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Molecular Basis of Cisplatin Resistance in Testicular Germ Cell Tumors
research-article
Author(s):
Violeta Bakardjieva-Mihaylova
1 ,
Karolina Skvarova Kramarzova
1 ,
Martina Slamova
1 ,
Michael Svaton
1 ,
Katerina Rejlova
1 ,
Marketa Zaliova
1 ,
Alena Dobiasova
1 ,
Karel Fiser
1 ,
Jan Stuchly
1 ,
Marek Grega
2 ,
Blanka Rosova
3 ,
Roman Zachoval
4 ,
Petr Klezl
5 ,
Vaclav Eis
6 ,
Eva Kindlova
7 ,
Tomas Buchler
8 ,
Jan Trka
1 ,
Ludmila Boublikova
1
,
8
,
*
Publication date (Electronic):
06 September 2019
Journal:
Cancers
Publisher:
MDPI
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Abstract
The emergence of cisplatin (CDDP) resistance is the main cause of treatment failure and death in patients with testicular germ cell tumors (TGCT), but its biologic background is poorly understood. To study the molecular basis of CDDP resistance in TGCT we prepared and sequenced CDDP-exposed TGCT cell lines as well as 31 primary patients’ samples. Long-term exposure to CDDP increased the CDDP resistance 10 times in the NCCIT cell line, while no major resistance was achieved in Tera-2. Development of CDDP resistance was accompanied by changes in the cell cycle (increase in G1 and decrease in S-fraction), increased number of acquired mutations, of which 3 were present within ATRX gene, as well as changes in gene expression pattern. Copy number variation analysis showed, apart from obligatory gain of 12p, several other large-scale gains (chr 1, 17, 20, 21) and losses (chr X), with additional more CNVs found in CDDP-resistant cells (e.g., further losses on chr 1, 4, 18, and gain on chr 8). In the patients’ samples, those who developed CDDP resistance and died of TGCT (2/31) showed high numbers of acquired aberrations, both SNPs and CNVs, and harbored mutations in genes potentially relevant to TGCT development (e.g., TRERF1, TFAP2C in one patient, MAP2K1 and NSD1 in another one). Among all primary tumor samples, the most commonly mutated gene was NSD1, affected in 9/31 patients. This gene encoding histone methyl transferase was also downregulated and identified among the 50 most differentially expressed genes in CDDP-resistant NCCIT cell line. Interestingly, 2/31 TGCT patients harbored mutations in the ATRX gene encoding a chromatin modifier that has been shown to have a critical function in sexual differentiation. Our research newly highlights its probable involvement also in testicular tumors. Both findings support the emerging role of altered epigenetic gene regulation in TGCT and CDDP resistance development.
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SETD2-Dependent Histone H3K36 Trimethylation Is Required for Homologous Recombination Repair and Genome Stability
Introduction DNA double-stranded breaks (DSBs) are potentially lethal lesions if unrepaired, and their misrepair can give rise to genome instability, a hallmark of cancer (Jeggo and Lavin, 2009). To maintain genome stability in response to such lesions, cells employ either homologous recombination (HR) or nonhomologous end-joining (NHEJ) pathways to repair DSBs. HR is initiated by 5′ end resection to generate a 3′ single-stranded DNA (ssDNA) overhang. Resection is a two-step process initiated by removing a short oligonucleotide through the activities of the Mre11-Rad50-Nbs1 (MRN) complex and CtIP. Extensive resection is performed by Exo1 or DNA2-BLM in conjunction with Replication Protein A (RPA), which binds ssDNA and removes secondary structures (Sugiyama et al., 1998; Symington and Gautier, 2011). This, in turn, leads to RAD51 nucleofilament formation, which promotes strand invasion of a homologous chromatid, leading to accurate repair (Heyer et al., 2010). During classical NHEJ (C-NHEJ), the broken ends are rapidly bound and protected by the Ku70/Ku80 heterodimer, which acts as a platform to recruit the DNA-PK catalytic subunit (DNA-PKcs). Damaged ends are then processed and subsequently joined by the Ligase 4 (Lig4), XRCC4, XLF complex in a template-independent manner, which can lead to inaccurate repair (Lieber, 2010). DSBs may also be repaired through alternative end-joining pathways, such as microhomology-mediated end-joining (MMEJ), which do not require Ku or Lig4. Like HR, MMEJ is initiated by resection, and end-joining is mediated through annealing of short direct repeats of microhomology. MMEJ leads to deletions and is frequently associated with chromosomal rearrangements (McVey and Lee, 2008). DSB repair is further facilitated through chromatin modification by chromatin remodeling complexes, by incorporation of histone variants, and by histone modification (Smeenk and van Attikum, 2013). Histone H3K36 dimethylation has recently been proposed to facilitate NHEJ, where Metnase (SETMAR) directly mediates H3K36 dimethylation near the break site, leading to recruitment and stabilization of NBS1 and Ku70 (Fnu et al., 2011). Histone H3K36 is also trimethylated, which in mammalian cells is performed uniquely by the SETD2/HYPB methyltransferase (Edmunds et al., 2008). H3K36me3 is associated with transcriptional elongation and is found in gene coding regions, peaking at 3′ ends (Edmunds et al., 2008). SETD2-dependent H3K36 trimethylation facilitates a number of processes within the cell, including splicing, repression of intragenic transcripts, and chromatin accessibility (Li et al., 2013; Wagner and Carpenter, 2012). SETD2 is also mutated in a number of cancer types, including breast, lung, acute lymphoblastic leukemia, clear cell renal cell carcinoma, and glioma, supporting its role as a tumor suppressor (Al Sarakbi et al., 2009; Dalgliesh et al., 2010; Fontebasso et al., 2013; Newbold and Mokbel, 2010; Zhang et al., 2012). SETD2-dependent H3K36 trimethylation has recently been shown to regulate DNA mismatch repair (Li et al., 2013). However, SETD2-deficient cancers exhibit a wide range of mutations, including insertions, deletions (indels), and chromosomal aberrations (Sato et al., 2013; Zhu et al., 2014), suggesting an additional role for SETD2 in genome stability. Additionally, H3K36 methylation is regulated by the KDM4/JMJD2 family of histone demethylases. These contain JmjN-JmjC and tandem-Tudor domains that specifically remove the tri- and dimethyl forms of both H3K9 and H3K36, with preference for the trimethyl form being observed in the case of KDM4A/JMJD2A (Couture et al., 2007; Klose et al., 2006; Whetstine et al., 2006). KDM4 family proteins are frequently overexpressed in cancers and are associated with poor patient survival (Berry and Janknecht, 2013; Black et al., 2013). Here, we investigate the role of SETD2-dependent H3K36me3 in maintaining genome stability. We establish a role for H3K36me3 in HR repair by facilitating resection. In addition, we define a role for SETD2 and RAD51 in maintaining genome stability through suppressing MMEJ at break sites. Results SETD2 Suppresses Break-Induced Mutations Tumor suppressors can suppress carcinogenesis by preventing genome instability, a hallmark of cancer. To investigate the impact of loss of SETD2, a tumor suppressor, on genome stability following DSB induction, we developed an I-SceI-induced loss of function assay (HPRT+:I-SceI). Intron-encoded endonuclease 1 from Saccharomyces cerevisiae (I-SceI) is a rare-cutting endonuclease that has no predicted recognition site in the mammalian genome (Jasin, 1996). An I-SceI recognition sequence was inserted into the endogenous HPRT exon 6 in HT1080 (fibrosarcoma) cells. The I-SceI site maintains the HPRT reading frame and makes only a single amino acid change that does not impair the function of HPRT (Figure 1A). Following I-SceI induction, inaccurate DSB repair generates HPRT-negative mutants, which can be selected for using 6-thioguanine (6-TG), and indels can be detected by PCR amplification using primers flanking the break site (Figure 1A). Following small interfering RNA (siRNA) knockdown and I-SceI induction, the frequency of HPRT loss (mutation frequency) in SETD2-depleted cells was 1.65%, significantly higher (p = 0.0014) than that of nontargeting controls (1.1%). These findings resembled those in RAD51-depleted cells in which the frequency of HPRT loss was also significantly increased to 1.88% (p = 0.0022) compared to nontargeting controls (Figure 1B). SETD2 and RAD51 codepletion (siS+SiR) also significantly increased the mutation frequency to 1.52% (p = 0.037), which was similar to the frequency observed following either SETD2 or RAD51 depletion (Figure 1B). We confirmed that I-SceI protein expression and cleavage efficiency were the same in cells transfected with either control siRNA or SETD2 siRNA (Figures S1A–S1C). In addition, HPRT gene transcription was not affected by SETD2 knockdown, as shown by quantitative RT-PCR (qRT-PCR) of spliced and unspliced HPRT mRNA (Figures S1D and S1E). Therefore, the mutation frequency in this system is likely to be directly associated with misrepair of the DSB at the I-SceI site. Further, we studied the mutation patterns across the break site of 30 individually isolated HPRT-negative clones from each background by PCR amplification and sequencing. HPRT-negative clones from cells treated with nontargeting control siRNA indicated the presence of microdeletions of 2–5 bp, consistent with DSB repair by C-NHEJ. Larger microdeletions were also observed that were associated with regions of microhomology located either side of the break site, consistent with DSB repair through MMEJ (Figures 1C and S1F). Sequence analysis indicated that SETD2-depleted cells exhibited a significant increase in average deletion length (11 bp, p < 0.0001) compared to control cells (5 bp) (Figures 1C and 1D). RAD51-depleted cells exhibited a further significant increase in the average deletion length (22 bp, p = 0.014) (Figures 1C and 1D). RAD51 and SETD2 codepletion generated deletion lengths (an average of 8 bp) comparable to those observed following SETD2 knockdown, which in turn was significantly higher than the nontargeting control (p = 0.026) (Figures 1C and 1D). Insertions at the HPRT break site were also detected. However, the frequency and size of the insertions was not significantly altered between SETD2-depleted cells, SETD2/RAD51 codepleted and nontargeting controls. We next examined the mechanism for misrepair in cells exhibiting deletions. We found that the frequency of microhomology-mediated end-joining (MMEJ) was significantly increased in SETD2 and RAD51-depleted cells (p < 0.05) (Figures 1E and S1G). This suggested that HR-deficient cells use MMEJ as an alternative repair mechanism, which gives rise to a mutation signature of sequence loss between regions of microhomology. These results support a role for SETD2 in tumor suppression by maintaining genome stability through preventing deleterious mutations arising in response to DSBs. These findings further identify a similar break-induced mutation signature between SETD2- and RAD51-depleted cells, suggesting an early role for SETD2 in HR repair. SETD2 Is Required for Homologous Recombination Repair To test the role for SETD2 in DSB repair, we depleted SETD2 in U2OS and HeLa cells using two independent siRNAs and measured the clonogenic survival following exposure to DNA-damaging agents. SETD2 knockdown significantly reduced survival after treatment with mitomycin C (MMC), camptothecin (CPT), or ionizing radiation (IR) in U2OS cells and HeLa cells compared to nontargeting controls (Figures 2A and S2A). These results identify a role for SETD2 in the cellular response to DNA damage and are consistent with a role for SETD2 in DSB repair. SETD2 knockdown was validated by western blotting (Figure 2B). Western blotting also showed that SETD2 knockdown significantly reduced global levels of H3K36me3, confirming SETD2 as the major H3K36me3 methyltransferase in human cells (Figure 2B). Because RAD51 is more likely to be affected among DNA repair genes by siRNA off-target effects (Adamson et al., 2012), we also confirmed that SETD2 knockdown by siRNA did not affect the RAD51 protein levels (Figure 2B). We confirmed that SETD2 knockdown had no effect on the expression of major HR proteins, including BRCA2, RAD50, CtIP, MRE11, and RPA and the NHEJ protein KU80 (Figure S2B). We also observed no changes to the cell cycle after SETD2 knockdown (Figure S2C). Both MMC and CPT induce S phase-specific DSBs, which are predominantly repaired by HR (Arnaudeau et al., 2001; Moynahan et al., 2001). To test a possible role for SETD2 in HR repair, we used a well-characterized GFP-based reporter for HR (DR-GFP). The HR reporter contains an I-SceI recognition sequence, which upon I-SceI expression is cleaved to generate a DSB. DSB repair by HR using a direct repeat within the reporter cassette as a template results in an intact GFP gene (Figure 2C)(Pierce et al., 1999). SETD2 knockdown by two independent siRNAs significantly reduced HR repair of an I-SceI induced DSB by 67%–77% compared to nontargeting control cells (p < 0.0003) (Figure 2D). As a control for the HR reporter, knockdown of RAD51 reduced HR activity by 97% (p < 0.0001), whereas DNA-PKcs inhibitor NU7441 had no effect on HR (Figure 2D). In contrast, SETD2 knockdown had no effect on repair of an I-SceI induced DSB in an NHEJ reporter (IRES-TK-EGFP) in which joining of two I-SceI sites separated by a thymidine kinase gene (TK) results in GFP expression (Figures 2E and 2F)(Ogiwara et al., 2011). As a control, RAD51 had no effect on the same NHEJ reporter, whereas the DNA-PK inhibitor NU7441 reduced NHEJ activity by 98% (p < 0.0001) (Figure 2F). These results identify a role for SETD2 in HR repair. An Early Role for SETD2 in Homologous Recombination To further investigate the role of SETD2 in DSB repair, we examined the rate of repair following IR. We used γH2AX foci as a marker of DNA damage and found that following exposure to 5 Gy IR, SETD2 knockdown significantly delayed removal of γH2AX foci at 48 hr (p < 0.003) compared to nontargeting controls, supporting a role for SETD2 in efficient DSB repair (Figure 3A). We next investigated the effect of SETD2 depletion on the recruitment of HR repair proteins following DNA damage. SETD2 knockdown resulted in significantly reduced RAD51 foci formation 4 and 8 hr after exposure to IR compared to nontargeting controls (p < 0.0013) (Figure 3B). SETD2 knockdown also significantly reduced RAD51 foci formation after exposure to CPT (p < 0.05) (Figure S2D). We also examined the effect of SETD2 depletion on RPA recruitment, where we induced DSBs in S phase cells using CPT or MMC. A significant reduction in RPA foci formation was observed after CPT or MMC treatment following SETD2 knockdown compared to nontargeting controls (p < 0.023) (Figures 3C and S2E). SETD2 knockdown leads to reduced H3K36me3 in the nuclei as confirmed by costaining the cells with antibodies against H3K36me3 and RPA (Figure S2F). Together, these findings support a role for SETD2 in promoting HR through facilitating recruitment of RPA and RAD51 to DNA damage sites. To further confirm the impact of SETD2 loss on the recruitment of HR factors to the site of DSBs, we performed chromatin immunoprecipitation (ChIP) near the I-SceI site in the DR-GFP cassette (Figure S3A). The I-SceI site is located within the DR-GFP cassette under the control of the highly active CMV promoter, and thus H3K36me3, which is associated with active transcription, is predicted to be present throughout the cassette. After induction of DSBs by I-SceI, no change in H3K36me3 or SETD2 levels was detected proximal or distal to the break site after 8, 24, or 48 hr following I-SceI transfection (Figure S3B). The results were also quantified by ChIP-qPCR (Figure S3C). Although other groups also observed no changes to H3K36me3 level after DSB, globally (Fnu et al., 2011) or locally at the break site (Pei et al., 2011; Aymard et al., 2014), we were surprised by the lack of reduction in H3K36me3 or SETD2 knowing that regional transcriptional silencing has been observed at sites of DSBs (Shanbhag et al., 2010). We speculate that the unexpected maintenance of H3K36me3 at the DSB site by SETD2 may serve as an anchor to mediate HR repair. We monitored recruitment of major HR factors to the DSB site in the ChIP system. RPA and RAD51 were recruited to the DSB site, resulting in increased binding to the “up” site 18 hr after I-SceI transfection, compared to uncut controls (Figure 3D). SETD2 depletion substantially reduced the recruitment of RPA and RAD51 after the I-SceI cut (Figure 3D). SETD2 protein depletion was validated by western blotting (Figure S3D), and was further confirmed by the absence of SETD2 or H3K36me3 at the break site (Figure 3D). These findings are consistent with the RPA and RAD51 foci data, indicating that SETD2 is required to recruit RPA and RAD51 to DSB sites. Together, these findings confirm a presynaptic role for SETD2 in facilitating HR repair of DSBs. H3K36me3 Is Required for Homologous Recombination Repair To investigate if the methyltransferase activity of SETD2 is required for its role in HR, we generated two doxycycline-inducible U2OS cell lines, where either a wild-type SETD2 cDNA or a methyltransferase-dead SETD2 cDNA was inserted behind a Tet operator in the genome of U2OS cells. Upon addition of doxycycline to the cell culture medium, the Tet operator was derepressed and the wild-type or mutant SETD2 was expressed. Both cDNA constructs were mutated so that they are refractory to SETD2 siRNA#3. The methyltransferase-dead SETD2 was mutated at two amino acid residues located within the catalytic site of the SET domain, which are conserved with yeast Set2. The corresponding mutations in yeast were shown to abolish the methyltransferase activity (Rea et al., 2000;Strahl et al., 2002) (Figure 4A). Both arginine (R) and cysteine (C) residues were mutated in the methyltransferase-dead (MD) cDNA by site-directed mutagenesis (see Supplemental Experimental Procedures). Western blot analysis confirmed that doxycycline-induced expression of wild-type SETD2 could restore the level of H3K36me3 after siSETD2 treatment, whereas expression of methyltransferase-dead SETD2 could not restore the level of H3K36me3 (Figure S4A). We depleted endogenous SETD2 with siRNA#3 and examined the rescue of the phenotype by the two constructs. Although the wild-type construct increased foci formation of RPA and RAD51 after DNA damage in siSETD2 cells, the methyltransferase-dead mutant did not (Figures 4B and 4C). This is consistent with the observation that the wild-type construct partially restored HR efficiency whereas the methyltransferase-dead (MD) mutant did not (Figure 4D). Five to six clones containing each construct were independently isolated and tested. All WT clones were able to restore HR efficiency, but none of the MD clones showed rescue (Figures S4B and S4C). The observation that the methyltransferase-dead SETD2 mutant could not rescue HR efficiency suggested that H3K36me3 is required for efficient HR. To test this further, we used SETD2-independent approaches to disrupt H3K36me3 levels. First, we overexpressed KDM4A, an oncogene and a demethylase specific for tri- and dimethylated histone H3K36 and H3K9, predicting that overexpression of KDM4A would reduce the level of H3K36me3 and thus impair HR. Therefore, a doxycycline-inducible KDM4A-expression U2OS cell line was generated. Upon addition of doxycycline, KDM4A was overexpressed, which led to a global reduction in H3K36me3 levels (Figure 4E). We then compared the recruitment of the HR factors in this cell line with or without KDM4A overexpression. We found that KDM4A overexpression inhibited the formation of RPA and RAD51 foci after DNA damage (Figures 4F and 4G). Accordingly, KDM4A overexpression also reduced HR efficiency as assessed by the HR reporter (p = 0.0078) (Figure 4H). Using the same cell line without KDM4A cDNA integration, we found that doxycycline itself had no effect on HR efficiency of U2OS cells (Figure S4D). These findings indicate that overexpression of the oncogene KDM4A significantly reduces levels of H3K36me3 and HR. Second, we utilized a dominant-negative mutation in the histone H3.3 gene (H3.3K36M) that was recently shown to reduce H3K36me3 specifically without affecting other histone methylations (Lewis et al., 2013). Accordingly, in cells expressing the mutant H3.3K36M transgene, global levels of H3K36me3 were depleted compared to the wild-type H3.3 control (Figure 4I). Cells expressing H3.3K36M also exhibited delayed RPA and RAD51 foci formation following DNA damage (Figures 4J and 4K), consistent with similar delays seen in KDM4A overexpressing cells or SETD2-depleted cells. Neither KDM4A overexpression nor H3.3K36M expression affected cell cycle progression as assessed by BrdU incorporation (Figures S4E and S4F), thus excluding the possibility that the reduction in RPA, RAD51 foci formation and HR efficacy was due to cell cycle effects. The common feature of all three different systems (SETD2 knockdown and rescue, KDM4A overexpression, and H3.3K36M expression) was the reduction in the recruitment of HR proteins and HR efficiency associated with reduced H3K36me3. Together these findings confirm a role for H3K36me3 in HR. SETD2 Promotes DSB End Resection Lens epithelium-derived growth factor p75 (LEDGF) binds to H3K36me3 constitutively through its PWWP domain. Upon DNA damage LEDGF recruits CtIP, which facilitates the resection step during HR repair (Daugaard et al., 2012). Accordingly, we were able to coprecipitate LEDGF and H3K36me3, independently of DNA damage in U2OS cells (Figure 5A). LEDGF binding to chromatin was reduced upon SETD2 depletion, consistent with preferential binding of LEDGF to H3K36me3 (Eidahl et al., 2013), and was DSB independent (Figure 5B). We predicted that the reduced LEDGF binding following SETD2 depletion would reduce recruitment of CtIP upon damage. We therefore used micro-irradiation to induce localized DSBs and studied the recruitment of CtIP using a U2OS cell line expressing GFP-tagged CtIP (Sartori et al., 2007). We found CtIP to be localized to sites of DNA damage marked by γH2AX (Figure 5C). In contrast, SETD2 knockdown resulted in reduced recruitment of CtIP to the DNA damage site as marked by γH2AX (Figure 5C). If SETD2 facilitates HR through LEDGF binding and CtIP recruitment, we would expect SETD2 to be epistatic with CtIP. Indeed, codepletion of SETD2 and CtIP (siC+siS) showed the same level of reduction in HR as CtIP-depletion alone (Figures 5D and S5A). These findings are consistent with the hypothesis that SETD2-dependent H3K36me3 promotes recruitment of CtIP by LEDGF to break sites. Reduced CtIP recruitment, together with the reduced RPA foci formation following SETD2 depletion, suggested that SETD2-dependent H3K36 trimethylation might promote DSB resection. To directly measure resection, we used an in vivo resection assay (Zhou et al., 2014) in which levels of ssDNA were assessed at increasing distances from a specific DSB induced by the AsiSI restriction enzyme, which was found to recruit RAD51 (Iacovoni et al., 2010; Aymard et al., 2014). Following DSB induction (+4OHT), ssDNA was readily detected 231 bp from the AsiSI break site (DSB-II in Aymard et al., 2014) and at reduced levels further from the break site. Following CtIP depletion, ssDNA levels were significantly reduced at 231 bp (p = 2.2 × 10−5) and 918 bp (p = 0.057) from the break site compared to control siRNAs (Figure 5E). Following SETD2 depletion, ssDNA levels were also significantly reduced at 231 bp (p = 0.021) and 918 bp (p = 0.037) from the break site compared to control siRNAs (Figure 5F), and SETD2 depletion did not affect DSB induction at this site as assayed by XRCC4 ChIP (Figure S5B), thus confirming a role for SETD2 in facilitating DSB resection. Discussion The SETD2 gene encoding the H3K36 trimethyltransferase is among the most mutated in human cancers (Lawrence et al., 2014). Yet, how SETD2 functions to suppress the genome instability frequently observed in SETD2-deficient cancer cells is unknown (Sato et al., 2013; Zhu et al., 2014). Here, we describe a role for SETD2-dependent H3K36 trimethylation in facilitating error-free DSB repair and genome stability through efficient HR. Our data support a model in which LEDGF is constitutively anchored to H3K36me3-decorated chromatin through its PWWP domain. Upon DNA damage, chromatin-bound LEDGF recruits CtIP, which promotes resection—an essential step in HR repair (Figure 6A). In the absence of SETD2-dependent H3K36me3, the chromatin association of LEDGF is compromised, and DNA damage-induced CtIP recruitment is impaired, leading to reduced resection and HR and an increase in error-prone MMEJ and subsequent genome instability (Figure 6B). Our data do not, however, exclude the possibility that other H3K36me3 binding factors may also contribute to efficient HR. The involvement of H3K36me3 in HR raises the question as to whether this histone mark is DSB inducible or preset to determine the choice of repair pathways. We did not detect any appreciable increase in H3K36me3 levels at the break site following I-SceI induction. These findings are consistent with previous observations indicating that H3K36me3 levels do not change following DNA damage, either globally (Fnu et al., 2011) or locally (Aymard et al., 2014; Pei et al., 2011). Given that SETD2-dependent H3K36 trimethylation strongly correlates with actively transcribed regions of the genome (Edmunds et al., 2008; Yoh et al., 2008), our findings thus support a spatial link between HR and transcriptionally active, H3K36me3 decorated regions of the genome. Indeed, because HR repair is usually error free, this would be expected to protect genome integrity following DSB induction or replication fork collapse at such sites, which would be important in maintaining cell homeostasis. Genome-wide sequencing has suggested certain mutation signatures are associated with specific cancer types (Alexandrov et al., 2013) but lacks the power to identify driver mutations that give rise to that mutation signature. Here, we use a system to determine the impact of individual gene dysfunction on mutation patterns arising during the repair of a single DSB. We show that, by depleting SETD2 or RAD51 HR proteins, DSB induction leads to significantly increased mutation frequencies associated with deletions within six cell cycles (5 days). These findings identify a role for SETD2, like RAD51, in suppressing break-induced mutations. Further mechanistic analysis indicated that deletions arising from SETD2 or RAD51 depletion resulted from a significant increase in error-prone MMEJ. We propose that MMEJ arises from failed HR. SETD2 loss results in partial resection, short ssDNA 3′ ends and reduced RPA foci. These partially resected ends are insufficient to promote efficient HR and are no longer substrates for C-NHEJ. Instead, these partially resected ends become substrates for MMEJ. Similarly, following RAD51 depletion, whereas resection proceeds normally, HR is blocked presynaptically, resulting in the accumulation of longer unrepaired resected ends. Such ends are no longer substrates for C-NHEJ, and, because HR cannot proceed, the ends are instead rejoined by MMEJ. Extensive resection following RAD51 loss facilitates annealing between regions of microhomology further from the break site, thus leading to larger deletions than those observed for SETD2. The observations that HR was significantly reduced whereas C-NHEJ was unchanged following SETD2 or RAD51 depletion are consistent with this model. Together, these findings suggest a role for HR in maintaining genome stability through suppression of MMEJ. This “real-time” mutation study can therefore provide important insights into how loss of these genes contributes to somatic mutation and intratumor heterogeneity, and it highlights the importance of HR repair genes in maintaining genome stability. SETD2 is a tumor suppressor and its loss is an important step in the progression of a number of cancer types (Dalgliesh et al., 2010; Fontebasso et al., 2013; Gerlinger et al., 2012). Our findings provide mechanistic insights into how SETD2 functions as a tumor suppressor. Moreover, all three Tudor domain-containing KDM4/JMJD2 proteins (A, B, and C) are frequently overexpressed in various cancers, including breast, colorectal, lung, and prostate (Berry and Janknecht, 2013). Our findings predict that overexpression of these KDM4 family members causes genome instability and tumorigenesis through loss of H3K36me3-mediated HR repair. The identification of a role for H3K36me3 in facilitating HR repair, in addition to its recently described role in mismatch repair (Li et al., 2013), establishes the H3K36me3 chromatin mark as an important functional node for maintaining genome stability. Experimental Procedures I-SceI-Inducible HPRT Mutation Assay The reporter HT1080 cell line with a functional but I-SceI-cleavable HPRT gene was generated by inserting an 18 bp I-SceI recognition site into exon 6 of the HPRT gene using gene targeting. Reporter cells were transfected with the I-SceI plasmid 48 hr after siRNA knockdowns. The cells were allowed to repair in a nonselective medium for 5 days before seeding and selecting for HPRT-negative cells using 6-TG (15 μg/ml). The mutation frequency was calculated as the number of HPRT-negative colonies divided by number of cells plated, after correcting for plating efficiency. To determine the mutation sequence, individual single-cell colonies were isolated, and genomic DNA spanning the DSB junction was PCR-amplified using the primers flanking the I-SceI site. Sequence alignment of the PCR products was conducted using the DNA Data Bank of Japan (DDBJ) ClustalW program. In Vivo DNA End Resection Assay DIvA cells (DSB Inducible via AsiSI [Aymard et al., 2014]) were transfected with siRNA using the Cell Line Nucleofactor kit V (Amaxa) according to the manufacturer’s instructions. Forty-eight hours after siRNA transfection, cells were treated or not with 300 nM of 4-hydroxytamoxifen (4OHT) (Sigma; H7904) for 4 hr. DNA was purified using QIAGEN DNeasy kit. Single-stranded DNA generated at an AsiSI-induced DSB was analyzed using the procedure described in Zhou et al. (2014), with the modifications described in Supplemental Experimental Procedures. The percentage of ssDNA was calculated with the following equation: ssDNA% = 1/(2(ΔCt−1) + 0.5) ×100, where ΔCt is calculated by subtracting the Ct obtained from mock-digested sample from the BanI-digested sample. Generation of Inducible KDM4A Overexpression Cell Lines U2OS-Flp-In/T-REx cells express the Tet repressor and contain a single Flp-In site (Gordon et al., 2009). U2OS-Flp-In/T-REx/DRGFP cells were generated by integrating the pDR-GFP expression plasmid (Pierce et al., 1999) into the U2OS-Flp-In/T-REx cells. The KDM4A cDNA (Genecopoeia) was cloned into the pDESTfrtto destination vector by a Gateway LR reaction (Life Technologies). U2OS-Flp-In/T-REx or U2OS-Flp-In/T-REx/DRGFP cells were cotransfected with the resulting KDM4A destination plasmid and pOG44 (Invitrogen). Stable clones that express the KDM4A protein under the control of the T-REx system were subjected to the treatments indicated. Data Analysis Data points from individual assays represent mean ± SEM. Statistical significance between two conditions was assessed by a two-tailed unpaired t test. ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001, and n.s. represents p ≥ 0.05. Full details of all other experimental procedures are given in the Supplemental Experimental Procedures. Author Contributions S.X.P., S.A., L.-P.Z., S.S., and F.A. designed and performed experiments. The HPRT+:I-SceI assay was developed by A.C.G.P. The FLIP-IN cells and plasmids were developed by C.Z.B., T.C.H., A.C.G.P., T.H., N.B.L.T., and G.L. supervised the work. T.C.H. and S.X.P. wrote the manuscript with discussion and input from all authors. Experiments in Figure 1 were performed by S.A., with technology developed by A.C.G.P.; Figure 2 was performed by S.X.P. and S.A.; Figure 3 was performed by S.X.P. and L.-P.Z.; Figure 4 was performed by S.S. and S.X.P., with technology developed by C.Z.B.; Figure 5 was performed by S.S., L.-P.Z., S.X.P., and F.A.; Figure 6 was produced by S.X.P. Experiments in Figure S1 were performed by S.A. and S.S.; Figure S2 was performed by S.X.P. and S.A.; Figure S3 was performed by L.-P.Z.; Figure S4 was performed by S.X.P. and S.S.; Figure S5 was performed by F.A. and S.X.P.
- Record: found
- Abstract: found
- Article: not found
Prevalence of Homologous Recombination–Related Gene Mutations Across Multiple Cancer Types
The prevalence of homologous recombination DNA damage repair (HR-DDR) deficiencies among all tumor lineages is not well characterized. Therapy directed toward homologous recombination DDR deficiency (HRD) is now approved in ovarian and breast cancer, and there may be additional opportunities for benefit for patients with other cancers. Comprehensive evaluations for HRD are limited in part by the lack of a uniform, cost-effective method for testing and defining HRD. Molecular profiles of 52,426 tumors were reviewed to identify pathogenic mutations in the HR-DDR genes ARID1A, ATM, ATRX, BAP1, BARD1, BLM, BRCA1/2, BRIP1, CHEK1/2, FANCA/C/D2/E/F/G/L, MRE11A, NBN, PALB2, RAD50, RAD51, RAD51B, or WRN. From solid tumors submitted to Caris Life Sciences, molecular profiles were generated using next-generation sequencing (NGS; average read depth, 500×). A total of 17,566 tumors were sequenced with NGS600 (n = 592 genes), and 34,860 tumors underwent hotspot Illumina MiSeq platform testing (n = 47 genes). Of the tumors that underwent NGS600 testing, the overall frequency of HR-DDR mutations detected was 17.4%, and the most commonly mutated lineages were endometrial (34.4%; n = 1,475), biliary tract (28.9%; n = 343), bladder (23.9%; n = 201), hepatocellular (20.9%; n = 115), gastroesophageal (20.8%; n = 619), and ovarian (20.0%; n = 2,489). Least commonly mutated lineages included GI stromal (3.7%; n = 108), head and neck (6.8%; n = 206), and sarcoma (9.3%; n = 592). ARID1A was the most commonly mutated gene (7.2%), followed by BRCA2 (3.0%), BRCA1 (2.8%), ATM (1.3%), ATRX (1.3%), and CHEK2 (1.3%). HR-DDR mutations were seen in 17.4% of tumors across 21 cancer lineages, providing a path to explore the role of HRD-directed therapies, including poly-ADP ribose polymerase inhibitors, DNA-damaging chemotherapies, and newer agents such as ATR inhibitors.
- Record: found
- Abstract: found
- Article: not found
Bevacizumab: an angiogenesis inhibitor for the treatment of solid malignancies.
T Shih, Celeste Lindley (2006)
Bevacizumab is a recombinant humanized monoclonal antibody that was approved by the US Food and Drug Administration (FDA) in February 2004 for use as part of combination therapy with fluorouracil-based regimens for metastatic colorectal cancer (mCRC). This article reviews the mechanism of action, clinical pharmacology, and pharmacodynamic and pharmacokinetic properties of bevacizumab. It summarizes data on the clinical efficacy and tolerability of bevacizumab from Phase II/III trials in mCRC, breast cancer, non-small-cell lung cancer, and renal cell carcinoma and preliminary data from investigational studies in pancreatic cancer and soft-tissue sarcomas. The dosing and administration of bevacizumab also are discussed. A comprehensive search of the English-language literature indexed on MEDLINE (1966-April 2006) was performed using the terms Avastin, bevacizumab, vascular endothelial growth factor, angiogenesis, and bevacizumab plus colorectal cancer, breast cancer, non-small-cell lung cancer, pancreatic cancer, and renal cell carcinoma. Published abstracts from American Society of Clinical Oncology annual meetings from 2002 to 2006, FDA submission documents, and the product information for bevacizumab also were reviewed. Pertinent review articles, preclinical studies, and editorials and all published Phase II/III clinical trials were selected for review. The reference lists of identified articles were examined for additional publications. Bevacizumab exhibits linear pharmacokinetics in the dose range between 0.3 and 10 mg/kg q2-3wk and steady state, which is reached in approximately 100 days. The estimated t(1/2) of bevacizumab is approximately 20 days. After correction for body weight, clearance and V(d) are reported to be a respective 26% and 22% higher in men than in women. Statistical analyses have not been performed, however, and the clinical impact of these gender differences has not been assessed. No dose adjustment is currently recommended based on age, sex, or renal function. Bevacizumab has been reported to result in changes in exposure to concomitant chemotherapy, although formal drug-interaction studies have not been performed. Surrogate markers of disease progression or treatment response to bevacizumab have been studied, but no significant correlations with any outcome measure have been found. When combined with standard chemotherapy regimens, bevacizumab has been associated with significant improvements compared with chemotherapy alone in the efficacy end points of overall survival, progression-free survival, and response rates in patients with mCRC (all, P < 0.05). Based on these findings, bevacizumab is considered a first-line option for this disease. Combination bevacizumab regimens have been associated with clinical activity in breast cancer, non-small-cell lung cancer, renal cell carcinoma, pancreatic cancer, and soft-tissue sarcoma. The observed toxicities of bevacizumab include hypertension, proteinuria, mild to moderate hemorrhage, wound healing complications, and thromboembolic events. A bevacizumab dose of 5 mg/kg q2wk has been established as effective and well tolerated in the setting of mCRC. A variety of dosing schemes have been investigated in other solid neoplasms, but no consensus has been reached on the optimal bevacizumab regimen. Bevacizumab is effective and generally well tolerated as first-line therapy for mCRC. The results from clinical studies of bevacizumab as a single agent or as part of combination regimens for breast cancer, non-small-cell lung cancer, renal cell carcinoma, and other solid malignancies have been promising. The most effective regimens for various malignancies and predictive markers of treatment response have not been fully determined. Combining bevacizumab with chemotherapy or other novel targeted agents appears to be a rational approach that may enhance efficacy while limiting the traditional nonselective toxicities.
Author and article information
Contributors
Ludmila Boublikova:
On behalf of
: on behalf of TTIP—Testicular Tumors Investigation Group Prague
Journal
Journal ID (nlm-ta): Cancers (Basel)
Journal ID (iso-abbrev): Cancers (Basel)
Journal ID (publisher-id): cancers
Title:
Cancers
Publisher:
MDPI
ISSN
(Electronic):
2072-6694
Publication date
(Electronic):
06
September
2019
Publication date Collection:
September
2019
Volume: 11
Issue: 9
Electronic Location Identifier: 1316
Affiliations
[1
]Department of Pediatric Hematology and Oncology, CLIP, 2nd Faculty of Medicine, Charles
University and University Hospital Motol, 150 00 Prague, Czech Republic (V.B.-M.)
(K.S.K.) (M.S.) (M.S.) (K.R.) (M.Z.) (A.D.) (K.F.) (J.S.) (J.T.)
[2
]Department of Pathology and Molecular Medicine, 2nd Faculty of Medicine, Charles University
and University Hospital Motol, 150 00 Prague, Czech Republic
[3
]Department of Pathology and Molecular Medicine, 3rd Faculty of Medicine, Charles University
and Thomayer Hospital, 140 00 Prague, Czech Republic
[4
]Department of Urology, 3rd Faculty of Medicine, Charles University and Thomayer Hospital,
140 00 Prague, Czech Republic
[5
]Department of Urology, 3rd Faculty of Medicine, Charles University and University
Hospital Kralovske Vinohrady, 100 00 Prague, Czech Republic
[6
]Department of Pathology, 3rd Faculty of Medicine, Charles University and University
Hospital Kralovske Vinohrady, 100 00 Prague, Czech Republic
Author notes
Author information
Martina Slamova
https://orcid.org/0000-0003-2966-3687
Jan Stuchly
https://orcid.org/0000-0003-1896-9533
Article
Publisher ID: cancers-11-01316DOI: 10.3390/cancers11091316
PMC ID: 6769617
PubMed ID: 31500094
SO-VID: dd08c692-6263-4b94-b8d5-6eda8d97dee9
Copyright © © 2019 by the authors.
License:
Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license ( http://creativecommons.org/licenses/by/4.0/).
History
Date
received
: 10
June
2019
Date
accepted
: 14
August
2019
Categories
Subject:
Communication
Data availability: