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Genetic changes associated with relapse in favorable histology Wilms tumor: A Children’s Oncology Group AREN03B2 study

Summary Over the last decade, sequencing of primary tumors has clarified the genetic underpinnings of Wilms tumor but has not affected therapy, outcome, or toxicity. We now sharpen our focus on relapse samples from the umbrella AREN03B2 study. We show that over 40% of relapse samples contain mutations in SIX1 or genes of the MYCN network, drivers of progenitor proliferation. Not previously seen in large studies of primary Wilms tumors, DIS3 and TERT are now identified as recurrently mutated. The analysis of primary-relapse tumor pairs suggests that 11p15 loss of heterozygosity (and other copy number changes) and mutations in WT1 and MLLT1 typically occur early, but mutations in SIX1, MYCN, and WTX are late developments in some individuals. Most strikingly, 75% of relapse samples had gain of 1q, providing strong conceptual support for studying circulating tumor DNA in clinical trials to better detect 1q gain earlier and monitor response.


In brief
Wilms tumors have many potential driver mutations. Gadd et al. focus on samples from relapse tumors revealing that over 40% contain mutations in SIX1 or in MYCN network genes, drivers of progenitor proliferation, and that 75% have 1q gain, supporting optimization of new tools for identifying 1q gain.

INTRODUCTION
The goal of this study is to identify genomic characteristics of relapse in Wilms tumors to better identify those at risk of relapse and to better understand the biology of relapse. Wilms tumor (WT; nephroblastoma) is the most common pediatric renal tumor; approximately 95% are of favorable histology (FHWT), and these are the focus of this study. The remaining 5% show histologic evidence of anaplasia, commonly associated with mutations or deletions of TP53. 1 Individuals with FHWT are treated with a chemotherapy backbone including vincristine and actinomycin (stage I and II individuals); doxorubin, cyclophosphamide, and etoposide may be added for stage III/IV individuals, and advanced-stage individuals also receive radiation to sites of disease. 2 Although individuals with FHWT enjoy an overall survival rate of 90%, this comes at a considerable cost, particularly for those with advanced disease. 3 The priority is to identify biological factors that would improve our ability to predict relapse, better tailor relapse therapy, and reduce the significant toxicity associated with relapse therapies.
Two genetic loci have long been associated with the pathogenesis of FHWT, the WT1 gene located at 11p13 and the IGF2/H19 imprinted region on 11p15; each locus is associated with syndromes when present in the germline (reviewed in Gadd et al. 4 ). Loss of imprinting (LOI) or loss of heterozygosity (LOH) of 11p15 is observed in the considerable majority of all WTs and results in overexpression of IGF2. 5,6 Although 11p15 imprinting abnormalities clearly play a critical role in WT development, the observation of 11p15 LOH in normal tissue from some individuals with WT 7 and the lack of tumors arising in mutant mice with LOI of the imprint control region 8 imply that biallelic expression of IGF2 alone is insufficient for tumor development.
Over the last decade, a number of investigators have reported next-generation sequencing of large numbers of WTs. [9][10][11][12][13] These studies indicate that WTs typically arise after acquisition of more than one genetic event. Rather than a limited number of shared driver mutations, WTs have a large number of candidate driver genes that have in common functional involvement in early renal development, often through epigenetic regulation of transcription (chromatin modifications, transcription elongation, and microRNAs [miRNAs]). The most highly represented mutations have been identified in WT1, DROSHA, DGCR8, SIX1/SIX2, CTNNB1, FAM123B (WTX and AMER1), and MYCN. However, only half of individuals with FHWTs have mutations in one of these genes, and many FHWTs lack clear driver mutations. This observation prompted analysis of high-resolution SNP arrays of large numbers of WTs to identify additional regions recurrently gained and lost. [14][15][16] This revealed several recurrent genetic regions of gain or loss, but the underlying pathogenetic genes and/or pathways remain elusive for most loci.
A molecular feature that has been used to stratify treatment of FHWTs in clinical trials is LOH of chromosomes 1p and 16q. 17 Intensification of therapy for individuals with combined LOH 1p and 16q improves the outcome in all stages of FHWT. 17 Although highly specific for predicting relapse, 1p/16q LOH is present in only 4.6% of FHWTs and in only 9.4% of relapses. 18,19 More recently, gain of chromosome 1q has been associated with inferior event-free survival (EFS) and overall survival (OS) of individuals with WTs. 20,21 This was subsequently validated by the National Wilms Tumor Study-5 and the International Society of Paediatric Oncology (SIOP) WT 2001 Trial. Both studies identified 1q gain in 28% of individuals overall and demonstrated diminished EFS for individuals with 1q gain. 22,23 Upcoming protocols will determine whether modifying the initial therapy based on 1q gain will improve outcomes.
All of these studies have largely relied on randomly selected samples taken at the time of diagnosis, samples that may not contain the clonal events resulting in poor outcome. The current study seeks to determine whether examining relapse samples can provide further information regarding the pathogenesis, progression, and therapeutic responsiveness for individuals with FHWTs. These studies are possible because of the AREN03B2 umbrella biology and classification study that served as the entry portal to all Children's Oncology Group (COG) individuals registered on therapeutic protocols for renal tumors from 2006 through 2017; it now includes banked samples from over 6,000 individuals. The overall goal for the current study is to analyze samples from individuals registered as FHWT on AREN03B2 who relapsed as FHWT.

Clinical samples
Individuals with currently valid and verified consent who relapsed with FHWT and who had samples banked at the Biopathology Center (BPC) were considered eligible. To gain maximal information from as many samples as possible, independent discovery and validation sets were defined. Discovery set Relapse and germline samples from 51 unique individuals passed the quality control steps. Two individuals had samples from two different relapse episodes (PAWPUL and PATEIS). In 45 of 51 individuals, DNA was also available from the primary tu-mor sample that likewise passed quality control and therefore represent complete trios. RNA sequencing was performed on 49 of 51 relapse samples and 12 of 45 primary samples for which an adequate sample was available. Adequate samples were available for miRNA extraction for all 51 relapse samples and 12 paired primary samples. Validation set Independent of the discovery set, 31 additional individuals with relapse samples that passed DNA quality control were eligible for the validation set but not for the discovery set (STAR Methods). For the majority of these, the samples consisted of two unstained slides and an H&E slide.
Somatic variants in the discovery set Relapse samples Whole genomic sequencing (WGS) was performed on 53 relapse samples from 51 individuals. This resulted in 3,846 small variants, 301 of which passed the filtering criteria (STAR Methods). The details of all 301 variants are provided in Table S1. Sixteen genes were affected by these 301 somatic variants in more than one individual, and these are provided in Table 1; all were verified by RNA sequencing (RNA-seq) or Sanger sequencing. Nine of these 16 genes involved genes and variants previously identified and fully described in previous large studies of primary FHWTs. [9][10][11][12][13] In particular, the SIX1 177 Q/R hotspot mutation was identified in 7 of 51 individuals, and the MYCN 44 P/L hotspot mutation was identified in 5 of 51 individuals. The remaining 7 genes have not been reported previously to be mutated in WTs (MGA, TCF12, RBL1, HCFC1, MAPKBP1, COBLLI, and DIS3), each identified in two individuals.
Recurrent structural variants were also identified and are provided in Tables 1 and S2. These include tandem duplications of MYCN (7 individuals), deletions involving WT1 (2 individuals), and deletions including all or part of the WTX gene (7 individuals). WTX also contained a non-recurrent nonsense SNV mutation (Table 1). Primary samples WGS was also performed on 45 available paired primary samples, which resulted in 1,804 small variants, 249 of which passed the same filtering criteria applied to the relapse samples (STAR Methods). The details of all 249 are provided in Table S1; structural variants are provided in Table S2.
The tumor mutation burden (TMB) per megabase (assuming 25.8-Mb non-redundant coding regions in WGS) was calculated for the primary and relapse samples of all discovery set individuals. This demonstrated a low TMB for all tumors (ranging from 0.04-0.89 in the primary samples and 0.08-1.74 in the relapse samples), provided in Figure S1. Genes recurrently involved in more than three individuals in primary or relapse samples are illustrated in Figure 1, and those involved in more than two individuals are illustrated in Figure S1.
Germline mutations of the discovery set Peripheral blood samples (43) or normal kidney samples (8) of the 51 discovery set individuals were examined for small variants in genes recognized to be predisposing to WTs 24 and genes identified in individuals with WTs known to predispose to adult tumors. 25 These include BLM, BRCA2, BUB1B, CDC73,  CHEK2, CTR9, DICER1, DIS3L2, GPC3, GPC4, MUTYH, PALB2,  PIK3CA, PMS2, REST, TP53, TRIM37, and WT1. This germline predisposition analysis revealed a pathogenic variant in DICER1 in an individual who also had a different somatic DICER1 mutation in the primary tumor (epithelial predominant) and relapse (blastemal predominant) samples (PAUSLU). A likely pathogenic variant in CHEK2 (rs587782471, associated with predisposition to breast cancer) was also identified. Variants of unknown significance were identified in WT1 (1), BLM (2), REST (1), and TRIM28 (1); none demonstrated a reduction to homozygosity in the primary and relapse samples.
We also searched the germline for variants in the genes showing recurrent somatic variants listed in Table 1. Germline DIS3 variants of unknown significance were identified in 4 individuals (in addition to the two individuals with somatic DIS3 hotspot 488 D/N mutations). The rs141067458 stop-loss germline variant was identified in two individuals (PASYKN and PAVBXS); two individuals had the rs35288597 coding-change variant at amino acid 438 (PAUGMT and PAYTJD). Last, germline variants of unknown significance were detected in CHD4 (rs372219150) and RBL1 (rs761881234), each in one individual. Details of germline variants are provided in Table S2; all were identified in peripheral blood samples.
Copy number changes in the discovery set highlight 1q gain Segmental copy number analysis for the relapse sample and the paired primary tumor sample (when available) was computed from WGS data by the GDC. Regions reported previously as gained or lost in WTs [14][15][16] were evaluated and are provided in Table 2. These data confirm numerous gains and losses of entire chromosomes or chromosomal arms in WTs, particularly gain of 1q, 6, 7q, and 12 and loss of 1p, 16q, and 22. When comparing the copy number changes identified in the relapse sample with the primary sample (when available), the only copy number change that was significantly different was gain of 1q. In relapse samples, 38 of 51 (74.5%) demonstrated 1q gain compared with 21 of 45 (47%, p = 0.008, Fisher's exact test) of the available primary tumor samples. The rate identified in primary tumors (47%) and relapse tumors (74.5%) in the current study is also greater that the overall rate of 28% identified previously in all WTs. 22,23 The number of male and female individuals with 1q gain in the discovery set (15 of 21 females and 23 of 30 males) was not significantly different (p = 0.7499, Fisher's exact test). Comparing the copy-neutral LOH (CNLOH) or LOH for 1p or 16q within the relapse sample with the primary sample did not demonstrate significance Comparison of primary-relapse tumor pairs reveals late acquisition of some recurrent mutations Of the 51 discovery set individuals, 45 had available DNA from the primary tumor and were thus evaluable for comparison with the relapse tumor. In several individuals, a mutation was present in the relapse sample but not in the paired primary tumor. This discordancy was observed for SIX1 (3 of 6 evaluable individuals), MYCN (2 of 7 evaluable individuals), and WTX (3 of 8 evaluable individuals), illustrated in Figure 1 for genes with 3 or more mutations and in Figure S1 for genes with 2 or more mutations. To further verify the absence of the mutation in the primary sample, the unfiltered data of all evaluable discordant tumor sets were searched, and no variants were identified that were filtered out because of one of the criteria. For other genes, all evaluable tumor sets were concordant: WT1 (5), DROSHA (3), CTNNB1 (3), CHD4 (2), MLLT1 (2), and DGCR8 (1). All genes identified in relapsed WTs for the first time in this study (MGA, TCF12, RBL1, HCFC1, MAPKBP1, COBLII, and DIS3) were discordant in at least one individual. These observations indicate that there are 17 of 45 individuals whose primary tumor sample lacked a recurrent mutation from Table 1. This prompted a number of analyses of these samples with the aim of identifying underlying pathogenic variants in these 17 individuals. First, we identified variants that have been detected in FHWTs in prior studies that may have been filtered out as non-recurrent in this study. This identified a mutation in NONO that was observed in primary and relapse samples of PAVLIN. Second, we analyzed the raw variant calling format (VCF) files for recurrent mutations in non-coding regions. This resulted in identification of a promoter mutation in TERT (rs1242535815 G>A) in four individuals, including 3 of 51 individuals with relapse samples and 3 of 45 samples from primary tumors. (One relapse tumor with TERT mutation lacked a primary tumor, and the TERT mutation was present only in the primary tumor of one individual.) This mutation is a G>A change 124 bp upstream of the TERT transcription start site. The details of the TERT mutations are included in Tables 1 and S1. Retrospective analysis of the 56 FHWTs that underwent WGS in the Therapeutically Applicable Research to Generate Effective Treatments (TARGET) analysis revealed one tumor that carried this variant. 13 Gene expression of TERT in the 5 samples with promoter

. Recurrent alterations in WT discovery set individuals
Genetic alterations identified in at least 3 individuals within the primary (P) and relapse (R) tumors are illustrated in this OncoPrint. The numbers on the right provide the percentage of P or R samples that have alterations in the relevant gene. These data are also expanded in Figure S1, which provides all genes recurrently involved in each designated individual. Article ll OPEN ACCESS mutations who had RNA available was significantly higher than those lacking promoter mutations (p = 0.0067) ( Figure 2).

Individuals lacking mutations in their primary samples often have multiple copy number changes
The 16 remaining available primary tumors that lacked evidence of recurrent SNVs or structural variants were examined for copy number changes. We found that 11 of 16 demonstrated CNLOH of 11p15. In each case, the germline sample lacked 11p15 CNLOH. Excluding changes on 11p, the 16 samples had an average of 4.5 segmental copy number changes per tumor in the primary sample; only one individual lacked copy number changes (PAUWCD). Figure S1 illustrates the key mutations and copy number changes for each individual and provides the co-occurrence of those mutations and copy number changes.
The data in Figure S1 were also analyzed to identify tumors that had an identifiable stable clone (genetic changes present in primary and relapse samples) and evidence of clonal evolution (an additional change in the relapse sample). Of the 18 evaluable individuals, the changes most frequently identified in the stable clone Included 11p15 CNLOH (7), gain of 12 (8), and gain of 18 (5). The most frequent additional genetic change identified only in the relapse sample included 1q gain (11), gain of 12 (4), gain of 18 (4), and mutations in WTX (4), MYCN (2), and SIX1 (2). Also of interest was a paucity of copy number changes in individuals with WT1 and MLLT1 mutations.

Targeted sequencing of validation set
Mutations in the genes identified in Table 1 were evaluated in the independent 31-individual validation set using targeted  Table S4.
It should be noted that 6 of 35 variants (2 each involving COBLL1 and MGA, and one each involving RBL1 and HCFC1) are in the National Center for Biotechnology Information's Single Nucleotide Polymorphism Database (dbSNP) and have an allelic fraction of 50% and are therefore suspected to be rare germline variants. They are retained because of their predicted effect according to the sorting intolerant from tolerant (SIFT) and/or Polyphen tools for predicting whether an amino acid substitution will affect protein function. Such variants would not have been identified in the discovery set because of their presence in the paired germline DNA. The discovery set tumors were evaluated for these germline variants, and none were identified. In Table 1, the combined frequency of mutations in the discovery and validation sets was compared with the frequency identified in the TARGET study (using the appropriate total tumor number of 533 FHWTs in the TARGET validation set and 56 FHWTs in the TARGET discovery set that were analyzed by WGS for structural variants.) Only SIX1, identified in 11 of 82 (13.4%) individuals, showed a frequency of mutations significantly higher compared with the frequency seen in tumor samples in TARGET (23 of 533 [4%], p = 0.0026). The number of male and female individuals with SIX1 mutation in the combined discovery and validation sets (6 of 36 females and 5 of 46 males) was not significantly different (p = 0.5227, Fisher's exact test).

Pharmacogenomic analysis
Of the 226 unique pharmacogenomic polymorphisms analyzed (STAR Methods), four were excluded (one was not in gnomAD, and three had missing calls in more than 10% of samples). None of the remaining 222 polymorphisms were significantly different (adjusted p < 0.05, binomial test) in the entire sample set compared with the gnomAD general population. A separate analysis comparing all individuals of European descent (N = 35) with the non-Finnish European gnomAD population similarly yielded no significantly different polymorphisms. Other ethnicity-specific comparisons were not run because of low sample numbers: admixed American (AMR), 7; African (AFR), 6; Asian, 1; unknown, 2. Analysis for reduction to homozygosity performed in the paired relapse sample and (when available) the primary sample did not demonstrate a significant shift in the polymorphism frequencies in the tumor samples.

DISCUSSION
Studies comprehensively reporting the genomic analysis of WTs over the last decade have greatly increased our knowledge of the underlying genetic underpinnings of WTs. Despite this, identification of therapeutic targets has been limited. To sharpen our focus, this study analyzes WT relapse samples, which should contain clonal molecular features contributing to relapse.

SIX1 and MYCN mutations are more frequently identified in relapsed WTs
Prior studies have illustrated that mutations in many of the genes involved in renal development also play a key role in development of WTs, including SIX1, WT1, MYCN, WTX, MLLT1, and CHD4. 10,13 The current study emphasizes that some of the same genetic mutations are also increased in prevalence at relapse, particularly those whose role is preserving the progenitor state. In particular, the highly homologous SIX1 and SIX2 genes are required for maintaining the progenitor state; 26 the identical 177 Q/R hotspot mutations in SIX1 and SIX2 (so far specific to WTs) have been identified previously in about 5% of primary FHWTs. 10,12 Previous reports of structural analysis of the SIX1 Q177R mutations suggest an effect on the DNA binding site, likely altering DNA binding specificity. 10 This mutation is accompanied by up-regulation in cell cycle genes, 12 supporting an activating function. We demonstrate the same SIX1 hotspot mutation in 11 of 82 (13.4%) of relapse samples in the current study, a significantly higher frequency compared with the 4% Figure 2. TERT gene expression by mutation or copy number status RNA-seq data, normalized using variance-stabilizing transformation, was used to generate boxplots with the ggplot2 R package (v.3.6.3). The top, middle, and lower lines of the box represent the 25th, 50th, and 75th percentiles, respectively. The upper whisker extends to the largest value no greater than 1.5 times the interquartile range from the box and the lower whisker extends to the smallest value no lower than 1.5 times the interquartile range from the box.  13 It is noteworthy that SIX1 mutations were identified in the relapse sample but not in the primary sample in 3 of 6 of evaluable individuals, suggesting that it is not required for tumor development in many individuals. Supporting this is the report of increased allelic fraction of SIX1 mutations in relapse compared with primary samples in a study of 8 primary-relapse pairs. 27 An increased frequency of SIX mutations was also identified in posttherapy WTs that were blastemal predominant compared with other histologic subtypes, suggesting that SIX mutations may confer resistance to chemotherapeutic agents. 10 Prior studies point toward multiple functional roles of SIX1 that may contribute to its increased prevalence in relapsed individuals in our study; these include alteration of DNA binding specificity, up-regulation of cell cycle genes, and resistance to chemotherapy.
The MYCN network is also involved in preservation of the progenitor state in the kidneys. [28][29][30] The activating MYCN 44 P/L hotspot mutation and MYCN tandem duplications have together been reported previously in 15% of primary WTs in studies that report SNVs and structural variants; 10,13,31 these changes were identified in 24% of relapse samples in the current study. A recent study of over 800 unselected WTs demonstrated the MYCN 44 P/L mutation to be significantly associated with relapse. 32 We identified the hotspot mutation only in the relapse sample and not in the primary sample in 2 of 3 evaluable individuals, whereas all four evaluable individuals with MYCN tandem duplication in the relapse sample also had this mutation in the primary sample. Additional individuals in this study had variants in other members of the MYC transcription factor network that are expected to result in cellular effects similar to MYCN overexpression, including MAX (2 individuals), MGA (5 individuals), and NONO (1 individuals). MAX binds DNA as a heterodimer with MYCN or MYCC, and this MYC$MAX transcription activator is involved in all known oncogene functions of MYC. 33 MGA, likewise frequently mutated in cancer, binds to MAX and regulates target gene expression. 34,35 NONO, an RNA-binding protein, binds to MYCN, leading to post-transcriptional up-regulation of MYCN mRNA and protein expression. 36 In total, the relapse samples of 25 of 82 individuals (30%) included in this study had evidence of mutations involving the MYCN network. Studies suggest that the mechanisms underlying the increased relapse rate in individuals with activation of N-MYC may be linked to interacting partners, including PEG10, YEATS2, FOXK1, CBLL1, and MCRS1, all of which correlate positively with MYCN expression in WTs. 32 FOXK1 in particular is known to regulate cancer initiation, development, angiogenesis, and drug resistance. 37,38 Knockdown of YEATS2 in lung cancer cells results in growth suppression and reduced survival, 39 all of which are key MYC functions. The interaction of MYC-N with YEATS2 may therefore contribute to oncogenesis by supporting cell growth and survival. Although all MYCN variants reported here were somatic events, germline MYCN duplication has been identified in a family predisposed to WTs. 40 Recurrent DIS3 germline and somatic mutations and TERT promoter mutations We identified recurrent somatic mutations in two functionally important genes that have not been recognized previously in WTs: DIS3 and TERT.

DIS3 mutations
Mutations of miRNA processing genes are an important category of mutations in WTs; these result in global reduction of mature miRNAs, but in particular let-7a. 11,12 Decreased let-7a may also result from up-regulation of LIN28B, which specifically binds pri/pre-let-7 miRNAs, [41][42][43] triggering their degradation by DIS3L2, 44 an exoribonuclease that is also rarely mutated in WTs. 9,10,13 Rare germline mutations in DIS3L2 result in Perlman syndrome, associated with increased risk of WTs. 45 A paralog of DIS3L2 is DIS3, a protein not reported previously to be mutated in WT but recognized as a recurrent mutation resulting in multiple myeloma. 46 A DIS3 somatic mutation (488 D/N) involving the catalytic domain of ribonuclease II (RNB domain) was identified in two individuals; this has been reported previously in individuals with multiple myeloma. 47 Two different germline variants of unknown significance were also identified in two individuals each. In particular, the rs141067458 stop-loss germline variant has been reported to result in lower DIS3 expression and to be associated with familial multiple myeloma, 48 although its role in WT development is unknown. Dis3 knockdown in Drosophila severely disrupts development of wing imaginal discs by regulating a small subset of microRNAs, in particular miR-252 and miR-982, 49 miRNAs with no known human orthologs.
TERT promoter mutations The current study also provides the first report of TERT promoter mutations in WTs. Telomerase maintains telomere length, thereby maintaining self-renewal potential. Somatic mutations in the promoter of TERT, the catalytic subunit of telomerase, have been reported in several tumor types and are predicted to increase promoter activity and TERT transcription 2-to 6-fold (reviewed by Ackerman and Fischer 50 ). These promoter mutations were associated with high expression of TERT in the current study ( Figure 2). Analysis of TERT expression in 78 FHWTs demonstrated a significant association between mRNA expression of TERT and relapse in univariate and multivariant analyses. 51 This significant association was verified in a subsequent study of 244 NWTS-5 individuals (96 relapse, 148 without relapse). 52 Additional genes recurrently mutated in the relapse samples of small numbers of WTs have likewise not been reported previously, although their significance remains unclear. These include mutations in (1) the basic helix-loop-helix (bHLH) binding domain of TCF12, reported previously to be associated with an aggressive tumor phenotype in anaplastic oligodendroglioma; 53 (2) HCFC1, whose loss results in proliferation of neural progenitor cells at the expense of differentiation; 54 (3) RBL1, a member of the retinoblastoma tumor suppressor family that modulates E2F transcription factor activity; 55 (4) MAPKBP1, one of over 20 genes linked to development of nephronophthisis; and (5) COBLL1, a gene associated with age-related macular degeneration.

Gain of 1q is highly prevalent in WT relapse
Observations highlighted by the current study, but certainly documented previously by others, 14 Article ll effort over the last decade. The most striking finding of this study is the prevalence of 1q gain in the relapse samples of WTs (75%) compared with the primary samples (47%) and compared with the overall prevalence of 1q gain reported previously in primary WTs (28%). The increased prevalence of 1q gain in individuals with increased stage and increased age, 22 and the increased allelic fraction seen in smaller studies of relapsed individuals, 27 strengthens the growing consensus that 1q gain is often associated with progression and solidifies its role in guiding therapy.
This study relies on retrospective analysis of prospectively obtained tumor samples from individuals registered on COG studies, which have historically relied on a single randomly selected tumor sample. This practice enables collection of the highest quality of sample (a fresh tumor collected shortly after surgery). However, this leaves large areas of a tumor unsampled and does not allow selection based on histology. Concern regarding the effect of tumor heterogeneity has therefore grown. To address this, Cresswell et al. 6 collected 70 tumor samples from 24 tumors in 20 individuals and demonstrated striking heterogeneity in their ability to detect 1q gain in multi-sampled tumors. In fact, had they only collected a single sample per tumor, 1q gain would have been detected in only about a third of the cases in which it was present. They estimated that at least three samples per tumor were needed to ensure that more than 95% of tumors with 1q gain would be detected. Sampling bias resulting from reliance on a single random tumor sample is the largest limitation of the current study. To correct this sampling bias in clinical trials is remarkably difficult for a number of practical reasons. Simply taking three samples from each tumor at the time of nephrectomy will not address all situations, particularly those relying on initial biopsy. To address this concern, efforts have recently focused on detecting circulating tumor DNA (ctDNA), and the possibilities and pitfalls have been discussed in the setting of pediatric cancer. 56,57 In a recent study of ctDNA in 50 individuals with high-stage WTs, only individuals with detectable ctDNA experienced relapse or died from disease. 58 Although the presence of 1q gain in the tumor predicted its presence in the serum, a number of individuals showed 1q gain in the serum but lacked 1q gain in the randomly selected tumor sample. 58 This supports the concept that measuring ctDNA at the time of diagnosis in individuals with WTs may enable detection of clonal 1q gain anywhere in the entire tumor burden. Studies examining the clinical utility of ctDNA have been included in the next therapeutic protocols for WTs. Copy number changes of other chromosomes, such as chromosomes 12 and 18, within the tumor may also be independently useful in predicting relapse and may augment our understanding of the development of WTs. These data will be easily captured because microarrays will be utilized in the next COG protocols to comprehensively evaluate copy number change in the tumor.
The power of this study is that it represents a comprehensive analysis of the largest number of relapse samples of WTs reported to date. In addition, the availability of primary and normal tissue from many individuals enables us to gain some insight into the temporal acquisition of mutations in WTs. This observation adds to an accumulating set of evidence that suggests that genetic variants may play important roles throughout tumor evolution, although there is not yet evidence to support a particular sequence of genetic events. In fact, the combinations of mutations or structural changes may be critical, rather than the temporal order of their accumulation. In particular, the co-occurrence of mutations in genes supporting continued progenitor proliferation with those preventing differentiation may be most important. Examples include SIX with DROSHA and WT1 with CTNNB1. 13 Limitations of the study The limitations of this study reside in the reliance on a single random sample at each episode (reviewed above) combined with the reliance on the relatively low coverage provided by WGS, precluding assessment of clonal evolution in this study. Another limitation of this study is the relatively small number of cases analyzed, given the nature of the tools applied, which generally require a large number of samples to achieve confidence. This limits the type of conclusions that can be confidently drawn in this study, particularly for pharmacogenomic variants. It also limits our ability to provide biologic verification using the RNA and miRNA expression patterns.

STAR+METHODS
Detailed methods are provided in the online version of this paper and include the following:  Article ll funded in whole or in part with federal funds from the National Cancer Institute, National Institutes of Health, under contract HHSN261201500003I. The content of this publication does not necessarily reflect the views or policies of the Department of Health and Human Services, nor does mention of trade names, commercial products, or organizations imply endorsement by the US government. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. The authors thank the Children's Oncology Group (COG) protocol coordinators, research coordinators, clinical research assistants, and other health professionals who contributed to acquiring samples used in this study. The authors in particular wish to thank the extended AREN03B2 team, who contributed to the central review and clinical annotation of these samples, as well as Jeff Dome, chair of the COG renal tumor disease committee during acquisition of some of the samples utilized in this study. We also acknowledge the excellent efforts of project managers from the BPC (Yvonne Moyer and Kae Tegtmeier), as well as support from Kristina Tracy from the Genomics Platform of the Broad Institute of MIT and Harvard. Most importantly, we are very grateful to the individuals and their families for consenting to sample deposition through AREN03B2 to the BPC.

Materials availability
This study did not generate new unique reagents.
Data and code availability d Sequencing data, de-identified human clinical information, and sample quality control information have been deposited on the publication page of this study at the Genome Data Commons (https://portal.gdc.cancer.gov/projects/MP2PRT-WT) GDC and are publicly available as of the date of publication. Accession numbers are listed in the key resources table. d This paper does not report original code. d Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request

METHOD DETAILS
Eligibility for this study included the following d Patients registered on AREN03B2 as FHWT who relapsed with FHWT (per central review) following therapy, with a current valid verified consent. (The development of contralateral disease following chemotherapy was not considered relapse for this study.) d Patients received both vincristine and actinomycin following diagnosis. (Patients who relapsed after surveillance alone or single agent vincristine were not included). d Patients have relapse sample banked by the Biopathology Center (BPC).
Of patients meeting the above criteria, several had limited availability of a germline or normal tissue comparator sample, or of a relapse sample. To maximally utilize relapse samples, a discovery and a validation set were defined. For the discovery set, the following additional criteria were included: 1) sufficient relapse sample banked at the BPC to perform whole genomic sequencing, and 2) a source of germline DNA (normal kidney or peripheral blood) banked by the BPC. Available samples from the primary tumor prior to therapy were not required for the discovery set, but when available were also included in the analysis. The validation set included all patients that met the first set of criteria but failed one or more of the second set of criteria.
Specimen processing and quality control Quality control was performed on each tumor specimen from either a frozen section prepared by the BPC or from a permanent section taken from a formalin-fixed, paraffin-embedded (FFPE) block. Hematoxylin and eosin (H&E) stained sections were reviewed to confirm that the tumor specimen was histologically consistent with FHWT. Percent tumor nuclei, percent necrosis, and other pathology annotations were assessed. Tumor samples with R60% tumor nuclei and %20% necrosis and normal tissue samples with 0% tumor nuclei were submitted for nucleic acid extraction at the BPC.
RNA and DNA were extracted from tissue using a modification of the AllPrep DNA/RNA kit (Qiagen). The flow-through from the Qiagen DNA column was processed using a mirVana miRNA Isolation Kit (Invitrogen) for frozen tissue and High Pure miRNA Isolation Kit (Roche) for FFPE samples. DNA was extracted from blood using the QIAamp DNA Blood Midi Kit (Qiagen). RNA samples were quantified by measuring Abs260 with a UV spectrophotometer and DNA was quantified by Quant-iT PicoGreen Assay Kit using the FilterMax F5 Multi Mode Microplate Reader. DNA specimens were resolved by 1% agarose gel electrophoresis to confirm high molecular weight fragments. A custom SNP panel (using Complex iPLEX Gold Genotyping Reagent from Agena and Extend primary mix from Integrated DNA Technologies) verified that tumor DNA and germline DNA representing a case were derived from the same patient. RNA was analyzed via the RNA6000 Nano assay (Agilent) on the Agilent Bioanalyzer for determinations of an RNA Integrity Number (RIN) for frozen tissue and the DV 200 values for FFPE samples. Cases yielding 1.0 mg of tumor DNA from FFPE, 1.2 mg of tumor DNA from frozen tissue, 2.0 mg RNA, and 1.0 mg of germline DNA were preferred in this study; samples with lower yields were also included if they passed all other quality control steps.
The BPC processed tumor samples from a total of 115 cases, of which 85 cases qualified, 11 requiring macrodissection. All qualified cases were sent for genomic analysis. Of the 30 cases that were disqualified, 19 cases failed pathology, 2 cases were too small to extract, 1 case did not have a germline sample available, and 8 did not meet molecular quality metrics. Genomic DNA (350 ng in 50mL) was used as the input into DNA fragmentation with acoustic shearing performed using a Covaris focused-ultrasonicator, targeting 385 bp fragments. Following fragmentation, a clean-up step was performed using Ampure XP SPRI beads. Library preparation was performed using a commercially available kit (KAPA Biosystems Hyper Prep without amplification module), and with palindromic forked adapters with unique 8-base index sequences embedded within the adapter (Unique Dual Indexed Adapter Kits, Roche). Following sample preparation, libraries were quantified using quantitative PCR (KAPA Biosystems Quantitative PCR kit), with probes specific to the ends of the adapters using a ViiA7 qPCR machine, and automated using Agilent's Bravo liquid handling platform. Based on qPCR quantification, libraries were normalized to 2.2 nM and pooled into 24-plexes. The pools were loaded onto NovaSeq 6000 S4 flowcells using the Hamilton Starlet liquid handling system, to produce 151 bp paired-end reads and each sample was sequenced to a coverage of 30x.
Output from Illumina software was processed by the Picard data-processing pipeline to yield CRAM files containing demultiplexed, aggregated aligned reads and submitted to the Genomics Data Commons (GDC). GDC identified somatic DNA variants for each primary and relapse tumor sample using the paired normal sample. Single nucleotide variants (SNVs) were called using CaVEMan, 59 indels using Pindel, 60 structural variants using BRASS, 61 and copy number variation using ASCATngs, 62 (Bioinformatics Pipeline: DNA-Seq Analysis -GDC Docs (cancer.gov). The following files were generated: aligned harmonized BAM files, raw SNVs in VCF format, raw indels in VCF format, raw structural variants in VCF and browser extensible data paired-end (BEDPE) format, gene-level copy number data in TSV format, and genomic segmented copy number in TXT format (https://portal.gdc.cancer.gov/projects/ MP2PRT-WTGDC). Discovery set RNA sequencing RNA sequencing was performed by the University of North Carolina. Fresh frozen RNA analytes were assayed for RNA integrity, concentration, and fragment size. Samples for total RNA-seq and small RNA-sequencing were quantified on a TapeStation system (Agilent, Inc. Santa Clara, CA). RNA Integrity score (RIN) averaged 8.1. Samples with RINs >8.0 were considered high quality. Input concentrations greater than 100 ng/ul were ideal and the amount of material ranged between 0.85 and 2.52 ug of RNA. Initial fragment size determined if additional fragmentation was needed.
For total RNA-sequencing, library construction was performed using the Stranded Total RNA Prep with RiboZero Gold protocol (Illumina) and Truseq RNA UD indexes (IDT for Illumina) following the manufacturer's instructions. Libraries were prepared on an Agilent Bravo Automated Liquid Handling System. Quality control was performed at every step and the libraries were quantified using a TapeStation system. Indexed libraries were prepared and run on HiSeq4000 paired end 75 base pairs to generate a minimum of 150 million reads per sample library with a target of greater than 90% mapped reads. Typically, these were pools of three to four samples. The raw Illumina sequence data were demultiplexed and converted to fastq files with bcl2fastq v.2.20.0, and adapter and low-quality sequences were removed.
For miRNA-sequencing, miRNA-seq library construction used the NEXTflex Small RNA-Seq Kit (v3, PerkinElmer, Waltham, MA). Samples were bar-coded with individual tags following the manufacturer's instructions. Libraries were prepared on Sciclone Liquid Handling Workstation Quality control was performed at every step, and the libraries were quantified using a TapeStation system and an Agilent Bioanalyzer using the Nextflex Small RNA analysis kit (Perkin Elmer). Pooled libraries were then size selected according to NEXTflex Kit specifications using a Pippin Prep system (Sage Science, Beverly, MA). Typical pool size was 20 libraries. Indexed libraries were loaded on the Hiseq4000 to generate a minimum of 10 million reads per library with a minimum of 90% reads mapped. The raw Illumina sequence data were demultiplexed and converted to FASTQ files using bcl2fastq v.2.20.0. Resultant data were analyzed using a variant of the small RNA quantification pipeline developed for TCGA. 66 Samples were assessed for the number of miRNAs called, species diversity, and total abundance. Samples passing quality control were uploaded to the GDC repository where the files were processed using the following pipeline: Bioinformatics Pipeline: mRNA Analysis -GDC Docs (cancer.gov).

VALIDATION SET TARGETED SEQUENCING
Targeted sequencing was performed at the Broad Institute of MIT and Harvard. For covering variants within a gene, the exons plus 10 bp at exon-intron boundaries were sequenced and DNA variants were called using the GATK Mutect2 tumor-only calling pipeline 67 and were filtered as described for the discovery set. Hotspot mutations underwent direct sequencing. Exon 3 of CTNNB1 was sequenced. Structural variants resulting in small copy number gains or losses (WTX and MYCN) were addressed by sequencing MYCN exon 2 and WTX exon 2, which were involved in all the segmental copy number changes identified in both the current study and in the TARGET discovery sets. 13 For WT1, all exons were sequenced. The location of all the primer targets analyzed are provided in Table S4.
For library construction, genomic DNA (50-200 ng in 50mL) was used as the input into DNA fragmentation performed acoustically using a Covaris focused-ultrasonicator, targeting 150 bp fragments. Library preparation was performed using a commercially available kit (KAPA HyperPrep Kit with Library Amplification) and IDT's duplex UDI-UMI adapters. Unique 8-base dual index sequences embedded within the p5 and p7 primers (IDT) were added during PCR. Enzymatic clean-ups were performed using AMPureXP SPRI beads with elution volumes reduced to 30mL to maximize library concentration. Library quantification was performed using the Invitrogen Quant-It broad range dsDNA quantification assay kit with a 1:200 PicoGreen dilution. Following quantification, each library was normalized to a concentration of 35 ng/mL, using Tris-HCl, 10 mM, pH 8.0.
After library construction, hybridization and capture were performed using the relevant components of IDT's XGen hybridization and wash kit and following the manufacturer's suggested protocol, with several exceptions. A set of 12-plex pre-hybridization pools were created. These pre-hybridization pools were created by equivolume pooling of the normalized libraries, Human Cot-1 (from the XGen hybridization and wash kit), and blocking oligos (xGen Universal blockers, IDT). The pre-hybridization pools underwent lyophilization using the Biotage SPE-DRY. Post lyophilization, the custom target bait (https://www.twistbioscience.com) along with hybridization mastermix was added to the lyophilized pool prior to resuspension. Library normalization and hybridization setup were performed on a Hamilton Starlet liquid handling platform, while target capture was performed on the Agilent Bravo automated platform. Post capture, PCR was performed to amplify the capture material using a mastermix containing HiFi HotStart Ready Mix (Kapa Biosystems), and dual index forward and reverse primers (IDT). Library pools were then quantified using qPCR (Quantitative PCR kit, KAPA Biosystems) on a ViiA7 qPCR machine. Based on qPCR quantification, pools were normalized using a Hamilton Starlet to 2 nM. The pools were loaded onto lanes of the HiSeq X sequencer to produce 151 bp paired-end reads and samples sequenced to a coverage goal of 500x MTC. Output from Illumina software was processed by the Picard data-processing pipeline to yield BAM files containing demultiplexed, aggregated aligned reads. Samples passing quality control were uploaded to the GDC repository (GDChttps://portal.gdc.cancer.gov/projects/MP2PRT-WT).

Somatic variant analysis
For analysis, the raw unannotated simple somatic mutation pindel files (for small indels) and CaVEMan VCF files (for SNVs) were obtained from the GDC and filtered to include regions associated with gene exons plus 10 bp (UCSC canonical gene coordinates, GRCh38.d1.vd1) and annotated as ''PASS''. The filtered variants were annotated using the ENSEMBL Variant Effect Predictor (VEP) tool (Variant Effect Predictor -Homo_sapiens -Ensembl genome browser 105). 68 Variants were removed that were indicated by the ENSEMBL VEP to be 1) present outside of the coding region or canonical splice site, or 2) synonymous, or 3) present in the general population with an allelic frequency (AF) > 0.01, or 4) had a dbSNP ID without a COSMIC ID and predicted to be both tolerated by SIFT and benign or possibly damaging by Polyphen. The total read count, alternate allele count, and alternate allele fraction were calculated from the VCF annotation. The tumor mutation burden was calculated as the number of nonsynonymous exonic mutations per 25.8 Mb non-redundant coding regions. 69,70 Analysis for variants discordant between primary and relapse samples was performed using the samtools mpileup method 71 to extract the BAM reads at the region of the discordant variant.
For analysis of structural variants, raw BRASS structural variants were obtained from the GDC in BEDPE format. Structural variants without an assembly score or an assembly score <90 or with <4 reads supporting the variant were removed. Recurrent tandem duplications, deletions, inversions and translocations within the relapse samples were identified. Genetic alterations for key genes were visualized using Oncoprinter from the CBioPortal for Cancer Genomics. 72,73 Somatic copy number analysis Gene-level and segmented genomic copy number files from WGS data were obtained from the GDC (https://portal.gdc.cancer.gov/ projects/MP2PRT-WT) GDC. Loci of interest based on prior large studies of copy number changes relevant to WT [14][15][16] were evaluated by filtering the segmented copy number files to include segments R8 MB. Specific gene copy number variations were provided in the gene-level copy number files from the GDC. Copy number was classified as gain, loss, no gain, no loss, LOH, copy-neutral LOH.

Germline analysis
The germline (normal kidney or peripheral blood) CRAM files were processed by the DRAGEN v3 germline pipeline (DRAGEN Germline (illumina.com)) using default settings. The resulting GVCF files were filtered to include 1) 15 genes classified as Wilms tumor predisposition genes (BLM, BRCA2, BUB1B, CDC73, CTR9, DICER1, DIS3L2, GPC3, GPC4, PALB2, PIK3CA, REST, TP53, TRIM37, WT1, 24 2) three additional genes with germline mutations identified in patients with WT that have been associated with predisposition to adult tumors 25 (CHEK2, MUTYH, PMS2) and 3) the variants contained in the genes identified in Table 1. These variants were annotated using Ensembl VEP, and filtered in the same manner as described for somatic variants. The clinical impact was designated in the categories of benign, likely benign, variant of unknown significance, and pathogenic or likely pathogenic according to American College of Medical Genetics and Genomics and the Association for Molecular Pathology germline variant classification guidelines. 74 Those variants evaluated as benign or likely benign were filtered out. The general population allelic frequencies provided by ENSEMBL were verified in gnomAD gnomAD (broadinstitute.org); 75 variants not present in gnomAD are labeled ''Novel''.