Landscape of somatic single-nucleotide and copy-number mutations in uterine serous carcinoma

Exome Sequencing of USC.

Fifty-seven patients with uterine serous carcinoma were studied. Their clinical features are presented in SI Appendix, Table S1. Upon surgical removal of tumors, primary cell lines were prepared (15 tumors) or tumors were frozen (42 tumors). Exome sequencing was performed on all tumors; for 34 of these, DNA samples from normal tissue were available and sequenced. Exome sequencing was performed using the NimbleGen/Roche capture reagent followed by 74 base paired-end DNA sequencing on the Illumina HiSeq platform (6). By design, tumor samples were sequenced to greater depth of coverage to permit detection of somatic mutations in tumors despite admixture of normal and tumor cells in these samples. For tumors and normal DNA, each targeted base was sequenced by a mean of 187 and 100 independent reads, respectively (SI Appendix, Table S2). Of all targeted bases in tumors, 94.5% were read by 20 or more independent reads; mean per-base per read error rates were 0.42% for normal DNA and 0.48% for tumor DNA. Segments of loss of heterozygosity (LOH) were called from the difference in B-allele frequency between tumor-normal pairs (SI Appendix, Fig. S1), allowing estimates of tumor purity, which were above 60% for frozen tumors and higher for primary cell lines. Somatic mutations were identified by finding variant reads in tumors that were significantly more frequent than expected by chance (Materials and Methods). At the coverage levels studied, there was no significant relationship between tumor purity and the number of somatic variants detected, consistent with sufficient depth of coverage having been achieved to identify the vast majority of somatic mutations. Variants in genes implicated in the pathogenesis of USC were verified by direct Sanger sequencing and were found to be expressed in all available USC cell lines.

Tumors with Hypermutator Phenotype.

The number of protein-altering somatic mutations per tumor markedly deviated from a normal distribution (Fig. 1A). In the discovery set of 34 USC with matched normal DNA, 30 tumors had fewer than 100 protein-altering somatic mutations (median 36), whereas 4 had more than 3,000 somatic mutations each. Only one of these tumors was from a cell line (with limited propagation), and none came from patients who had received chemotherapy before sample acquisition. These tumors with high mutation burden were also notable for having no LOH segments or copy-number variants (CNVs), a feature found in only five other tumors. These features suggest a hypermutator phenotype due to deficiency of mismatch repair (MMR) or polymerase ε (POLE) genes (7, 8). Consistent with this, these hypermutated tumors showed a paucity of T:A > A:T or C:G > A:T transversions (SI Appendix, Fig. S2) (9). Examination of the POLE and MMR genes showed no germ-line mutations; however, somatic mutations in these genes were highly prevalent in these tumors (mean of 4 per tumor, including 4 premature termination mutations for MMR genes and a mean of 4.5 per tumor for POLE) and more frequent than expected by chance (P = 2.23 × 10⁻³) (SI Appendix, Table S3). Among the cancers without matched normal DNA, one showed a similarly high prevalence of rare protein-altering variants (>3,000) and a skewed distribution of rare protein-altering transversions. Thus, 9% of USC in this cohort have a hypermutator phenotype. Because of the skewing effect of the large number of mutations in these tumors, they were not included in subsequent analyses of mutation burden.

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Fig. 1.

Somatic variation pattern underlying USC. (A) Distribution of the number of protein-altering somatic mutations in 34 normal-tumor USC pairs. Subplot left, mutation spectrum in four hypermutator phenotype samples; subplot right, mutation spectrum in 30 moderately mutated samples. (B and C) Thirty tumors with moderate somatic burden are arranged by the total number of somatic point mutations from left to right. The four hypermutator phenotype tumors are excluded in this analysis. (B) Significantly mutated genes are listed vertically by the order of damaging or conserved P values shown in Table 1. (C) Genes with significant CNVs and genes of interest are listed. Copy neutral status is shown as lavender rectangles. Five samples without CNV information are marked by crosses.

Analysis of Single-Nucleotide Variants.

Among somatic mutations in the 30 remaining matched tumors, we identified recurrences of somatic mutations at the same positions. Accounting for the rate of protein-altering somatic mutations in these tumors (1.1 × 10⁻⁶) and the size of the exome, the likelihood of seeing the mutation twice by chance at any position among these tumors is <10⁻³. We identified six genes with recurrent somatic mutations (Table 1). These included well-established activating mutations in PIK3CA (10), the catalytic subunit of phosphoinositide-3 kinase (five tumors); the well-established G12V mutation in KRAS (three tumors) (11); and a mutation at R465 in FBXW7 in four tumors (12). FBXW7 is the targeting component of a SCF-type 3 ubiquitin ligase, and R465 occurs in the WD40 domain involved in substrate recognition; mutation at this site prevents targeting of cyclin E for ubiquitination and degradation (SI Appendix, Fig. S3) (12, 13). Recurrent mutations also occurred at two sites in PPP2R1A, the constant regulatory subunit of serine-threonine phosphatase 2a. The P179R and S256F mutations occurred four and two times, respectively, and have been previously reported (14). These mutations occur at the interface where PPP2R1A interacts with regulatory B subunits that target the phosphatase to specific substrates; inhibition of this interaction by SV40 small t antigen plays a role in viral transformation (SI Appendix, Fig. S4). Additional somatic mutations were found on the surface of PPP2R1A that interacts with the B or C (catalytic) subunit (Table 1). TP53, the well-characterized tumor suppressor gene, had five different positions mutated two or more times, and there were 19 additional single somatic mutations in this gene. Eighty-two percent of these mutations were in segments of somatic LOH (Table 1 and Fig. 1B).

In addition to these previously described recurrent mutations, a recurrent mutation was found in CHD4/Mi2b (chromodomain-helicase–DNA-binding protein 4), an ATP-dependent chromatin-remodeling protein that is a major subunit of the Mi2b/nucleosome remodeling and deacetylase (NuRD) complex. Mutations in CHD4/Mi2b have not been previously associated with cancer. In addition, there were 10 other somatic or rare mutations in CHD4 among matched and unmatched tumors (see further discussion below).

We next sought genes with overall increased somatic mutation burden in the 30 matched tumor-normal pairs. In this analysis, we determined the probability of seeing ≥n mutations in each gene, taking into account the overall rate of protein-altering somatic mutations in the matched tumor normal set (1.1 × 10⁻⁶) and the length of the protein-coding region in each gene. We also adjusted for the level of expression of each gene from expression data in normal human endometrium (15) because we found a higher somatic mutation rate among genes with lower expression, consistent with effects of transcription-coupled DNA repair reducing the mutation rate among expressed genes (16). We considered P values < 2.4 × 10⁻⁶ to represent a significant increase in mutation burden compared with that expected under the null hypothesis, accounting for the testing of ∼21,000 genes. We added to this set variants in the 22 unmatched tumors that occurred in genes that had at least one somatic mutation in the matched set and that had never been seen in >7,000 exomes in the Yale University and National Heart, Lung, and Blood Institute exome databases. Because we found no novel variants in any of these genes in the 30 germ-line samples of tumor-normal pairs, we infer that virtually all of these represent somatic mutations.

In the resulting set, the six genes with recurrent mutations were among the most frequently mutated genes. Included in this set was CHD4, which had six somatic mutations and five more novel variants found in the 22 unmatched tumors (Table 1 and Fig. 1B). Many of these 11 CHD4 mutations (Fig. 2), which all appear to be heterozygous, impair at least some normal CHD4 functions. CHD4 is a SWI2/SNF2 ATPase and part of the larger helicase superfamily 2 whose members share a similar catalytic core containing two RecA-like helicase domains. Conserved catalytic “signature” motifs have been well described and contain many residues required for catalysis of ATP hydrolysis and helicase activity (20). Three CHD4 mutations (R957Q, R1127G, and R1162W) alter residues in these signature motifs (motif B, V, and VI, respectively) that are conserved from yeast to humans, and whose mutation has been shown to impair normal function. Similarly, there is a mutation in the second plant homeodomain (PHD) finger that normally binds methylated histone H3K9. This C464Y mutation disrupts one of the key cysteines that coordinate Zn²⁺ binding. In addition, there are four mutations (Q1106R, I1109T, and two instances of F1112L) clustered in a short α-helix in ATPase lobe 2; we speculate that alteration of this helix might alter interaction with another protein in the complex. Additional mutations include two in a C-terminal bridge that links ATPase lobes 1 and 2 at positions that appear to stabilize this segment. Finally, there is a premature termination near the normal C terminus (Fig. 2). The high prevalence of CHD4 mutations, the clear implication of disrupted normal function by many CHD4 mutations, and the common genomic amplifications of the CHD4 gene (see CNV results below) implicate CHD4 mutations in USC.

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Fig. 2.

Mapping of USC mutations onto the crystal structure of CHD4. (A) Schematic representation of somatic mutations found in CHD4. The horizontal bar represents full-length CHD4 protein with functional domains shown as boxes. Somatic mutations found in USC are marked in red text. All mutations are missense mutations except E1628X, which is a nonsense mutation. (B) C464 locates in the second PHD finger, which binds directly to histone H3 methylated at K9. An NMR structure of the second PHD domain of CHD4 (blue ribbon plot) has been determined [Protein Data Bank (PDB) ID: 2L75] (17). Red dot represents C464Y mutation. (Lower) A close-up view of the zinc–C464 interaction. (C) Somatic mutations in catalytic core of CHD4 mapped to the crystal structure of a related protein, human CHD1 (sequence identity of the ATPase lobes is 42%, homology is 57% over 572 residues; PDB ID: 3MWY) (18). Blue, ATPase lobe 1; light purple, ATPase lobe 2; cyan, chromodomains; green, C-terminal bridge. Three mutations (red dots) in CHD4 fall within known conserved motifs (motifs B, V, VI) (19); three mutations were found in unknown helicase motif i, and two mutations were found in unknown motif ii. (D) A close-up view of mutations in five motifs in C. Somatic mutations are labeled in red text; amino acid positions in parentheses represent homologous positions in CHD1.

Another gene of interest was TAF1, an X-linked gene, which had four different somatic mutations and three additional variants in unmatched tumors (Table 1). TAF1 is the largest component and core scaffold of the TFIID basal transcription factor complex and has DNA-binding activity, histone acetyltransferase (HAT) activity, two kinase domains, and ubiquitin-activating/conjugating activity (21). It is known to be required for progression through the G1 phase of the cell cycle, promoting cyclin D expression (22). Most of the seven TAF1 mutations lie in the HAT domain at positions that are extraordinarily well conserved—all are conserved in vertebrates and nearly all are conserved in yeasts (Fig. 3). Although the function(s) of these mutations in USC are uncertain, overexpression of TAF1 has been previously reported in human lung and breast carcinoma and found to be associated with poor tumor differentiation and high mitotic activity (25).

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Fig. 3.

Schematic representation of TAF1 functional domains and mutation conservation analysis. (A) Functional domains in TAF1 are represented by colored boxes with domain names noted below (23). HAT domain, histone acetyltransferase domain. Mutations found in USC are labeled at the top by red text. (B) Multiple sequence alignment across vertebrate and invertebrate species around the seven mutations found in USC. Mutation positions in human TAF1 are labeled in red at the top. Sequence aligned by Clustal W 2.0 (24).

Additional genes that meet thresholds for significantly increased burden in the entire set include PTEN, CDKN1A, and SPOP, as well as HCFC1R1, CTDSPL, YIPF3, and FAM132A, genes not previously implicated in cancer. For each of these genes, mutations are predominantly at highly conserved positions, there are few if any silent mutations in the same gene, and quantitative PCR demonstrated expression of each of these genes in all available USC cell lines.

Analysis of CNVs.

We next assessed somatic CNVs. For the 25 tumors in which read coverage distribution showed distinct modes (SI Appendix, Fig. S5), CNVs were identified by comparing coverage depth of individual capture intervals from tumor and normal samples (Materials and Methods); CNVs were supported by significant deviation of the B allele frequency from the genome-wide average. The significance of CNVs affecting specific chromosome segments was assessed by Monte Carlo simulation, randomly distributing CNVs of the empirically observed sizes and numbers in each tumor in 10⁸ permutations to assess the distribution expected by chance alone. A significance threshold was established that provided a false discovery rate <0.25. Within each significant copy-number gain or loss, all CNVs that contained the most frequently altered segment were removed, and the remaining CNVs were reassessed to see if independent signals could be detected. We identified 13 chromosome segments with more frequent gains of copy number and 12 with more frequent deletions than expected by chance (Fig. 4). Among these, we found focal amplification of the segment of chromosome 17 that contains ERBB2 in 11 of the 25 tumors (44%) (SI Appendix, Fig. S6), large duplications that include the PIK3CA locus in 60%, and a small duplication of chromosome 19 containing CCNE1 in 48% (SI Appendix, Table S4). There was also amplification of a large segment of chromosome 8 containing MYC in 11 (44%) tumors and amplification of a segment of chromosome 12 that included CHD4 in 7 (28%) tumors (Fig. 1C). Among deletions, TP53 was deleted in 44% of tumors. The most frequent somatic deletions were small (0.5 Mb) deletions on chromosomes 19 and 22, which occurred in 68% and 72% of tumors, respectively (SI Appendix, Table S5 and Fig. S7). Most interestingly, the chromosome 19 interval contains MBD3, which is a component of the same SWI/SNF complex as CHD4 (19). The chromosome 22 interval includes a number of interesting genes, including three in the MAP kinase pathway, HDAC10, and PPP6R2.

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Fig. 4.

Copy-number profile of 25 USC tumors. Frequency of copy-number gain (red) and copy-number loss (blue) are plotted along the genome. Horizontal dotted line, genome-wide significance level for CNV gain (red) and CNV loss (blue). Genes of interest in significant CNV peak regions are labeled.