that exerts its effect on the critical WT1 enhancer is Sp1. Although Sp1 did activate the constructs with mutated enhancers or lacking the enhancer, their activated levels were still only ~16% of the level achieved with the intact -81/+191 bp construct. Therefore, despite that each mutated construct contains 273 bp of 70+% GC-rich WT1 sequence with many potential Sp1 binding sites, more than 80% of Sp1 transactivation is mediated by the intact CTC repeat. This experiment confirms that Sp1 is capable of strongly transactivating a reporter gene linked to the small WT1 enhancer containing a single, alternative Sp1 site.


Fig. 15. Effect of Sp1 on the WT1 proximate promoter, with and without functional enhancer sequence. Drosophila Schneider S2 cells were transiently transfected with 10 µg of reporter DNA in the absence (black bars) or presence of 100 ng Sp1 expression vector (shaded bars). When cotransfected with Sp1, the construct with an intact WT1 enhancer (-81/+191 bp) had ~6 times the reporter activity of the constructs containing mutated enhancers or lacking the enhancer. Mean results of two experiments are reported.
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CTC Repeats in Other Growth-related Genes Also Bind Sp1

Through homology searches, we have identified a number of genes that also contain CTC repeats in their promoters, several at a distance from an identified TSS comparable to that found in WT1. As a group, these genes are transcribed from TATA-less promoters, are tissue-restricted in their expression, and are growth-related, which suggests that, as an Sp1 site, the CTC repeat may serve a unique regulatory role. Genes in this group are egfr (49), pdgf-A chain (62, 63), egr-1 (64), c-myc (65), c-myb (66), and vav (50). Thy-1 has a similar opposite strand site downstream of its TSS (67). To determine whether other CTC repeats were high affinity Sp1 sites, we tested ~40-bp oligonucleotide promoter fragments containing CTC repeats from egfr and the vav proto-oncogene in EMSA and EMSA competition studies. As expected, these other CTC elements were readily capable of forming the same protein-DNA complex as the WT1 element and could compete effectively for Sp1 binding to the WT1 element (data not shown), indicating they too were high affinity Sp1 sites.

Sp1 Expression during Nephrogenesis

Saffer et al. showed that Sp1 tissue levels vary up to 100-fold during murine development, as assessed by immunohistochemistry and RNA dot blotting (68). At postnatal day 32, high Sp1 protein levels were detected in spleen, early hematopoietic cells, testes, and uterine decidua and moderate levels in kidney glomerular cells (68); WT1 is expressed in all these tissues (2, 29, 36, 69-71). High levels of Sp1 were also found in tissues that do not express WT1, such as gastric and lung epithelia and spermatids (68), but no WT1-expressing tissue had little or no Sp1 expression, with the possible exception of testis Sertoli cells. Consequently, we hypothesized that Sp1 might be spatially and temporally regulated in nephrogenesis and thereby account, to some extent, for the WT1 expression pattern.

To determine if a correlation exists between Sp1 and WT1, we examined Sp1 protein expression by immunohistochemistry in day 18 fetal (Fig. 16, A-C) and adult rat kidney (Fig. 16, D-F). At embryonic day 18, all stages of nephrogenesis can be observed (Fig. 16A). Near-contiguous sections were examined for Sp1 immunoreactivity or hematoxylin and eosin staining (data not shown) to allow identification of structures. Control sections stained with the secondary antibody alone were essentially negative (Fig. 16, B and E). In Fig. 16A, more Sp1 is found in the outer, undifferentiated part of the section (o) than in the inner, differentiated region (i). Sp1 staining is most prominent in nuclei of uninduced mesenchyme cells (Fig. 16A, closed arrow), which do exhibit weak WT1 expression (30-32), and early-induced mesenchyme cells (open arrow), which have increasing amounts of expressed WT1 (29). Sp1 staining remains dense in early tubule structures and is reduced in more differentiated cortical tubules (Fig. 16A), becoming barely detectable in mature proximal tubules (Fig. 16, D and F, open arrows). In contrast, Sp1 expression is maintained at moderately high levels in developing (Fig. 16C, center) and mature glomeruli (Fig. 16, D and F, center). The only renal cell type that expresses WT1 in adulthood is the podocyte (30, 31). To determine whether the Sp1-staining glomerular cells included podocytes, we counterstained the Sp1-stained adult rat kidney sections with periodic acid-Schiff reagent (Fig. 16F). Periodic acid-Schiff reagent stains the carbohydrate-rich basement membranes, which allows distinction of glomerular endothelial cells, which reside inside glomerular capillary loops, from podocytes, which are outside the loops. In Fig. 16F, closed arrows denote podocytes staining with moderate Sp1 levels. Glomerular mesangial cells (Fig. 16F, arrowhead), which do not express WT1, had high Sp1 levels. Thus, the WT1-expressing structures have at least moderate levels of Sp1, but not all high Sp1-expressing cells express WT1. Because Sp1 expression completely overlaps that of WT1, Sp1 might therefore be a prerequisite for WT1 expression. In addition, falling Sp1 levels may account for loss of WT1 expression, as observed in developing proximal convoluted tubules (Fig. 16, D and F). These findings support the importance of Sp1 to WT1 expression.


Fig. 16. Sp1 immunoreactivity in fetal (A-C) and adult rat kidney (D-F). A, Sp1 immunohistochemistry of day 18 fetal rat kidney (original magnification, ×25). Developing tubules stain prominently in the outer region (o), but less so in the more mature inner region (i). High Sp1 levels are also found in uninduced mesenchyme (closed arrow) and condensing mesenchyme (open arrow). B, negative control for A showing no background. A comparable day 18 fetal rat kidney section was stained in the absence of the primary Sp1 antibody (shown at same magnification). C, higher magnification view (original magnification, ×100) of Sp1-immunostained day 18 fetal rat kidney showing an immature glomerulus (center). The developing glomerular podocyte layer (arrow) exhibits moderately high Sp1 staining. D, adult rat renal cortex (original magnification, ×100) stained for Sp1. A mature glomerulus (center) has high Sp1 levels, while mature proximal tubules (arrow) have little Sp1. E, negative control section for D performed in the absence of primary antibody shows little background staining. F, periodic acid-Schiff staining of an Sp1-stained section, comparable to D, allowing visualization of glomerular basement membranes. Glomerular podocytes (closed arrows) have moderate Sp1 levels, while proximal tubule cells (open arrow) have low Sp1 levels. Mesangial cells (arrowhead) have high Sp1 levels.
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DISCUSSION

We are reporting several novel findings. First, using an unbiased approach, we have identified a critical Sp1 site in the WT1 promoter that accounts for the majority of its transcriptional activity, and which is the most important enhancer over 24.4 kb of WT1 5'-flanking and first intron sequence, based on these studies in stably transfected cultured cells. Second, although previous work has suggested the CTC repeat may be an Sp1 site (see below), we believe we have provided conclusive evidence that the CTC repeat is an Sp1 site, that it binds the same form of Sp1 as the consensus GC box, and that it does so with an affinity at least as high as the consensus GC box. Third, we have found that similar CTC repeats reside in the immediate 5'-flanking regions of a subset of tissue-restricted, growth-related genes, suggesting this site may be a critical, operationally distinct Sp1 site. Finally, we have shown for the first time that Sp1 expression is temporally and spatially regulated during nephrogenesis in a pattern that not only supports a role in WT1 regulation, but also suggests Sp1 itself may be important in kidney development. These studies indicate that the CTC repeat element and the associated ubiquitous transcription factor Sp1 are critical to expression of the tissue-restricted WT1 gene.

Although ubiquitous, Sp1 may participate in tissue-specific WT1 expression in several ways. Foremost, our immunocytochemistry data indicate that Sp1 is spatially and temporally regulated in nephrogenesis and further support that Sp1 may be necessary for WT1 expression. Similarly, Sp1 is expressed at different levels in gastric (68) and hematopoietic development (72) and influences differential gene expression in these cells (72, 73). Monocyte-specific expression of the CD11b (74) and CD14 promoters (75) also depends on Sp1. Change in Sp1 levels, therefore, is one likely way Sp1 could contribute to tissue-specific WT1 expression. There is also growing evidence of other mechanisms through which Sp1 may influence differential gene expression. A great number of Sp1-interacting proteins have been identified, many of which are tissue-restricted transcription factors, such as GATA-1 (72, 76), GATA-2, and GATA-3 (76), HNF4 (77), the retinoic acid (78), triiodothyronine (79), and estrogen receptors (80), Ets-1 (81), and MyoD (82). Such factors, perhaps bound to elements beyond the 24-kb region we examined, may interact with Sp1 at the critical promoter site we have identified. For example, a GATA-1-responsive enhancer was found in the WT1 3' region (38, 83), outside the sequences we tested. GATA-1 that binds the WT1 enhancer in hematopoietic cells, or GATA-3 in the kidney (84), might interact with promoter-bound Sp1. We have also observed that the CTC repeat confers a striking 20-fold activation in 786-O cells,3 a renal cell carcinoma line lacking a functional copy of the von Hippel Lindau (VHL) tumor suppressor gene (86). This activation is ~3-fold higher than that in any other transiently transfected cell line3 and suggests the VHL gene product might normally inhibit the CTC enhancer or Sp1 itself. A functional relationship between these two transcription factors, which we are examining, would suggest that differential gene expression involving Sp1 may in addition include interactions with other ubiquitous factors with differing tissue levels. Sp1 might also affect WT1 expression in other ways. Sp1 levels and/or activity can be increased by oncogene expression (87-89), growth factors (90-92), or cytokines (93, 94). Whether any of these stimuli affect WT1 expression remains unexplored.

Somewhat surprisingly, we and others (37, 38) have found that the WT1 promoter-reporter constructs are transcriptionally active when transfected in either non-WT1-expressing or WT1-expressing cell lines. In contrast, we have used a similar construct (-1.9/+0.2 kb WT1-lacZ) to generate four transgenic mouse lines and were unable to detect transgene expression in any mouse tissue, despite having observed expression in transfected cultured cells.3 What are possible explanations for this discrepancy in transfected versus transgenic WT1-reporter expression? In cell lines, it is well recognized that stably integrated DNA exists in an "active" conformation (95-98), which would favor ubiquitous expression. Selection of neo-resistant clones requires that integration occur in a transcriptionally active site, and co-transfected DNA should integrate there as well (99, 100). WT1 may also be regulated by DNA methylation, as the gene contains CpG islands (17, 18). Lack of methylation of transfected DNA may have contributed to its ubiquitous expression (98, 101). A WT1 intron 3 silencer (102) was not included in our constructs and therefore would not have accounted for the difference in expression between transgenic and transfected cells. Some additional repressive chromatin feature may form in transgenic cells subjected to differentiation forces that cannot form on transfected DNA in a monomorphic cell line. The cell lines' endogenous WT1 genes do appear to be subject to a higher level of regulation, however. We mapped the DNase I-hypersensitive sites of the endogenous WT1 locus in these lines and found differential hypersensitivity of the WT1 promoter.2 In sum, although we do not have WT1 sequence that confers tissue-specific expression, we have identified the key non-tissue-specific cis element that resides in a promoter region accessible only in WT1-expressing cells.

Based on its high GC content, the WT1 promoter was previously considered as a target for regulation by zinc finger transcription factors (35, 103, 104). Rupprecht et al. (104) used large amounts of CMV-driven WT1 expression vectors to show that WT1 negatively autoregulates murine WT1 promoter-reporter constructs. The WT1-mediated repression depended predominantly on sequence from +31 to +201 bp, although DNase I footprinting over the -513 to +201 bp region using purified WT1 protein revealed seven protected regions spanning 25-70 bp each, including the homologous CTC repeat we identified ((104), Fig. 1A). Similarly, Malik et al. (103) examined negative autoregulation of the human WT1 promoter using an inducible WT1 system and found that high WT1 protein levels reduced reporter activity ~50% and that repression also depended on WT1 promoter sequence from +38 to +195 bp. Therefore, WT1 may well autoregulate its own promoter, but high WT1 protein levels are required. Finally, in their initial description of the GC-rich WT1 promoter, Hofmann et al. (35) tested whether Sp1 might be important for its activity. Over the -453 to +191 bp region, purified Sp1 protein protected nine elements, 15-27 bp in size. The essential CTC repeat we identified was mostly protected in this assay, but there was no indication of its importance. Cotransfection of 10 µg of CMV-driven Sp1 with a similar -453/+191 bp construct increased reporter expression about 3-fold (35). Our work, in contrast, identifies as the major enhancer of the WT1 promoter a single high affinity Sp1 site. Moreover, mutation of this site (Fig. 6) indicates that endogenous amounts of Sp1 acting through this CTC repeat account for ~80% of WT1 promoter activity.

CTC repeats similar to the WT1 element are found in the proximate promoters of other growth-related genes (see above). The egfr immediate 5'-flank contains four closely spaced CTC repeats that are S1 nuclease-sensitive (105). These elements are important to overall egfr promoter transcription, as their deletion reduced egfr promoter-reporter activity 3-5 fold (105). These sites were considered to bind Sp1, as well as an unidentified factor, because purified Sp1 protein footprinted this region (105). The c-myc gene promoters were subjected to DNase I footprinting, and a number of the footprints identified with crude nuclear extract could be reproduced with purified Sp1 protein, including a similar CTC repeat (65). However, these authors did not demonstrate, as we have, that the crude nuclear extract protein that binds the CTC repeat is Sp1.

CTC repeats may even be targets for transcriptional repression by WT1 protein, but large amounts of WT1 expression vector are required (24, 103, 104, 106) (see above). In contrast, we have not observed WT1, but only Sp1 binding to the CTC repeat, even in crude nuclear extracts from high WT1-expressing Wilms' tumor (gift of A. J. Garvin, Medical University of South Carolina), LP9 mesothelial cells (2) (gift of J. Rheinwald), or fetal kidney, supporting that Sp1 preferentially binds the CTC repeat.3 It is still intriguing, however, that the CTC repeat is a potential site for Sp1-WT1 competition, as has been reported elsewhere for Sp1 with Egr-1 (107) and other factors (79, 108-110). Although sequence other than the CTC-repeat has been shown necessary for WT1-mediated repression (103, 104, 106, 111), two WT1 sites, one 5' and one 3' of the TSS, may be required for repression, as proposed by Wang et al. (112). Because WT1 homodimerizes (113), the presence of adjacent WT1 binding sites may increase WT1's affinity for the sites. Most convincingly, transfected WT1 represses the endogenous egfr gene, and this effect is mediated by CTC repeats (24). The ability to localize either Sp1 or WT1 near a TSS may therefore be a critical role of the CTC repeat.

Might the CTC repeat element itself serve a regulatory function? CTC repeats and similar polypyrimidine tracts can exist in non-B DNA structural conformations (85, 114-116), which have long been proposed to act as transcriptional regulatory elements. Because these structures permit some single-strandedness, polypyrimidine tracts may exhibit S1 nuclease sensitivity (28, 85, 106, 114-116) and therefore be particularly accessible to transcription factor binding. However, the WT1 CTC repeat, its surrounding sequence (Fig. 1A), and even the adjacent plasmid DNA do not contain features typical of polypyrimidine tracts with higher order structure. For example, no direct repeat of the (CTC)3 sequence is found nearby that might serve as a target for DNA slippage, as proposed for several promoters (85, 115), including that of egfr (106). No inverted repeat described for cruciform structures (28) is found either. In addition, unlike the WT1 region, the continuous polypyrimidine tracts that have been shown to be S1 nuclease sensitive are long simple repeats, i.e. 45 bp of d(TC)n·d(GA) (116) and 33 bp of d(GA)n·d(TC)n (114). For these reasons, the (CTC)3 element function is most likely attributable to interacting transfactors.

In conclusion, we have identified a critical enhancer in the WT1 gene as a single, alternative Sp1 site located immediately upstream of the start of WT1 transcription. This site, a CTC repeat, exhibits at least as high an Sp1 affinity as the consensus GC box and is found in other TATA-less, tissue-restricted genes. The Sp1 expression pattern during nephrogenesis lends support to Sp1 serving a key role in WT1 transcription and perhaps in regulation of other genes critical in nephrogenesis.


FOOTNOTES

*   This work was supported by National Institutes of Health grants K08 DK02280 to HTC, F32 DK09263 to SAB, and R01 DK45617 to VPS. The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EMBL Data Bank with accession number(s) U77682[GenBank].


Dagger    To whom correspondence should be addressed: Renal Division, Beth Israel Hospital and Harvard Medical School, Dana 517, 330 Brookline Ave., Boston, MA 02215. Tel.: 617-667-2105; Fax: 617-667-7843; E-mail: vsukhatm{at}bih.harvard.edu.
1    The abbreviations used are: kb, kilobase(s); bp, base pair(s); CAT, chloramphenicol acetyltransferase; CMV, cytomegalovirus; TSS, transcription start site(s); EMSA, electrophoretic mobility shift assay; WGA, wheat germ agglutinin; WAGR, Wilms' tumor, aniridia, genitourinary abnormalities, and mental retardation.
2    S. A. Bossone, H. T. Cohen, and V. P. Sukhatme, unpublished data.
3    H. T. Cohen and V. P. Sukhatme, unpublished data.

Acknowledgments

We thank H. Rennke for review of immunohistochemistry results; K. Call, D. Haber, S. Patwardan, J. Rheinwald, M. Segal, and R. Tjian for reagents; and S. Jackson, J. Saffer, and D. Tenen for valuable advice.


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