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Annals of Surgical Oncology 10:136-143 (2003)
© 2003 Society of Surgical Oncology

ORIGINAL ARTICLES

Fine Mapping of Wilms’ Tumors With 16q Loss of Heterozygosity Localizes the Putative Tumor Suppressor Gene to a Region of 6.7 Megabases

Shawn D. Safford, MD, Dominique Goyeau, MS, Alex J. Freemerman, PhD, Rex Bentley, MD, Mary Lou Everett, MS, Paul E. Grundy, MD and Michael A. Skinner, MD

From the Department of Surgery (SDS, DG, AJF, MLE, MAS), Division of Pediatric Surgery, and Department of Pathology (RB), Duke University Medical Center, Durham, North Carolina; and Department of Pediatric Oncology (PEG), Cross Cancer Institute, Alberta, Canada.

Correspondence: Address correspondence and reprint requests to: Michael A. Skinner, MD, PO Box 3815, Duke University Medical Center, Durham, NC 27710; Fax: 919-681-8353; E-mail: skinn009{at}mc.duke.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Background: The aim of this study was to more precisely map the region of 16q loss of heterozygosity (LOH) in Wilms’ tumors and to examine the expression of putative tumor suppressor.

Methods: We performed polymerase chain reaction–based LOH analysis on the 185 sample pairs from 21 to 80 megabases (Mb) on chromosome 16q. Expression of two candidate tumor suppressor genes located within the identified consensus region of 16q LOH was examined by immunohistochemistry.

Results: We identified 16q LOH in 7 (4%) of 185 Wilms’ tumors not previously thought to demonstrate such genetic loss. The smallest common region of genetic loss was located between 67.3 and 74.0 Mb on chromosome 16. Within this 6.7-Mb region, there reside only three recognized tumor suppressor genes: E-cadherin, P-cadherin, and E2F4. E-cadherin demonstrates statistically significantly reduced expression in Wilms’ tumors with 16q LOH.

Conclusions: We have localized the consensus region of 16q LOH in Wilms’ tumor to a 6.7-Mb locus and have identified three candidate Wilms’ tumor suppressor genes within this narrowed region. Our data support E-cadherin as a candidate tumor suppressor gene in Wilms’ tumor; however, further studies are needed to definitively prove its role as the tumor suppressor gene associated with 16q LOH.

Key Words: Wilms’ tumor • Loss of heterozygosity • Chromosome 16 • Neoplasia • Tumor suppressor


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Wilms’ tumors frequently exhibit nonrandom chromosomal deletions at sites on chromosomes 1, 11, and 16, and these areas of genetic loss have been examined for the presence of tumor suppressor genes.1,2 Indeed, through a combination of cytogenetic and molecular techniques, the Wilms’ tumor suppressor genes WT1 and WT2 have been localized to chromosome 11p13 and chromosome 11p15, respectively. Assays for chromosomal loss of heterozygosity (LOH) have been especially helpful in this regard.3–5

The long arm of chromosome 16 exhibits LOH in 17% to 20% of Wilms’ tumors.6,7 Two studies have shown that 16q LOH correlates with a poorer prognosis, suggesting there is a gene or genes in the region whose loss increases tumor aggressiveness.7–9 Whereas most investigations have found only large areas of loss on 16q by using widely spaced genetic markers, a more recent study has localized the region of 16q LOH to two interstitial deletions: these two areas of loss were localized between 38.9 and 75.2 megabases (Mb) and between 80.7 and 112.9 Mb on chromosome 16.2

The goal of this study was to more precisely narrow the genetic area of 16q LOH so that we could identify the specific tumor suppressor genes in the region that may be responsible for the aggressive behavior in a subset of Wilms’ tumors. Using narrowly spaced microsatellite repeat markers, we refined the consensus area of 16q LOH to 6.7 Mb, and within this region were three candidate tumor suppressor genes. Moreover, we examined the expression pattern of the most likely 16q tumor suppressor candidates E-cadherin and P-cadherin in a set of 18 Wilms’ tumors.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Subjects
All DNA samples were acquired from the National Wilms’ Tumor Study (NWTS) Group Biological Samples Bank. A total of 196 normal and tumor DNA pairs were selected from a group that had been evaluated previously as part of the Pediatric Oncology Group study 9046, "A Molecular Genetic Analysis of Wilms’ Tumors." Eleven samples were previously shown to demonstrate 16q LOH through screening in the NWTS core laboratory with six widely spaced microsatellite repeat markers. In this study, we evaluated the remaining 185 pairs of tumor and normal DNA for 16q LOH to detect interstitial deletions too small to be identified in the core laboratory.

Eighteen additional tissue samples for immunohistochemical analysis were also acquired from the NWTS Group Biological Samples Bank. Stage, histology, and presence of LOH on chromosomes 1p, 11p, and 16q were determined by NWTS before our study. Nine tissue samples were known to exhibit 16q LOH, and nine demonstrated retention at all the tested 16q loci. We were unable to evaluate three of the retention group because of extensive necrosis within the sample. The remaining 15 samples were evaluated by immunohistochemical staining for expression of the E-cadherin and P-cadherin proteins, and DNA was extracted and used to evaluate for CpG methylation of the E-cadherin promoter. The investigation was approved by the Duke University Institutional Review Board.

DNA Analysis and Allelotyping
Polymerase chain reaction (PCR)-based LOH analysis was performed through amplification of polymorphic microsatellite repeat markers. For this study, we used 21 narrowly spaced markers to evaluate 185 Wilms’ tumors that had not demonstrated LOH in an initial screen at the NWTS core laboratory. The primers were located between 21 and 80 Mb on chromosome 16q (Table 1), covering the narrowest region of 16q LOH that had previously been described.2 The locations of primers and the genes that lie between microsatellite repeat markers were determined by Map Viewer at the National Center for Biotechnology Information (NCBI) Web site (http://www.ncbi.nih.gov/Tools/index.html).


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  		TABLE 1. Polymorphic microsatellite repeat markers used to map the area of 16q LOH
 
The forward primer was conjugated to a rhodamine dye for analysis on the ABI PrismTM 377XL DNA sequencer (Applied Biosystems, Foster City, CA). PCR was performed in 25-µl reactions containing 2.5 µl of AmpliTaqTM 10x buffer, 20 ng of DNA, .2 mM of each deoxynucleotide triphosphate, 100 ng of each primer, and .5 to 1 U of AmpliTaq GoldTM polymerase (PerkinElmer, Boston, MA) and the optimal MgCl2 concentration. Each PCR reaction was started with a 10-minute "hot start" at 95°C. The 50 cycles proceeded with a 1-minute denaturation at 94°C, with an annealing temperature, depending on the primer, between 50°C and 65°C for 30 seconds, and elongation for 30 seconds at 72°C.

The samples were separated on a 4%, 36-cm, .2-mm polyacrylamide gel in an ABI Prism 377XL DNA sequencer. The samples were diluted with water from 1/4 to 1/13, depending on the rhodamine conjugate, and 1 µl of the diluted sample was mixed with 3 µl of loading mix containing 5:1:1 formamide, Tamara GeneScan-500TM size standard (Applied Biosystems), and loading dye. The mixed samples were denatured at 90°C for 2 minutes immediately before loading. The wells were flushed and loaded with 1 to 3 µl of sample. The resulting gel file was analyzed with the ABI Prism GeneScan AnalysisTM software (Applied Biosystems). The normal and tumor DNA peaks were compared and scored with regard to the presence of allelic homozygosity, the retention of heterozygosity, or LOH. Retention of heterozygosity was defined as the presence of two alleles in each of the normal and tumor samples. If only one allele was seen in the normal DNA, the patient was homozygous at the locus, and this was noninformative in determining whether there was genetic loss. LOH was scored if the peak height of one of the tumor alleles was <40% of the height when compared with the corresponding allele in the normal DNA.

E-Cadherin and P-Cadherin Immunohistochemistry
Wilms’ tumor tissue samples from NWTS were stained with E-cadherin mouse immunoglobulin (Ig)G2a (BD Transduction Laboratories, Lexington, KY) and P-cadherin mouse IgG1 (BD Transduction Laboratories, Lexington, KY). Tissues were stained by using standard protocols from BioGenex Laboratories (San Ramon, CA). Briefly, the deparaffinized tissues were blocked with Ready-to-Use Peroxide BlockTM (BioGenex Laboratories), and the primary antibody was applied at a dilution of 1/50 for both E-cadherin and P-cadherin. The signal was amplified by sequentially applying a biotinylated secondary rat anti-IgG mouse antibody followed by a streptavidin–horseradish peroxidase tertiary antibody (Multi-Link Super Sensitive KitTM; BioGenex Laboratories). The slides were then developed with 3,3'-diaminobenzidine tetrahydrochloride and counterstained with hematoxylin. Negative controls were performed on all tissue samples by using the same conditions without the primary antibody for E-cadherin and P-cadherin.

The stained slides were evaluated by an independent pathologist who was blinded to the chromosomal pattern of each Wilms’ tumor sample in the analysis of both E-cadherin and P-cadherin. E-cadherin scoring was based on the percentage of positively staining tubular cells. The standard scoring system for E-cadherin consisted of 0, no staining; 1, <33% of tubular cells stained; 2, 34% to 66% of tubular cells stained; and 3, >67% of tubular cells stained. Staining intensity was not evaluated for E-cadherin because of the homogeneous strength noted in those cells that stained positive. P-cadherin expression was scored on the basis of the staining intensity and pattern for each of the cellular subcomponents of Wilms’ tumors. Staining intensity was scored on a standard of 0, no staining; 1, weak; 2, moderate; and 3, strong. The pattern of expression was scored on the basis of 0, no staining; 1, focal staining; and 2, diffuse staining. The tumors were then grouped on the basis of the presence or absence of 16q LOH, and an average score was calculated for each group.

Analysis of CpG Methylation of E-Cadherin
We used the CpGenome DNA Modification KitTM (Intergen Co., Purchase, NY) to perform the bisulfite modification reaction. DNA samples prepared from the tumors that had undergone immunohistochemical evaluation were used in this analysis. We performed the bisulfite reaction exactly as described in the CpGenome DNA Modification Kit.

To amplify DNA from the bisulfited samples, we used the CpG Wiz Amplification KitTM (Intergen Co.). Each DNA sample was amplified by using three primer sets: unmethylated, methylated, and wild-type. The unmethylated primer set will anneal to unmethylated DNA that has undergone the bisulfite chemical modification that we performed previously. The methylated primer set will anneal to methylated DNA that has undergone the bisulfite chemical modification. The wild-type primer set serves as the control for the efficiency of the DNA bisulfite modification and will anneal to any DNA, methylated or unmethylated, that has not undergone the chemical bisulfite modification. The chemical modification step was considered adequate if the wild-type primer sample was <50% intensity of the unmethylated and methylated samples for each DNA sample.

Each 25-µl reaction mixture consisted of 2.5 µl of 10x Amplitaq (PerkinElmer, Boston, MA) PCR buffer, 2.5 µl of deoxynucleotide triphosphate mix, 1 µl of primer mix, .2 µl of AmpliTaq Gold polymerase, and 2 µl of bisulfite modified DNA (50 ng/µl). These were then placed in the thermocycler block, and PCR was performed with a hot start of 95°C for 10 minutes and then 35 cycles of denaturing at 95°C for 45 seconds and annealing at 60°C for 45 seconds, followed by extension at 72°C for 60 seconds.

Statistics
All statistics were analyzed with StatsDirectTM (CamCode, Cambridge, UK) statistical analysis software. We used the Mann-Whitney U-test to evaluate the difference between the E-cadherin and P-cadherin staining scores in groups of Wilms’ tumors with retention and with 16q LOH. We defined statistical significance as a P value of <.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Physical Region of Genetic Loss in Wilms’ Tumors With 16q LOH
One hundred eighty-five tumor and normal DNA sample pairs were obtained that had been previously screened for 16q LOH with widely spaced microsatellite genetic markers. No genetic loss was identified in this previous analysis; however, we reasoned that by using more closely spaced markers, we could identify smaller interstitial regions of genetic loss located between the markers used in the initial screening study. We used 21 microsatellite repeat markers to examine the samples. We performed our initial study by using 8 moderately spaced markers to screen all 185 samples covering an area from 21 to 80 Mb (Table 1, group 1). This analysis showed two tumors with genetic loss in the region between 50 and 80 Mb, so we re-evaluated all 185 samples by using an additional 13 markers covering only this region to identify the smallest possible length of genetic loss (Table 1, group 2). This focused use of the microsatellite repeat markers identified seven tumors with discreet areas of loss that had not been previously shown to contain LOH. Figure 1 shows these seven tumors with areas of genetic loss and retention. The area of common LOH within these tumors was between 67.3 and 74.0 Mb.



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  		FIG. 1. Common area of 16q loss of heterozygosity (LOH). These figures represent the common area of 16q LOH that was discovered through LOH analysis. (A) This diagram represents tumor sample 50067 with its corresponding narrowed region of 16q LOH. The arrows point to the physical locations of the microsatellite repeat markers on chromosome 16q. The plots below the arrows represents the data generated by the ABI Prism sequencer, with normal DNA at the top of each plot and tumor DNA on the bottom. D16S2620 and D16S2624 demonstrate retention of the DNA segments, and primers D16S3043 and D16S752 exhibit loss of heterozygosity at their loci. The dark band represents the smallest area of 16q LOH in this tumor. (B) This figure shows the seven Wilms’ tumor samples that were found to exhibit discrete interstitial areas of 16q LOH. The solid black area represents the area of LOH, with the smallest consensus region extending from 67.4 to 74.0 Mb. The location of each candidate tumor suppressor gene is shown by an arrow, and its physical map location is listed under each name.

 
Within this 6.7-Mb region of common LOH on 16q, 42 identified and 129 hypothetical genes were described, according to the analyzed human DNA sequence data on the NCBI Web site. Of the 42 identified genes, only 3 genes were previously known to be tumor suppressor genes. These tumor suppressor genes include E-cadherin, P-cadherin, and E2F4.10,11 On the basis of previous studies in gastric, breast, bladder, and endometrial cancers, we proposed that E-cadherin, P-cadherin, or both would be the candidate tumor suppressor genes most likely to be responsible for a poorer prognosis in a subset of Wilms’ tumors.10–14

Expression of the Candidate Tumor Suppressor Genes E-Cadherin and P-Cadherin
We might reasonably expect decreased protein expression of the relevant 16q tumor suppressor gene in Wilms’ tumors with 16q LOH compared with tumors not exhibiting such genetic loss. Thus, we investigated expression of the candidate genes, P-cadherin and E-cadherin, to further investigate their potential contribution to Wilms’ tumor biology. Using an immunohistochemical technique, we evaluated the expression of E-cadherin and P-cadherin in nine Wilms’ tumors with and six tumors without 16q genetic loss.

E-cadherin was expressed only in the tubular subcomponent of Wilms’ tumors, and there was no staining in the blastemal or stromal regions of the tumors. Therefore, we examined the percentage of tubular cells expressing E-cadherin in Wilms’ tumors with and without 16q LOH. Wilms’ tumors with 16q LOH had significantly fewer tubular cells that stained positive for E-cadherin (P = .01; two-sided Mann-Whitney U-test). Interestingly, E-cadherin staining within cells was either extremely strong or completely absent. Thus, Wilms’ tumors with 16q LOH demonstrated a mosaic pattern of E-cadherin expression (Fig. 2) in which tubular cells in the same tubular complex had either strong expression or a complete absence of E-cadherin expression. Tumors with 16q LOH had a lower average E-cadherin score compared with tumors with 16q retention (Table 2). These results support the candidacy of E-cadherin as the Wilms’ tumor suppressor gene that is responsible for the worsened prognosis and decreased relapse-free survival associated with 16q LOH.



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  		FIG. 2. E-cadherin and P-cadherin staining pattern of the tubular components of Wilms’ tumor. These photographs depict the E-cadherin and P-cadherin staining patterns seen in Wilms’ tumors with 16q loss of heterozygosity (LOH) and retention (400x). The E- and P-cadherin–positive cells stain brown, and cells are blue. (A) The differential tubular staining can be appreciated between tubular complexes in the same area within the tumor exhibiting 16q LOH. (B) This is in contrast to the uniform staining pattern that most tumors with retention of 16q LOH demonstrated. (C) This photograph represents the positive control for tubular E-cadherin staining in normal kidney. (D) P-cadherin demonstrates increased staining in samples with 16q LOH. (E) Wilms’ tumors with retention of 16q express less P-cadherin than tumors with loss of 16q. (F) This photograph shows the positive uniform staining of P-cadherin in the normal kidney tubules.

 

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  		TABLE 2. E-cadherin expression in Wilms’ tumorsa
 
In many neoplasms in which there is LOH that contributes to the physical deletion of a tumor suppressing gene, there is also inactivation of the nondeleted allele to allow unchecked tumor growth. One common method of gene inactivation of E-cadherin is through hypermethylation of its promoter region. To investigate the possible mechanism responsible for downregulation of the nondeleted E-cadherin allele, we performed CpG methylation analysis of the Wilms’ tumors. There are six CpG islands in the E-cadherin promoter; the third CpG island, in particular, plays a key role in E-cadherin downregulation in other tumors.13,15,16 We used a PCR-based method to examine CpG methylation in the third CpG island in Wilms’ tumors. We analyzed all 18 of the Wilms’ tumors from the immunohistochemistry studies for methylation patterns. Nine of the tumors had 16q genetic loss, and the remaining nine specimens did not have such genetic loss. Fifteen of 18 tumors showed no evidence of CpG methylation in island 3, whereas 3 of the tumors in the 16q retention group yielded DNA unsuitable for analysis (Fig. 3).



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  		FIG. 3. Analysis of CpG methylation in Wilms’ tumors. This figure represents polymerase chain reaction–based analysis of CpG methylation of Wilms’ tumors. This gel contains all of the tumor samples that had the lowest E-cadherin expression. Each sample was amplified with the U (unmethylated) primer, the M (methylated) primer, and the W (wild-type) primer. Each lane is labeled with these primers. The arrows indicate the size standards for the gel. No methylation was present in any of the samples. bp, base pairs.

 
P-cadherin is another known tumor suppressor gene within the narrowed region of 16q LOH. We used immunohistochemical staining to assess P-cadherin protein expression in the nine Wilms’ tumors with 16q LOH and compared the results with those of six tumors with 16q retention. In these experiments, variations were noted in the intensity and pattern of expression in both the tubular and blastemal cells. P-cadherin expression was significantly higher in both the blastemal and tubular cells of the Wilms’ tumors with 16q LOH (Fig. 2). The staining scores for the 16q LOH tumors averaged 2.71 for the blastema and 1.63 for the tubules, and for the tumors without genetic loss, the staining scores averaged 1.33 for the blastema and .70 for the tubules (P < .01). The stromal subcomponent is relatively acellular and had barely appreciable staining levels. The increased expression in tumors with 16q LOH argues against P-cadherin as the important 16q candidate Wilms’ tumor suppressor gene.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Wilms’ tumors have been shown to lose large segments of chromosomes 1p, 11p, and 16q. These genetic losses were generally identified through assays for the LOH, and the physical loss of an allele is generally thought to constitute the first hit of a two-hit phenomenon resulting in the inactivation of a tumor suppressor gene.2–4,6,7,9,17 Investigations of the specific areas of chromosomal loss have led to previous discoveries of tumor suppressor genes, including WT1 and WT2 on chromosome 11p.2,3

We have focused our efforts on chromosome 16q because of the strong evidence that 16q LOH affects relapse-free survival and prognosis in children with Wilms’ tumor.7–9 Previous investigators have shown that 16q LOH is present in 17% to 24% of Wilms’ tumors.7,9,18 In most cases, the entire long arm of chromosome 16 was deleted. Because so much genetic material and so many genes were deleted, few clues were elicited to help identify precisely which critical gene or genes were lost to account for the increased Wilms’ tumor aggressiveness. In an earlier study mapping the region of 16q LOH in a large set of Wilms’ tumors, we narrowed this area of 16q LOH to two consensus regions on chromosome 16q between 38.9 to 75.2 Mb and 80.7 to 112.9 Mb.2 In this study, we have further narrowed the area of 16q LOH to a 6.7-Mb locus.

According to the NCBI database, this small consensus area contains 42 identified and 129 hypothetical genes. Of the described and hypothetical genes, there are only three known tumor suppressor genes in this area: E-cadherin, P-cadherin, and E2F4. We have focused on the most widely known tumor suppressor genes, E-cadherin and P-cadherin, in an attempt to establish or exclude their contribution to the increased aggressiveness of Wilms’ tumors with 16q LOH. Once we conclusively evaluate these genes, we will continue our systematic evaluation of the other known genes within our consensus area.

The E-cadherin complex participates in cell-cell adhesion and maintenance of normal cellular architecture; disruption of the complex can result in increased tumor aggressiveness. For example, in vitro and animal studies have shown that reduced cell-cell adhesion can lead to invasion and metastasis.12,19 Moreover, reduced E-cadherin expression has been associated with increased aggressiveness in human gastric and breast cancer.12,13

The contribution of the cadherins to tumor aggressiveness can be understood by examining the cell-cell adhesion mechanism. E-cadherin selectively binds to other E-cadherin molecules through its extracellular domain. Intracellularly, E-cadherin interacts with both ß and {gamma} catenins that bind to actin through {alpha} catenin. This complex together maintains the three-dimensional architecture of the tumor, and if any of the subcomponents of the E-cadherin complex are lost or mutated, the normal cellular architecture becomes disrupted, and the cells can potentially invade surrounding tissue and metastasize.19,20 Indeed, previous studies in Wilms’ tumors have shown frequent mutations in the ß-catenin gene, further supporting the importance of the E-cadherin system in controlling Wilms’ tumor aggressiveness.21 In this study, we present additional evidence for the biological importance of E-cadherin in Wilms’ tumors, noting that there is a significant reduction in the number of cells expressing the protein in tumors with 16q LOH. However, this does not conclusively prove E-cadherin’s role as the tumor suppressor gene involved with 16q LOH.

Multiple mechanisms for downregulation of E-cadherin expression have been described in breast, liver, and gastric cancer.10–13,16 These tumors demonstrate apparent epigenetic regulation of E-cadherin expression. The most commonly described method of E-cadherin downregulation is through hypermethylation of the E-cadherin 5' CpG island.12 Our data support this mechanism of an epigenetic overlay that accounts for the nonclonal mosaic expression pattern seen in Wilms’ tumor samples. In our study, however, we did not find any evidence of hypermethylation in any of the Wilms’ tumors, regardless of retention or LOH of 16q. Similarly, in hepatocellular carcinoma, the hypermethylation pattern does not correlate with downregulation of E-cadherin.22 Other mechanisms exist that could cause decreased E-cadherin expression, such as gene mutation or alteration in the posttranslational management of the gene product.

P-cadherin has not been well characterized, but on the basis of its homology with the cadherin gene family, P-cadherin is another candidate tumor suppressor gene located in the region of 16q lost in some Wilms’ tumors. In our studies, P-cadherin expression was increased in tumor samples with 16q LOH. These data argue against P-cadherin as the tumor suppressor gene associated with 16q LOH. A similar pattern of increased P-cadherin expression has been seen in breast carcinoma, where P-cadherin expression is associated strongly with poor survival and constitutes an independent prognostic factor for clinical outcome.11 Further study is needed to elucidate the mechanism by which P-cadherin is involved in tumorigenesis and mutation, and, in particular, we need to further elucidate the role that P-cadherin plays in the clinical outcome of Wilms’ tumors.

In conclusion, our study has narrowed the area of 16q LOH to a region of 6.7 Mb between 67.3 and 74.0 Mb. We have preliminary data suggesting that E-cadherin may be the tumor suppressor gene associated with 16q LOH; however, the mechanism for reduced expression is unclear at this time. Although we have demonstrated a statistically significant reduction in E-cadherin staining in those tumors with 16q LOH, these data do not definitively demonstrate a connection between E-cadherin expression and aggressiveness in Wilms’ tumors. We will need to correlate E-cadherin expression with relapse-free survival and outcome to definitively implicate E-cadherin as the 16q tumor suppressor gene in Wilms’ tumors.


   Footnotes
 
We have localized a consensus region of 16q loss of heterozygosity in Wilms’ tumors to a 6.7-megabase locus through polymerase chain reaction-based microsatellite analysis. Three known tumor suppressor genes reside within this region: E-cadherin, P-cadherin, and E2F4.

Received for publication March 15, 2002. Accepted for publication October 2, 2002.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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