Prostate cancer represents a substantial clinical challenge because it is difficult to predict outcome and advanced disease is often fatal. We sequenced the whole genomes of 112 primary and metastatic prostate cancer samples. From joint analysis of these cancers with those from previous studies (930 cancers in total), we found evidence for 22 previously unidentified putative driver genes harboring coding mutations, as well as evidence for NEAT1 and FOXA1 acting as drivers through noncoding mutations. Through the temporal dissection of aberrations, we identified driver mutations specifically associated with steps in the progression of prostate cancer, establishing, for example, loss of CHD1 and BRCA2 as early events in cancer development of ETS fusion-negative cancers. Computational chemogenomic (canSAR) analysis of prostate cancer mutations identified 11 targets of approved drugs, 7 targets of investigational drugs, and 62 targets of compounds that may be active and should be considered candidates for future clinical trials.
Prostate cancer is the most common solid cancer in men (diagnosed in 12%) and is often fatal (9% of male cancer deaths). It is difficult to manage clinically due to a poor current understanding of what dictates its highly variable natural history and of what underlies the development of castration-resistant disease1. Extensive data on the structure of prostate cancer genomes have been published2,3,4,5,6, including work from our own consortium7,8,9,10. These studies have identified a number of genetically distinct subgroups, including cancers with ERG, ETV1, ETV4, FLI1, SPOP, FOXA1 and IDH1 alterations. Overlapping with these categories, cancers may have alterations in PI3K and DNA repair pathways, with the latter significantly over-represented in advanced disease4. However, we have relatively limited understanding of the ordering of genetic events, with the exception that ETS gene alteration appears to represent an early event while mutations of AR are later, sometimes convergent events occurring in advanced and metastatic disease. Indeed, we have very little understanding of the evolution of mutational processes, the various genetic paths that cancers traverse on their way to progression, the levels of heterogeneity at different stages of development, or the effect of these factors on clinical outcome.
Gene status has been used in studies designed to improve the poor predictive value of conventional clinical markers (PSA, Gleason sum, stage) and to develop disease management strategies. For example, genetic alteration of BRCA1 or BRCA211, PTEN deletion12, amplification of AURKA together with the MYCN gene13, and coordinated loss of MAP3K7 and CHD114 have been reported to have prognostic value. A number of commercial prognostic tests based on gene expression profiles are also available15,16,17, and a classification framework has been proposed18. Improvements in the treatment of castration-resistant disease have been made through better targeting of AR regulation using abiraterone19 and enzalutamide20, while PARP inhibitors are effective against cancers harboring BRCA1 or BRCA2 mutations and other defects in DNA repair pathways21. However, notable advances have been made recently through the retasking of approved drugs22. In this study, we use previously unpublished whole-genome DNA sequencing data in combination with published data to provide insights into the mechanism of progression of prostate cancer to lethal disease and to design new molecularly based strategies for drug targeting.
We whole genome sequenced cancerous and matched normal samples from 87 primary prostate cancers from the UK and 5 from China, together with 10 hormone-naive prostate metastases and 10 castration-resistant metastases from the USA. Our analysis (see Methods) yields insights into the nature and order of acquisition of driver alterations, genomic heterogeneity in primary and metastatic cancers, changes in mutational signatures during progression and potential drug targets. In addition, we identify coding and noncoding drivers by combining single nucleotide variants (SNVs) and small insertions/deletions (indels) in our dataset with those from The Cancer Genome Atlas (TCGA)4 (425 samples), the COSMIC database23 (243 samples) and Stand Up to Cancer24 (SU2C-PCF, 150 samples) to give a combined dataset, hereafter referred to as the ‘joint dataset’, comprising 710 primary cancers and 220 metastases. Supplementary Table 1 summarizes the genes affected in both our study and the joint dataset.
For the 112 cancer–normal pairs in our cohort, we identified 392,753 SNVs, 54,952 indels and 10,921 chromosomal rearrangements (Fig. 1). The mean genome-wide substitution rate was 1.23/Mb, with a significant difference in mutational burden between the primary (0.99) and metastatic (2.30) samples (P = 4.4 × 10−15, Methods). Moreover, within the metastatic subset, mutation burden was higher in men treated with androgen deprivation therapy (ADT, also referred to as castration resistant prostate cancer, CRPC) than treatment-naive men (2.98 vs. 1.61, P = 0.015). There were also significantly more rearrangements in metastatic than in primary samples (P = 0.0059), while the proportion of breakpoints attributed to a chromoplexy-like event25 was indistinguishable between the two groups. Within the metastatic group, the ADT samples had more rearrangements than did the hormone-naive (P = 0.027), with no difference in the proportion of chromoplexy-like events (Fig. 1).
Genes of interest were determined through a comprehensive set of analyses to identify excess non-synonymous mutations in coding regions; excess missense mutations within a gene, indicative of an oncogenic driver; excess mutations in noncoding regions; regions with an excess of structural variants in either ETS+ or ETS– cancers; and regions with recurrent copy number aberrations (CNAs) in either ETS+ or ETS– cancers. Overall, we identified 73 genes with evidence for involvement in prostate cancer development (Fig. 2, Table 1 and Supplementary Table 2). Based on a literature search, each gene was assigned a high, medium or low level of previous supporting evidence (Table 1 and Supplementary Table 2). In addition to 22 genes with little or no previous evidence of involvement in prostate cancer (Table 1, low previous evidence), we provide corroborating evidence for 8 more genes previously lacking strong evidence of driving prostate cancer (Table 1, medium previous evidence).
Coding driver mutations
We identified 28 genes with an excess of non-synonymous coding mutations, 5 of which are previously unknown drivers in prostate cancer (Supplementary Table 2). TBL1XR1 was enriched in truncating SNVs and indels and is also located in a genomic region enriched for rearrangements in ETS+ cancers (chr3:172–179Mb) (Fig. 3). These rearrangements result in loss of heterozygosity (LOH) or, in one case, homozygous deletion, suggesting a cancer suppressor role for this gene. Another significantly mutated gene primarily affected by truncating mutations was ZMYM3, which encodes a component of CoREST, a transcriptional repressor complex including REST (RE-1 silencing transcription factor) and involved in suppression of neuronal differentiation-related genes in non-nervous tissues26. In addition, two further CRPC samples from the SU2C-PCF study24 had nonsense mutations and one sample in our study had a 70-kb exonic deletion in REST.
Two other genes with recurrent truncating mutations were IL6ST and CASZ1 (Fig. 3). The latter is a putative cancer suppressor in neuroblastoma27, while the former encodes glycoprotein 130, the signal-transducing subunit of the interleukin 6 (IL6) receptor. The pattern of mutations we observe in the joint dataset for IL6ST is dominated by truncating events. Moreover, this gene is located in a genomic region recurrently rearranged in ETS+ cancers, resulting in either LOH or homozygous deletion (four cases of each), suggesting a cancer-suppressive role. TBX3, previously reported to harbor mutations in breast cancer28, exhibited a mixed pattern of mutations with mostly missense mutations and two cancers harboring truncating events.
Analysis of missense mutations identified recurrent mutations in seven more genes, of which two are newly reported (Supplementary Table 2). CNOT3 exhibited mutation hotspots altering two amino acid positions, Glu20Lys (4 of 932 samples) and Glu70Lys (5 of 932 samples), as well as a nonsense mutation in a single sample (Fig. 3). CNOT3 has a known cancer-suppressive function in T-cell acute lymphoblastic leukemia29. Enrichment for missense mutations was identified in RPL11, which encodes a ribosomal protein and putative cancer suppressor upstream of the MDM2–TP53 pathway30. In contrast to findings of previous studies, the enrichment for missense mutations in both CNOT3 and RPL11 suggests oncogenic, rather than tumor suppressor, roles in prostate cancer. A comparison between coding mutations in metastatic and primary samples in the joint dataset identified enrichment in metastases for mutations in TP53, AR, KMT2C, KMT2D, RB1, APC, BRCA2, CDK12, ZFHX3, CTNNB1 and PIK3CB (Supplementary Table 2), confirming previous studies3, 24.
Noncoding driver mutations
Analysis of noncoding portions of the genome identified two regions significantly enriched for mutations. NEAT1, which produces a long noncoding RNA recently reported to be associated with prostate cancer progression31, was mutated in 13 of 112 ICGC samples, with significant over-representation in patients with metastatic disease (6 of 20 metastases vs. 7 of 91 primaries, Fisher’s exact test, P = 0.012; Fig. 3). Out of the metastatic tumors, NEAT1 mutations were found only in patients who had undergone ADT, consistent with the link between high NEAT1 expression and resistance to AR-targeting therapies31. Notably, two of these six samples had two separate NEAT1 mutations. The FOXA1 promoter also had significant evidence of selection. This gene modulates AR-regulated transcriptional signaling32 and has previously been found to harbor recurrent coding mutations5. In our series, we identified 14 samples with coding and 6 samples with noncoding mutations, with two samples (PD14721a and PD12813a) bearing both a coding and a noncoding mutation. We also identified mutations in the promoter for FOXP1, a gene with known cancer-suppressive effect in prostate tumorigenesis33, in three samples, but this was insufficient to reach statistical significance.
Structural variant enrichment in ETS+ and ETS– cancers
The density of rearrangements varies across the genome as a result of various factors, including chromatin state, GC content, gene density, replication timing and repetitive sequence. To remove the effect of these factors, we segmented interbreakpoint distance across the genome separately in ETS+ and ETS– cancers and identified regions with differential enrichment for rearrangements between the two subtypes. The functional importance of many of these regions was supported by an excess of truncating mutations or CNAs.
In addition to regions previously identified as enriched for rearrangements in ETS+ cancers (FOXP1, RYBP, SHQ1, PTEN and TP53)34,35,36,37, we identified two unreported regions. The region chr5:55–59Mb covers the genes PPAP2A, PDE4D, MAP3K1 and IL6ST (Fig. 3). In IL6ST, we also detected significant enrichment for coding mutations, suggesting this is the main target of the aberrations. At chr3:171–178Mb, TBL1XR1 was similarly enriched for both rearrangements and truncating mutations.
In ETS– cancers, we confirmed a previously reported enrichment for rearrangements containing CHD138, 39. A target of enriched rearrangements in the region chr1:149–158Mb is likely ETV3. In 5 of 9 cancers, ETV3 was exclusively affected by these events (4 LOH by deletion and 1 by translocation). Additionally, 1 cancer had a truncating mutation (Arg413fs*3) and 2 had missense mutations (Ala73Val and Leu37Gln). In total, 12 patients had localized alteration, 10 of whom had ETS– cancers. Moreover, in the joint dataset, there were four cancer samples with truncating mutations in this gene. In contrast to ETV4, the nature of variants in ETV3 is indicative of a tumor-suppressive role in prostate cancer. Manual inspection of the recurrently rearranged region chr3:76–84Mb identified ROBO1 and ROBO2 as possible targets (Fig. 3). In total, 16 of 112 samples had an event affecting one or other of these genes, and in 4 samples both were affected. Previously implicated in pancreatic ductal adenocarcinoma40, these two genes have not been previously reported in the context of prostate cancer.
Events enriched at chr6:80–114Mb indicate that ZNF292 is a possible target. Eleven of 112 patients (5 ETS+ and 6 ETS–) had loss of at least one chromosome copy, and in 2 patients there was a homozygous loss specifically targeting ZNF292. Moreover, the joint dataset contained 5 of 932 samples with a truncating mutation, further suggesting a cancer-suppressive function for this gene in prostate cancer. Another gene affected by recurrent rearrangements on 6q was SENP6, which encodes a small ubiquitin-like modifier (SUMO)-specific protease that removes SUMO polypeptides from conjugated proteins41 and may influence AR function42. Of note, 4 of 5 rearrangements in this region affected SENP6 only, leading to a significant reduction in expression; P < 0.0001 (Supplementary Fig. 1). Finally, located at chr6:126Mb, the nuclear receptor co-activator NCOA7 was altered in 6 samples, with 1 sample having homozygous loss.
Further regions enriched for rearrangements in ETS– cancers were chr2:133–144Mb (LRP1B), chr8:112–114Mb (CSMD3) and chr8:40–41Mb (MYST3). The first two genes are very large and fall within reported fragile sites43. Nevertheless, preferential enrichment of breakpoints in ETS– cancers may suggest either that underlying structure, such as AR binding sites or nucleosome structure, or that epistatic interactions between ETS fusion and other rearrangements affect the occurrence of rearrangements at these loci. Samples containing structural variants affecting MYST3 were found to have significantly reduced RNA expression; P = 0.025 (Supplementary Fig. 1).
Timing of copy number aberrations
To identify routes to progression in prostate cancer, we developed an approach by which to order the occurrence of CNAs by combining information on the clonality of CNAs, timing relative to whole-genome duplication (WGD), and timing of homozygous deletions relative to neighboring hemizygous losses. Information from all tumors was combined using a Bradley–Terry model to give the most likely ordering of events. By applying a set of logical rules (see Methods), we deciphered the temporal ordering of subclonal CNAs within each cancer. In general, homozygous deletions appeared late in oncogenesis, corroborating previous findings that homozygous deletions are associated with advanced disease44,45,46.
Clear differences emerged in the evolution of ETS+ and ETS– prostate cancers. Where present, the deletion between the TMPRSS2 and ERG genes in ETS+ cancers was an early (generally clonal) event, as was gain of chr8q in the locus 112–137Mb (Fig. 4a). The earliest homozygous deletions in ETS+ cancers included chr5:55–59Mb, corroborating the rearrangements targeting PPAP2A, PDE4D, MAP3K1 and IL6ST, and chr10:89–90Mb, which covers PTEN (Figs. 3 and 4a). In ETS– cancers, losses at chr5:60–100Mb (CHD1 and RGMB), chr13:32–91Mb (which includes BRCA2, RB1 and FOXO1) and chr6:73–120Mb are followed by losses at chr2:124–142Mb, then by gains at chr3:100–187Mb, and then whole chromosome gain of chr7 (Fig. 4b). Loss of CHD1 has been previously implicated in the initiation of ETS– prostate cancers, preventing ERG rearrangement in the prostate38, and our data confirm the exclusivity between ETS positivity and homozygous loss of CHD1 (Fig. 4c). In both ETS+ and ETS– cancers, WGD correlated with loss of chromosomal segments at chr1:94Mb, chr2:140Mb, chr12:12Mb, chr16:85Mb and chr17:7Mb (Fig. 4c). From timing analysis, these losses appear to occur synchronously with WGD in most cases. Gains at chr8:101Mb occurred before WGD, gains at chr3:131Mb occurred synchronously, and gains at chr7:88Mb tended to follow WGD.
Timing of point mutations and indels
SNVs and indels were clustered according to their cancer cell fraction using a Bayesian Dirichlet process47. The proportion of SNVs identified as subclonal showed considerable variation across cancers, but was significantly higher in primary than metastatic samples (P = 0.022, Wilcoxon rank sum test, Fig. 5a), as was the proportion of subclonal indels (P = 0.00033) and the fraction of the genome with subclonal CNAs (P = 0.0037, Supplementary Fig. 2). This is apparent evidence for a bottleneck in acquiring metastatic potential rather than a response to treatment, since levels of heterogeneity in untreated metastases are no lower than in androgen-deprived metastases (Fig. 5a).
The levels of heterogeneity observed in SNVs and indels were correlated (Pearson r = 0.57, P = 2.3 × 10−9, Fig. 5a). Higher levels of heterogeneity were observed among indels than SNVs (P = 2.4 × 10−8). However, the possibility cannot be ruled out that variant calling of indels may have greater sensitivity for low-allele-frequency variants than calling of SNVs.
Driver SNVs were identified as clonal or subclonal in each sample according to the cluster to which they were assigned, with 84 classified as clonal and 22 (21%) as subclonal. Our power to detect subclonal mutations is limited by sequencing depth, and the real number of subclonal driver mutations is likely much higher. The driver mutations identified as subclonal included 2 mutations in APC in the same sample, PD14713a. Notably, this cancer has undergone clonal loss of one copy of chr5q, followed by mutations in APC in two different subclones (Fig. 5b and Supplementary Fig. 3), suggesting convergent evolution. Five other samples each had two subclonal drivers: PD12808a has a missense mutation in ZNF292 and an essential splice site mutation in SMAD2; PD13401a has a nonsense mutation in PPP1R3A and a mutation in the promoter of NEAT1; PD13402a has a nonsense mutation in USP34 and an essential splice site mutation in ABI3BP (Fig. 5b); PD12820a has a missense mutation in USP48 and an essential splice site mutation in ASXL2; and PD13389a has a frameshift mutation in PHF12 and an essential splice site mutation in TBX3 (not shown). Subclonal mutations were also seen in several common drivers, including 1 in TP53 (PD13339a) and 1 in PTEN (PD12840a). By contrast, SPOP was mutated in 10 samples, always clonally and always in ETS– tumors (Fig. 2).
Analysis of the mutational signatures by non-negative matrix factorization (NMF) showed that, in addition to the ubiquitous ‘clock-like’ signatures 1 and 5, there was presence of the previously described48 signatures 2, 3, 8, 13 and 18. Signature-3-positive samples were enriched for germline or somatic mutations in BRCA1 and BRCA2 genes (4 of 6 samples), as reported previously48 (Fig. 1). However, the presence of high levels of microhomology-mediated deletions was even more strongly correlated with the presence of BRCA mutations (6 of 6 samples). Separating the mutations into early clonal, late clonal and subclonal epochs (see Methods) showed that the proportion of signature 1 mutations decreases over time, suggesting an increase of cancer-associated mutagenic processes relative to innate processes (P = 2.2 × 10−16, test for trend in proportions).
Signature 13, previously associated with the activity of the AID/APOBEC family of cytidine deaminases, was over-represented in advanced disease: 45% (9 of 20) in metastases vs. 14% (14 of 92) in primaries (Fisher’s exact test, P = 5.6 × 10−3). Similarly, signature 18, which has been previously associated with failure of base excision repair and the accumulation of mutations from 8-oxoguanine damage49, was enriched in advanced disease: 40% (8 of 20) in metastases vs. 11% (10 of 92) in primaries (Fisher’s exact test, P = 3.8 × 10−3). In a recent report of 560 breast cancer whole genomes, signature 8 correlated with DNA damage repair deficiency50. Androgen signaling is known to positively regulate multiple genes involved in DNA repair51, 52, while androgen deprivation impairs DNA double-strand break repair53. In support of these previous reports, the proportion of mutations assigned to signature 8 is consistently higher among later appearing (subclonal) populations of cells (55% ± 24%) than earlier (clonal) populations (28% ± 12%) (t-test, P = 1.3 × 10−4, Supplementary Table 3). The proportion of metastases with evidence for the action of signature 8 was higher than that for primary tumors, although not reaching statistical significance (8 of 20 metastases vs. 25 of 92 primaries, Fisher’s exact test P = 0.28). Increased prevalence of DNA-damage related genes in metastatic prostate cancer, as well as the observations made in this study, warrant an extensive study of mutational signatures in therapy-naive disease and CRPC in a larger dataset to explore the relevance of checkpoint inhibition as an alternative therapy for advanced prostate cancer.
CDH12 and ANTXR2 alterations were significantly associated with time to biochemical recurrence (Benjamini–Hochberg adjusted P = 0.0060 (CDH12) and 0.012 (ANTXR2), HR (hazard ratio) = 9.3 and 7.7, Cox regression model, Fig. 6) and were significant predictors of biochemical recurrence independent of cofactors Gleason, PSA at prostatectomy, and pathological T-stage (P = 0.00061 (CDH12) and 0.0015 (ANTXR2), HR = 7.3 and 6.5, respectively; Cox regression model, Supplementary Table 4). A Cox regression model containing a combination of CDH12, ANTXR2, SPOP, IL6ST, DLC1 and MTUS1 mutations was determined to be an optimal predictor of time to biochemical recurrence and was a significant improvement over a baseline model of Gleason, PSA at prostatectomy, and pathological T-stage (model χ2 test, P = 0.00053). The number of mutational signatures identified in a cancer was negatively correlated with time to biochemical recurrence in prostatectomy patients (P = 0.014, HR = 3.0; Cox proportional hazards model on number of processes greater than 3, Supplementary Fig. 4) and was an independent predictor (P = 0.0061, HR = 3.6; Cox proportional hazards model). The number of SNVs detected was also an independent prognostic biomarker (P = 0.031, HR = 1.005; Cox proportional hazards model). The numbers of both samples and events in this study are modest, and further analysis of larger cohorts is required to establish firmly these findings.
Druggable targets in the prostate cancer disease network
A key opportunity arising from systematic analyses of cancer genomics is the early identification of therapeutic intervention strategies. To this end, we applied established chemogenomic technologies using the canSAR knowledge base54 to map and pharmacologically annotate the cellular network of the prostate disease genes identified in this study. We derived the network using curated protein–protein and transcriptional interaction data. We included the protein products of the genes identified in this study and other key proteins that directly interact with these proteins or affect their function (see Methods and Supplementary Fig. 5). This resulted in a focused prostate network of 156 proteins. Each protein was annotated according to multiple assessments of ‘druggability’; i.e., the likelihood of the protein being amenable to small molecule drug intervention (Table 2 and Supplementary Table 5). We found that prostate cancer driver genes were embedded in a highly druggable cellular network that contains 11 targets of approved therapies and 7 targets of investigational drugs. As well as the AR and the glucocorticoid receptor (GR), the network contains targets of drugs approved for other indications, several of which (for example, BRAF, ESR1, RARA, RXRA, HDAC3) are under clinical investigation for prostate cancer.
Seven proteins in the prostate network are targets of drugs now in clinical trials. In particular, the ataxia-telangiectasia mutated (ATM) inhibitor AZD-0156, currently in phase 1 for safety assessment, is a likely candidate for exploration in prostate cancer owing to the recently described role of DNA damage repair, particularly in advanced prostate cancers21, 55. The network highlights targets of phosphatidylinositol-3-OH kinase pathway inhibitors (PI3K, AKT1) that are undergoing clinical investigation in prostate cancer, as well as IDH1 and MDM2 drug targets.
To give an indication of the potential of these drugs, we analyzed the most recent drug sensitivity data (GDSC56, see URLs). Eighteen drugs acting on our network were tested in GDSC on prostate cancer cell lines. Of these, 5 showed significant effect on growth inhibition and the remaining 13 drugs showed weak activity in at least one cell line (Supplementary Table 6). However, to validate fully the potential of these drugs, extensive drug sensitivity testing needs to be performed in disease-relevant cancer models that correctly reflect the patient population.
Potential future opportunities for prostate cancer therapy are also highlighted by 13 proteins that are under active chemical biology or drug discovery investigation (Table 2). These include menin (MEN1), a component of the MLL–SET1 histone methyltransferase complex. Mice with Men1 alterations develop prostate cancer57, and recent data have shown that menin expression is involved in CRPC58. A further 49 proteins are predicted to be druggable and therefore potentially amenable to drug discovery. These include the known prostate cancer protein SPOP, the transcription activator BRG1 (SMARCA4), CDK12 and the CREB binding protein CREBBP.
In summary, we find that 80 of the 156 proteins central to the prostate disease network are either targets of existing drugs or have the potential to be targeted in the future. To maintain an up-to-date view of this analysis, we provide a link to a live page in canSAR (see URLs).
The analysis of whole-genome sequence data from 112 prostate cancers has highlighted many of the genetic factors underlying the processes of carcinogenesis, progression, metastasis and the acquisition of drug resistance. Supporting evidence has been provided for 30 candidate driver genes with limited or no previous support, including the noncoding drivers NEAT1 and FOXA1.
Through the timing of genomic aberrations, we have a picture of the possible routes to progression in prostate cancer. Most driver mutations may occur either clonally or subclonally, but mutations in SPOP and ETS fusions occur early in cancer development and are exclusively clonal. Whereas the gain of 8q and ETS fusion appear to be sufficient to drive a dominant clonal expansion, ETS– cancers typically need a combination of large-scale losses, acquired over an extended period of time. Known cancer drivers are frequently observed subclonally, and two competing drivers are seen in several cancers. Metastases have less genomic heterogeneity, likely resulting from a bottleneck in achieving metastatic potential.
We observe changes in the mutational processes operative upon cancers during progression. Signature 8 was enriched in subclonal expansions, and signatures 13 and 18 were enriched in metastatic cancers. Cancers with germline or somatic BRCA1 or BRCA2 mutations were enriched for signature 3, demonstrating the effect of double-strand repair defects throughout cancer evolution.
Losses of CDH12 and ANTXR2 result in poorer recurrence-free survival. We identify 69 prostate cancer-associated proteins that are either targets for existing drugs or new potential targets for therapeutic development.
Analysis of the whole-genome sequences of over 100 prostate cancers has started to reveal the complex evolutionary pathways of these cancers. The early acquisition of driver aberrations, including ETS fusions and whole genome duplications, strongly affects the acquisition of subsequent aberrations. Acquisition of individual mutations affects both the subsequent likelihood of metastasis and response to treatment. Network analyses identified, in addition to previously known drivers, targets that could be exploited for clinical investigation with existing drugs as well as targets for new drug discovery, giving potential for the results of genome analysis to be translated rapidly into therapeutic innovation and patient benefit.
Patient cohorts, samples and ethics
We have complied with all relevant ethical regulations. We collected 92 cancer samples from prostatectomy patients treated at The Royal Marsden NHS Foundation Trust, London, at the Addenbrooke’s Hospital, Cambridge, at Oxford University Hospitals NHS Trust and at Changhai Hospital, Shanghai, China as described previously8, 68. Clinical details for the patients are shown in Supplementary Table 7. Ethical approval was obtained from the respective local ethics committees and from The Trent Multicentre Research Ethics Committee. All patients were consented to ICGC standards (see URLs). Twenty men from PELICAN (Project to ELIminate lethal CANcer)69, an integrated clinical-molecular autopsy study of metastatic prostate cancer, were subjects in the current study. Subjects consented to participate in the Johns Hopkins Medicine IRB-approved study between 1995 and 2005 (Supplementary Table 7). Subject A17 had a germline BRCA1 mutation, as previously reported70.
DNA preparation and DNA sequencing
DNA from whole blood samples and frozen tissue was extracted and quantified using a dsDNA assay (UK-Quant-iT PicoGreen dsDNA Assay Kit for DNA) following the manufacturer’s instructions with a fluorescence microplate reader (Biotek SynergyHT, Biotek). Acceptable DNA had a concentration of at least 50 ng/μl in TE (10 mM Tris, 1 mM EDTA) and an OD260/280 between 1.8 and 2.0. WGS was performed at Illumina, Inc. (Illumina Sequencing Facility, San Diego, CA, USA) or the BGI (Beijing Genome Institute, Hong Kong), as described previously, to a target depth of 50× for the cancer samples and 30× for matched controls8. The Burrows-Wheeler Aligner was used to align the sequencing data to the GRCh37 reference human genome71.
Variant calling pipeline
SNVs, insertions and deletions were detected using the Cancer Genome Project Wellcome Trust Sanger Institute pipeline as described previously8. In brief, SNVs were detected using CaVEMan with a cut-off ‘somatic’ probability of 95%. Postprocessing filters were applied. Insertions and deletions were called using a modified version of Pindel72. Variant allele frequencies of all indels were corrected by local realignment of unmapped reads against the mutant sequence. Structural variants were detected using Brass8. A positive ETS status was assigned if a breakpoint between ERG, ETV1 or ETV4 and previously reported partner DNA sequences was detected.
Mutation burdens were compared between primary and metastatic samples and between ADT and hormone-naive samples using a negative binomial generalized linear model (GLM), implemented with the R package MASS. Sample type was found to be an independent predictor of number of SNVs, as was age at time of sampling.
Timing of copy number events
We developed an approach to order the occurrence of CNAs by combining three sources of information: (i) clonality of CNAs; (ii) timing relative to whole-genome duplication, and; (iii) timing of homozygous deletions relative to neighboring hemizygous losses. Information from all tumors was combined using a Bradley–Terry model to give the most likely ordering of events during progression of prostate cancer.
The Battenberg algorithm was used to detect clonal and subclonal somatic CNAs and to estimate ploidy and cancer content from the next-generation sequencing data as previously described73. Briefly, germline heterozygous SNPs were phased using IMPUTE2 and a- and b-alleles were assigned. Data were segmented using piecewise constant fitting74 and subclonal copy-number segments were identified via a t-test as those with b-allele frequencies that differed significantly from the values expected of a clonal copy number state. Ploidy and cancer purity were estimated with the same method used by ASCAT75.
In this cohort, we defined WGD samples as those that had an average ploidy greater than 3. For tumors that had not undergone WGD, gains were defined as those regions that had at least one allele with copy number greater than 1, while losses were defined as those segments that undergone LOH. For tumors that had undergone WGD, losses were called in those segments with at least one allele with copy number of less than 2, whereas gains were called for those with an allelic copy number greater than 2. An extension of this logic was used for subclonal copy number segments; the evolving cellular fraction was always defined as that which deviated away from overall ploidy (defined as 2 for non-WGD samples and 4 for WGD samples). For example, if 75% of cells in a non-WGD tumor have a copy number of 3 + 1 at a given genomic loci, with the remaining 25% of cells having a copy number of 2 + 1, then we assume there has been clonal gain to 2 + 1 and then a subclone containing 75% of cells has undergone a further gain.
Three independent approaches were used to extract evolutionary data from each cancer sample. The first involved ordering clustered subclonal cancer fractions, the second used implicit ordering of clonal homozygous deletions in relation to losses, and the third estimated the relative timing of whole genome duplication. The logical arguments used in each approach are considered in turn. First, Battenberg algorithm-derived estimates for the cellular fraction and s.d. of each subclonal aberration were input to a Markov chain Monte Carlo hierarchical Bayesian Dirichlet process to group linked events together in an unsupervised manner. This defined clusters of different cell populations, each present at a calculated cancer cell fraction. The pigeonhole principle was then used to determine the hierarchical relationship between these clusters. Using this process, gains, losses and homozygous deletions were ordered with the following caveat to ensure that only independent events are ordered: if there was a clonal and subclonal gain (or loss) at the same locus, then only the clonal or initial gain (or loss) was ordered. Second, homozygous deletions have implicitly occurred after loss of heterozygosity at the same locus. Finally, the parsimony principle was used to define the allele counts that correspond to early and late changes in relation to WGD. For losses, if the minor allele copy number equals 0, then the loss occurred before WGD. Otherwise the loss occurred after WGD. Regarding gains, if the major allele copy number is twice or greater than ploidy, then the gain occurred before WGD. Otherwise, the gain occurred after WGD.
The above arguments allow us to gain insights into the order of copy number events in each individual tumor sample. To establish a consensus order across a cohort of tumor samples requires the ordering data to be integrated across all samples. As specific copy number events (location of breakpoints and the individual copy number states) tend to be unique to individual samples, we defined reference copy number segments that occurred recurrently. These were then used to build an overall contingency table.
The reference genomic segments were defined as regions that were recurrently aberrant. Regions of significant recurrence (false detection rate (FDR), P < 0.05) were determined by performing 100,000 simulations, placing the CNAs detected from each sample in random locations within the genome. The process was repeated for gains, LOH and homozygous deletions and the randomly generated copy number landscape compared to that arising from this cohort provided significance levels. Each significantly aberrant region was initially segmented using all breakpoints from all the events that contributed to that region. For instance, the significantly enriched region for LOH:chr8:0–44Mb contains over 300 breakpoints drawn from all the samples that contain LOH at chromosome 8p. We computed significantly recurrent regions and reference segments for both ETS+ and ETS– sample subgroups.
Performing pair-wise comparisons between all segmented results using the Bradley–Terry method described below proved computationally expensive, and therefore the total number of segments used in the pairwise comparison was rationalized by grouping reference segments to make combined segments of minimum length 1 Mb.
We then considered each tumor sample in turn. If any copy number event overlapped the reference genomic segments and was ordered in relation to any other event (that also overlapped regions of significance), those overlapped reference segments were ordered in comparison to other overlapped reference segments. In addition to these reference segments, the TMPRSS2–ERG deletion was ordered more stringently by considering only those segments that could result in the gene fusion, and not merely overlap the locus. In this manner, a contingency table of contests was constructed using reference genomic segments as the variables. We built contingency tables for both ETS+ and ETS– tumor samples to determine whether their evolutionary trajectory differed significantly.
An implementation of the Bradley–Terry model of pairwise comparison in R76 with bias reduced maximum likelihood estimated the ability or overall order of each individual reference segment.
The fraction of each cancer genome with subclonal CNAs was calculated as the total amount of the genome with subclonal CNA, as identified by the Battenberg algorithm, divided by the total amount of the genome that had CNAs. One sample (PD13397a, Supplementary Table 8) was identified as having very low cellularity, as it had a completely flat copy number profile and only 411 identified SNVs. Since CNAs could not called in this sample, it was not possible to adjust allele frequencies to cancer cell fractions (CCFs), and this sample was excluded from subclonality analysis. SNVs and indels were separately clustered using a Bayesian Dirichlet process, as previously described47. Clonal variants are expected to cluster at a CCF close to 1.0. However, in 18 tumors (Supplementary Table 8), there was no cluster in the range 0.95–1.05. The likely cause of a shift in CCF is inaccuracy in copy number calling, and these samples therefore failed quality control and were excluded from subclonality analysis. From Markov chain Monte Carlo sampling carried out in the Dirichlet process model, the posterior probability of each variant having a CCF below 0.95 was estimated. Variants with a probability above 80% were designated as ‘subclonal’, those with probability below 20% were designated ‘clonal’, and those with intermediate probabilities were designated as ‘uncertain’. The fraction of subclonal variants used in Fig. 5 and Supplementary Fig. 2 was then calculated after excluding uncertain variants.
The mutational spectra, defined by the triplets of nucleotides around each mutation of each sample, were deconvoluted into mutational processes as previously described48, 77. Clonal and subclonal variants were separated as defined above. Further separation of clonal mutations was performed for mutations in genomic regions that had undergone copy number gains. These mutations were classified as ‘early’ or ‘late’ depending whether their observed allele frequencies were more likely to indicate their presence on 2 or 1 chromosome copies, respectively, as assessed by binomial probability. Assignment of mutations to mutational signatures was carried out on each subset of mutations (early, late, clonal, subclonal), as well as on all mutations from each sample (Supplementary Table 3).
Clinical survival analyses
A Cox regression model was fitted to 71 features: every gene with mutations (breakpoints, substitutions or indels) with a potential functional impact (missense, nonsense, start-lost, in-frame, frameshift, or occurred in a noncoding transcript) or a CNA highlighted by the CNA analysis that occurred in three of more prostatectomy patients. The endpoint was biochemical recurrence. P-values were adjusted for multiple testing using the Benjamini–Hochberg method. Multivariate analyses were performed on all genes found to be significant using discretized Gleason (6, 7, 8 or 9), pathological T-stage (T2, T3) and PSA at prostatectomy as cofactors. Gene selection for the optimal predictor of time to biochemical recurrence was determined using Lasso78, a shrinkage and selection method for linear regression, starting with all genes that had a significant association with time to biochemical recurrence. Standard algorithms were used for survival analyses and statistical associations.
Identifying new oncogenes
The joint dataset was compiled from the aggregation of variants called in our samples with three other datasets, yielding a total of 930 samples, comprising 710 primary and 220 metastatic samples: (i) TCGA4, comprising 425 primary cancer samples, whole-exome sequencing with SureSelect Exome v3 baits on Illumina HiSeq 2000, average coverage ~100×; (ii) COSMIC database22, comprising 243 samples, curated set of mutations from several sources (see URLs); (iii) Stand Up to Cancer23 (SU2C-PCF), comprising 150 metastatic castration-resistant samples, paired-end, whole-exome sequencing with SureSelect Exome v4 baits on Illumina HiSeq2000, average coverage ~160×.
To identify coding and noncoding drivers from SNVs and indels, we used two previously described methods50. Coding drivers on the joint dataset (930 cancers) were identified using dNdScv, a dN/dS method designed to quantify positive selection in cancer genomes. dNdScv models somatic mutations in a given gene as a Poisson process. Inferences on selection are carried out separately for missense substitutions, truncating substitutions (nonsense and essential splice site mutations) and indels and then combined into a global P-value per gene. Noncoding recurrence was studied using NBR. Both dNdScv and NBR model the variation of the mutation rate across the genome using a negative binomial regression with covariates. First, Poisson regression is used to obtain maximum-likelihood estimates for the 192 rate parameters (r j ) describing each of the possible trinucleotide substitutions in a strand-specific manner. r j = n j /L j , where n j is the total number of mutations observed across samples of a given trinucleotide class (j) and L j is the number of available sites for each trinucleotide. These rates are used to estimate the total number of mutations across samples expected under neutrality in each element considering the mutational signatures active in the cohort and the sequence of the elements (E h = Σ j r j Lj,h). This estimate assumes no variation of the mutation rate across elements in the genome. Second, a negative binomial regression is used to refine this estimate of the background mutation rate of an element, using covariates and E h as an offset. Both methods identify genes or noncoding regions with higher than expected mutation recurrence, correcting for gene length, sequence composition, mutation signatures acting across patients and the variation of the mutation rate along the genome. A QQ plot confirmed that P-values obtained from this method in this cohort were not subject to inflation and consequent over-calling of driver genes (Supplementary Fig. 6).
Chromoplexy, characterized by highly clustered genomic breakpoints that occur in chains and are sometimes joined by deletion bridges, has been shown to be prevalent in prostate cancer25. To identify rearrangement drivers, we first used ChainFinder25 to account for any bias toward regions with chromoplexy and identified ‘unique’ rearranged regions per sample taking the midpoint between all the breakpoints ChainFinder assigns to the same chromoplexy event. Next, separately aggregating the ICGC samples with and without ERG fusions, we calculated interbreakpoint distance and performed piecewise constant fitting (PCF)74 to identify genomic regions that were recurrently rearranged in multiple samples. Rearranged regions with potential functional impact were identified using two criteria: a minimum threefold difference in the number of SVs per megabase of ERG+ and ERG– samples, and presence of at least one gene with multiple samples with truncating events—i.e., homozygous deletion, stop codon, frameshift indel or essential splice site mutation. In addition, several identified regions were significantly enriched for LOH in either ETS+ or ETS– samples, from copy number analysis (see above). The variants identified in key regions are depicted in Fig. 3.
Chemogenomics annotation of the prostate cancer network
To construct the network, we used the 71 protein products of the 73 genes identified in this study (henceforth referred to as prostate proteins) to seed a search for all possible interacting proteins in the canSAR interactome54. This interactome contains merged and curated data from the IMEx Consortium79, Phosphosite (see URLs) and other databases. It includes (i) interactions where there were more than two publications reporting experiments demonstrating the binary interaction between the two proteins, (ii) interactions where there is 3D protein structural evidence of a direct complex, (iii) interactions where there are at least two publications reporting that one protein is a substrate of the other, and (iv) interactions where there are at least two papers reporting that one protein is the product of a gene under the direct regulatory control of the other. It excludes (i) interactions that were inferred from a large immunoprecipitation experiment without follow-up to demonstrate the specific binary interaction, (ii) interactions inferred from text mining, and (iii) interactions inferred from co-occurrence in publications or from gene expression correlation.
The initial prostate cancer seeded network resulted in a large collection of 3,290 proteins that have some experimental evidence of interacting with at least one prostate protein. When we add extra proteins to the network, we wanted to ensure that we only add proteins that are more likely to function primarily through interaction with the proteins in the network rather than just be generic major hubs. To this end, we carried out the following steps. First, starting with the input (prostate protein) list, we obtained all possible first neighbors. We then computed, for each new protein, the proportion of its first neighbors that are in the original input list. To define the proteins that are most likely to function through our network, we calculated the chances of these proportions occurring in a random network. We did this by randomizing our interactome 10,000 times and computing how often the observed proportions can be achieved by chance (empirical P-value). We corrected the P-values for multiple testing and retained only proteins that had corrected FDR P < 0.05 (Supplementary Fig. 5). We performed network minimization to maintain only proteins that are strongly connected to more than one prostate protein or whose only connection is to one of the prostate proteins. We identified a prostate cancer network of 156 proteins. Using canSAR’s Cancer Protein Annotation Tool (CPAT)80, we annotated the 156 proteins with pharmacological and druggability data. We labeled proteins that are (i) targets of approved drugs, (ii) targets of drugs under clinical investigation, (iii) targets of preclinical or discovery stage compounds that are active at concentrations equal to or less than 100 nM against the protein of interest, or (iv) proteins that we predict to be druggable using our structural druggability prediction protocols80,81,82,83 but that have few or no published active inhibitors—these are potential targets for future drug discovery.
Further information on experimental design is available in the Nature Research Reporting Summary linked to this article.
Alignment and variant calling was performed using analysis pipelines in the Cancer Genome Project (CGP) at the Wellcome Trust Sanger Institute. Software versions applied to each sample are listed in Supplementary Table 9. The CGP pipelines can be downloaded (see URLs). Chromoplexy was called using ChainFinder version 1.0.1. ChainFinder can be downloaded (see URLs). The Battenberg algorithm was used to call clonal and subclonal CNAs in all samples. The Battenberg pipeline can be downloaded (see URLs). Putative drug targets were identified using CanSAR version 3.0. Data analysis was carried out using R, version 3.0.0.
Sequencing data that support the findings of this study have been deposited in the European Genome-phenome Archive with the accession code EGAS00001000262. Sample-specific accession codes are listed in Supplementary Table 7.
GDSC, http://www.cancerrxgene.org; canSAR, http://cansar.icr.ac.uk/cansar/publications/sequencing_prostate_cancers_identifies_new_cancer_genes_routes_progression_and_drug_targets/; ICGC, https://icgc.org/; dockstore-cgpwgs, https://github.com/cancerit/dockstore-cgpwgs; Battenberg pipeline, https://github.com/Wedge-Oxford/battenberg; Phosphosite, http://www.phosphosite.org/; ChainFinder, http://archive.broadinstitute.org/cancer/cga/chainfinder; COSMIC database, http://cancer.sanger.ac.uk/cosmic.
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The authors thank those men with prostate cancer and the subjects who have donated their time and their samples to the Cambridge, Oxford, The Institute of Cancer Research, John Hopkins and University of Tampere BioMediTech Biorepositories for this study. We also acknowledge support of the research staff in S4 who so carefully curated the samples and the follow-up data (J. Burge, M. Corcoran, A. George and S. Stearn). We thank M. Stratton for discussions when setting up the CR-UK Prostate Cancer ICGC Project. We acknowledge support from Cancer Research UK C5047/A14835/A22530/A17528, C309/A11566, C368/A6743, A368/A7990, C14303/A17197 (Z.K.-J., S. Merson, N.C., S.E., D.L., T. Dadaev, M.A., E.B., J.B., G.A., P.W., B.A.-L., D.S.B., C.S.C., R.A.E.), the Dallaglio Foundation (CR-UK Prostate Cancer ICGC Project and Pan Prostate Cancer Group), PC-UK/Movember (Z.K.-J.), the NIHR support to The Biomedical Research Centre at The Institute of Cancer Research and The Royal Marsden NHS Foundation Trust (Z.K.-J., N.D., S. Merson, N.C., S.E., D.L., T. Dadaev, S. Thomas, M.A., E.B., C.F., N.L., D.N., V.K., N.A., P.K., C.O., D.C., A.T., E.M., E.R., T. Dudderidge, S. Hazell, J.B., G.A., P.W., B.A.-L., D.S.B., C.S.C., R.A.E.), Cancer Research UK funding to The Institute of Cancer Research and the Royal Marsden NHS Foundation Trust CRUK Centre, the National Cancer Research Institute (National Institute of Health Research (NIHR) Collaborative Study: “Prostate Cancer: Mechanisms of Progression and Treatment (PROMPT)” (grant G0500966/75466) (D.E.N., V.G.), the Li Ka Shing Foundation (D.C.W., D.J.W.) and the Academy of Finland and Cancer Society of Finland (G.S.B.). We thank the National Institute for Health Research, Hutchison Whampoa Limited, University of Cambridge and the Human Research Tissue Bank (Addenbrooke’s Hospital), which is supported by the NIHR Cambridge Biomedical Research Centre; The Core Facilities at the Cancer Research UK Cambridge Institute, Orchid and Cancer Research UK, D. Holland from the Infrastructure Management Team, and P. Clapham from the Informatics Systems Group at the Wellcome Trust Sanger Institute. D.M.B. is supported by Orchid. C.V.’s academic time was supported by the NIHR Oxford Biomedical Research Centre (Molecular Diagnostics Theme/Multimodal Pathology sub-theme). We also acknowledge support from the Bob Champion Cancer Trust, The Masonic Charitable Foundation successor to The Grand Charity, The King Family and the Stephen Hargrave Trust (C.S.C., D.S.B.). P.W. is a Cancer Research Life Fellow. We acknowledge core facilities provided by CRUK funding to the CRUK ICR Centre, the CRUK Cancer Therapeutics Unit and support for canSAR C35696/A23187 (P.W., G.A.).