SIRT 7 promotes genome integrity and modulates non‐homologous end joining DNA repair

Abstract

Article25 May 2016Open Access Source DataTransparent process SIRT7 promotes genome integrity and modulates non-homologous end joining DNA repair Berta N Vazquez Berta N Vazquez Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Joshua K Thackray Joshua K Thackray Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Nicolas G Simonet Nicolas G Simonet Chromatin Biology Laboratory, Cancer Epigenetics and Biology Program (PEBC), Bellvitge Biomedical Research Institute (IDIBELL), Barcelona, Spain Search for more papers by this author Noriko Kane-Goldsmith Noriko Kane-Goldsmith Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Paloma Martinez-Redondo Paloma Martinez-Redondo Chromatin Biology Laboratory, Cancer Epigenetics and Biology Program (PEBC), Bellvitge Biomedical Research Institute (IDIBELL), Barcelona, Spain Search for more papers by this author Trang Nguyen Trang Nguyen Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Samuel Bunting Samuel Bunting Department of Molecular Biology and Biochemistry, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Alejandro Vaquero Alejandro Vaquero Chromatin Biology Laboratory, Cancer Epigenetics and Biology Program (PEBC), Bellvitge Biomedical Research Institute (IDIBELL), Barcelona, Spain Search for more papers by this author Jay A Tischfield Jay A Tischfield Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Lourdes Serrano Corresponding Author Lourdes Serrano orcid.org/0000-0002-3409-9366 Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Berta N Vazquez Berta N Vazquez Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Joshua K Thackray Joshua K Thackray Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Nicolas G Simonet Nicolas G Simonet Chromatin Biology Laboratory, Cancer Epigenetics and Biology Program (PEBC), Bellvitge Biomedical Research Institute (IDIBELL), Barcelona, Spain Search for more papers by this author Noriko Kane-Goldsmith Noriko Kane-Goldsmith Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Paloma Martinez-Redondo Paloma Martinez-Redondo Chromatin Biology Laboratory, Cancer Epigenetics and Biology Program (PEBC), Bellvitge Biomedical Research Institute (IDIBELL), Barcelona, Spain Search for more papers by this author Trang Nguyen Trang Nguyen Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Samuel Bunting Samuel Bunting Department of Molecular Biology and Biochemistry, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Alejandro Vaquero Alejandro Vaquero Chromatin Biology Laboratory, Cancer Epigenetics and Biology Program (PEBC), Bellvitge Biomedical Research Institute (IDIBELL), Barcelona, Spain Search for more papers by this author Jay A Tischfield Jay A Tischfield Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Lourdes Serrano Corresponding Author Lourdes Serrano orcid.org/0000-0002-3409-9366 Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Author Information Berta N Vazquez1, Joshua K Thackray1, Nicolas G Simonet2, Noriko Kane-Goldsmith1, Paloma Martinez-Redondo2, Trang Nguyen1, Samuel Bunting3, Alejandro Vaquero2, Jay A Tischfield1 and Lourdes Serrano 1 1Department of Genetics, Human Genetics Institute of New Jersey, Rutgers University, Piscataway, NJ, USA 2Chromatin Biology Laboratory, Cancer Epigenetics and Biology Program (PEBC), Bellvitge Biomedical Research Institute (IDIBELL), Barcelona, Spain 3Department of Molecular Biology and Biochemistry, Rutgers University, Piscataway, NJ, USA *Corresponding author. Tel: +1 848 445 9577; E-mail: [email protected] The EMBO Journal (2016)35:1488-1503https://doi.org/10.15252/embj.201593499 See also: S Paredes & KF Chua (July 2016) PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Sirtuins, a family of protein deacetylases, promote cellular homeostasis by mediating communication between cells and environment. The enzymatic activity of the mammalian sirtuin SIRT7 targets acetylated lysine in the N-terminal tail of histone H3 (H3K18Ac), thus modulating chromatin structure and transcriptional competency. SIRT7 deletion is associated with reduced lifespan in mice through unknown mechanisms. Here, we show that SirT7-knockout mice suffer from partial embryonic lethality and a progeroid-like phenotype. Consistently, SIRT7-deficient cells display increased replication stress and impaired DNA repair. SIRT7 is recruited in a PARP1-dependent manner to sites of DNA damage, where it modulates H3K18Ac levels. H3K18Ac in turn affects recruitment of the damage response factor 53BP1 to DNA double-strand breaks (DSBs), thereby influencing the efficiency of non-homologous end joining (NHEJ). These results reveal a direct role for SIRT7 in DSB repair and establish a functional link between SIRT7-mediated H3K18 deacetylation and the maintenance of genome integrity. Synopsis Sirtuins are deacetylase enzymes implicated in genome stability and life span regulation. Here, the roles of SIRT7 in genome function are studied by characterization of SirT7−/− mice. Loss of SIRT7 results in reduced embryonic viability, accelerated aging, and genomic instability. SIRT7 is recruited to sites of DNA damage in a PARP-dependent manner. SIRT7 participates in the DNA damage response by promoting NHEJ DNA repair. SIRT7-dependent H3K18 deacetylation is important for efficient 53BP1 recruitment at DNA damage sites. SIRT7-deficient cells show increased levels of replication stress, which may underlie progeroid phenotypes. Introduction Sirtuins are NAD+-dependent protein deacetylases and, in some cases, NAD+-dependent ADP ribosyltransferases. Mammals have seven sirtuin family members, which are denoted SIRT1 to 7. They are involved in sensing and responding to different types of cellular stressors, including fasting, genotoxic, and oxidative stress (Vaquero & Reinberg, 2009). A key function of sirtuins is the regulation and maintenance of genome stability under stress. As this regulation fails, genome integrity can diminish, resulting in devastating consequences on cellular fitness, cumulatively leading to organismal aging. Indeed, numerous studies from yeast to mice support a role for sirtuins in the amelioration of human aging-related pathologies (Guarente, 2013), and its deletion is associated with genome instability and compromised organismal viability (Cheng et al, 2003; McBurney et al, 2003; Mostoslavsky et al, 2006; Wang et al, 2008a; Serrano et al, 2013). Deficiency of SIRT7 has been linked to several pathologies including cardiac hypertrophy (Vakhrusheva et al, 2008b; Ryu et al, 2014), hepatic steatosis (Shin et al, 2013; Ryu et al, 2014), and deafness (Ryu et al, 2014). SIRT7 has been functionally linked to transcriptional regulation. SIRT7 is detected at promoters and coding regions of ribosomal genes, where it positively controls ribosome production through direct interaction with the PolI machinery (Ford et al, 2006; Grob et al, 2009; Chen et al, 2013). Conversely, SIRT7 negatively regulates the transcription of genes outside of the rDNA repeats via histone H3K18 deacetylation (Barber et al, 2012). Recent evidence indicates that SIRT7 may have the capacity to act as an oncogene as its expression is elevated in several human cancers (Roth & Chen, 2014). The oncogenic potential of SIRT7 has been attributed to the transcriptional regulation of a specific set of genes through direct interaction with the EKL4 transcription factor. However, whether or not the oncogenic potential of SIRT7 is due to the promotion of genome instability remains heretofore unexplored. Here, we investigate the role of SIRT7 in the maintenance of genome integrity by characterizing SirT7−/− SIRT7-knockout (KO) mice. This mouse model has been shown previously to develop fatty liver due to transcriptional-related ER stress (Shin et al, 2013). Our results show that these mice also suffer from reduced embryonic viability and the development of a progeroid-like phenotype in mice that survive to adulthood. Moreover, SIRT7-deficient cells present replication stress and impaired DDR. In addition, we show that SIRT7 deletion correlates with increased H3K18 acetylation at DNA damage sites and impaired NHEJ repair, revealing a novel molecular mechanism for H3K18Ac in the maintenance of genome integrity, in addition to transcriptional regulation. This report provides new evidence for the role of sirtuins in aging, both by acting as functional mediators between environmental and metabolic factors, and in the regulation of genome stability. Results Perinatal lethality and accelerated aging phenotype in SIRT7-deficient mice We first determined the consequences of SIRT7 loss (Appendix Fig S1A–C) on mouse embryonic development and lifespan. Remarkably, after intercrossing SirT7+/− parental mice, we observed that SirT7−/− pups were born at sub-Mendelian ratios in 129S1/SvImJ mice (9%, down from the predicted 25%, P-value < 0.001 by χ2 test, Fig 1A) and in C57/BL6 SirT7−/− mice (4%, down from the predicted 25%, Appendix Fig S1D), indicative of a defect in embryogenesis, which was still observed at gestational day 14.5 but to a substantially reduced extent (20%, down from predicted 25%; Fig 1A). The weight at birth was lower in SirT7−/− mice relative to WT (5.06 ± 0.1 and 6.8 ± 0.04 g, respectively; Fig 1B and C, and Appendix Fig S1E). Over the 26-week period examined, the rate of growth was similar in the two groups (slope); however, they maintained their weight differences over this time frame as indicated by the graph plateaus. Importantly, more than 20% of SirT7−/− mice died within the first month of life, and the remaining KOs died sooner (12–20 months period) than their WT littermates (Fig 1D). Taken together, these results show that SIRT7 depletion is detrimental to lifespan, suggesting a novel role for SIRT7 protein in perinatal development. Figure 1. SirT7−/− mice have increased perinatal lethality and reduced life span Mendelian ratios from SirT7+/− cross (n = 522 pups; n = 96, 14.5 embryos; P-value < 0.001 by χ2). WT and SirT7−/− mice at 10 days (top) and 2 months (bottom) of age. Weight distribution of WT and SirT7−/− mice (n = 3–10 female mice per time point and genotype; P-value = 0.003 by unpaired t-test; mean ± SD). Kaplan–Meier survival curves (n = 170 WT and n = 58 SirT7−/− mice; log-rank test P-value < 0.0001). Download figure Download PowerPoint Adult SirT7−/− mice showed phenotypic and molecular signs of accelerated aging such as premature (6 months) and pronounced curvature of the spine, kyphosis (100% penetrance; Fig 2A), and decreased gonadal fat pad content (Fig 2B and C). Nevertheless, SirT7−/− mice presented increased hepatic lipid content (Fig EV1A), as previously documented (Shin et al, 2013). In addition, 14 month-old SirT7−/− mice had reduced IGF-1 levels in plasma compared with WT littermates (Fig 2D), as has been observed in different human and mouse progeroid syndromes (Niedernhofer et al, 2006; Murga et al, 2009). Figure 2. Accelerated aging phenotype in SirT7−/− mice A. Representative 3D reconstructed CT scans showing increased kyphosis in 16-month-old SirT7−/− mice compared with WT. B. Representative gonadal fat pads from WT and SirT7−/− 16-month-old mice. C. Quantitation of body weight (left) and gonadal fat pad mass normalized to total body weight (right) (mean ± SEM; four samples per genotype). D. IGF-1 protein levels in serum measured by ELISA (mean ± SEM; 3–8 mice per genotype and age-group). E, F. Dot plots of Lin−Sca1+cKit+ (LSK) cells (middle and right) from WT and SirT7−/− bone marrow cells. Cells were gated for negative staining of lineage markers B220, CD3, CD11b, CD19, Gr-1, and Ter-119 (left), and analyzed for Sca1 and cKit expression (middle and right). (F) Quantitation of (E) (mean ± SEM; 4 mice per genotype). G. Bone marrow, thymus, and spleen cell number in young WT and SirT7−/− mice (mean ± SEM; 3–5 mice per genotype). H. mRNA expression of p16 gene normalized to GAPDH measured by RT–PCR from young and old WT and SirT7−/− fibroblasts (mean ± SEM; three samples per genotype). I, J. Senescence-associated β-galactosidase staining (I, blue) in HT1080 cells transfected with scramble control or SirT7 knockdown and grown for 7 days in selection media. (J) Quantitation of the number of senescent cells shown in (I) (mean ± SEM; three independent cell lines per genotype). Data information: *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001 by ANOVA single factor. Download figure Download PowerPoint Click here to expand this figure. Figure EV1. Accelerated aging phenotype in SirT7−/− mice Representative image of liver cryosections from 3-month-old WT and SirT7−/− mice stained with Oil Red O (red) and counterstained with hematoxylin (blue) (n = 3 livers per genotype). Scale bar 20 μm. Schematic representation of competitive bone marrow transplants, where equal numbers of WT or SirT7−/− bone marrow cells (CD45.2) were mixed with WT competitor cells (CD45.1) and transferred into lethally irradiated recipients via injection into the tail vein. Graph showing the reconstitution of the lymphoid compartment evaluated in peripheral blood by FACS using the CD45.2 marker 8 weeks later (mean ± SEM; 5 mice per genotype from two independent experiments). *P < 0.05; **P < 0.01; by ANOVA single factor. Download figure Download PowerPoint Aging is also associated with stem cell dysfunction (Sharpless & DePinho, 2007). In the bone marrow hematopoietic stem cell LSK (Lineage−, Sca1+, cKit+) population, aging is associated with increased cellularity, which is believed to compensate for the lack of regenerative potential as these cells age (Sudo et al, 2000). The fraction of LSK-positive cells was increased in young (4-month-old) SirT7−/− bone marrow compared with WT littermates (Fig 2E and F). However, the capacity of tissue regeneration of this cell population was reduced as measured by competitive bone marrow transplantation experiments (Fig EV1B and C). WT or SirT7−/− bone marrow-derived cells (CD45.2) were mixed with WT cells (CD45.1) in a 1:1 ratio and were used to reconstitute lethally irradiated mice (Fig EV1B). We found that SIRT7-deficient cells had a reduced (~50%) capacity to repopulate the lymphoid compartment compared with WT bone marrow cells (Fig EV1C). Moreover, analysis of primary and peripheral lymphoid organs revealed leukopenia (Fig 2G), as has been reported in other progeroid mouse models including SirT6−/− mice (Mostoslavsky et al, 2006; Murga et al, 2009). In addition, we measured p16INK4 mRNA levels in splenocytes and ear fibroblasts at 4 and 14 months (Fig 2H, fibroblast data not shown). p16INK4 is a cell cycle regulator that limits the regenerative capacity of many tissues (Krishnamurthy et al, 2004). p16INK4 transcript levels increased as the mice aged and were higher in SirT7−/− mice compared with WT. This difference was markedly greater in younger animals, suggesting premature cellular senescence in SirT7−/− mice. SIRT7 participates in the maintenance of genome integrity Despite the fact that we observed signs of premature senescence in SirT7−/−-derived cells (Fig 2H), we did not observe major differences in the cell growth (Appendix Fig S2A) and cell cycle profiles (Appendix Fig S2B) between SirT7−/− and WT MEFs, in agreement with previous reports (Vakhrusheva et al, 2008a). Remarkably, at later cell passages, we found a mild G2/M arrest and a twofold increase in both polyploid and apoptotic cells (sub-G1) in SirT7−/− MEFs compared with WT (Fig 3A and B). Consistently, SirT7−/− thymocytes had an increased susceptibility to apoptosis after exposure to different doses of X-ray irradiation (IR) compared with WT (Fig 3C), indicative of sustained DNA damage after IR. We further analyzed cell viability by performing colony formation assays after different X-ray doses in HT1080 cells that had been transfected with either scramble (Scr) or SirT7 shRNA. SIRT7-depleted cells formed fewer colonies compared with control cells, once more indicating that SIRT7-deficient cells are more sensitive to externally induced DNA damage (Figs 3D and EV3D). In agreement with the increase in the senescence marker p16INK4, knockdown (KD) of SIRT7 in HT1080 cells resulted in increased number of cells positive for the senescence-associated β-galactosidase marker (Fig 2I and J, and Appendix Fig S6A). Figure 3. Increased genome instability in SirT7−/− mice A, B. Dot plots of FACS cell cycle analyses of WT and SirT7−/− MEFs in passages 3 (P3) and 6 (P6) using EdU incorporation and 7AAD (A). Percentages of cells in sub-G1 (apoptotic cells, left square) and cells with DNA content above 4N (polyploid, right square). (B) Quantitation of experiment shown in (A) (mean ± SEM; three samples per genotype). C. Survival curve for WT and SirT7−/− thymocytes after X-ray irradiation (IR) at the indicated doses. Cell death was quantified by FACS using Annexin V and 7AAD staining 18 h postinsult (mean ± SEM; three samples per genotype from one of two independent experiments). D. Clonogenic assays in HT1080 cells transfected with scramble control or SirT7 knockdown and irradiated at the indicated X-ray doses, then plated at low density. After 9 days, colonies were stained with crystal violet and counted (mean ± SEM; three independent cell lines per genotype). E. Schematic describing the mouse APRT loss of heterozygosity (LOH) assays. Aprt+/− mice were used to measure the in vivo somatic mutation frequency in WT and SirT7−/− mice. Cells fully deficient for APRT are selected in culture by 2,6-diaminopurine (DAP), an adenine analog that is converted to a toxic product by APRT enzymatic activity. Mutant frequency is proportional to the number of DAP-resistant (DAPr) colonies. F. Quantitation of experiment shown in E (mean ± SEM; 9 WT and 3 SirT7−/− samples). Data information: *P < 0.05; **P < 0.01; ***P < 0.001 by ANOVA single factor. Download figure Download PowerPoint We proceeded to examine genome stability in the absence of SIRT7 at the organismal level. We investigated mutation burden in SirT7−/− mice by performing an in vivo mutagenesis assay using the adenine phosphoribosyltransferase (APRT) mouse model, a unique system designed to select for loss of heterozygosity in vivo (Shao et al, 1999) (Fig 3E). Mutant frequency was increased sixfold in SirT7−/− mice compared with WT (Fig 3F), indicating that SIRT7 deficiency increases functional loss of heterozygosity in vivo. Overall, these results reveal a functional link of SIRT7 to the maintenance of genome integrity, which might be a consequence of the accumulation of genomic damage. Indeed, we observed increased levels of DNA damage in SIRT7-deficient cells by comet assays (Fig 4A and Appendix Fig S3A). We next questioned whether the DNA damage might be due to impaired DDR, by monitoring the activity of ataxia telangiectasia mutated (ATM), an apical player of DDR (Shiloh & Ziv, 2013), and its downstream effector KRAB-associated protein 1 (KAP-1) (Ziv et al, 2006) (Fig 4B). The phosphorylation levels of ATM and target proteins increased upon induced DNA damage, and the induction was more elevated in SirT7−/− cells. Overall, these results indicate proper DDR in the absence of SIRT7, but confirm the increased DNA damage observed in SirT7−/− cells. This observation was further examined by measuring the double-strand break (DSB) marker and ATM-targeted protein, γH2AX (Lobrich et al, 2010), by immunofluorescence (IF). We performed a quantitative analysis of the spatial and cell cycle distribution of γH2AX foci in exponentially growing primary mouse fibroblasts (Fig 4C and D). Sites of active DNA replication during S-phase progression were identified by the incorporation of EdU, and G1 and G2 cells were segmented according to their nuclear volume based on DAPI staining of 3D-reconstructed individual nuclei (Serrano et al, 2011) (Appendix Fig S4A). We found a higher number of γH2AX foci throughout the cell cycle in SIRT7-deficient than in WT cells (Fig 4C and D, NoIR). The temporal dynamics of repair, as measured by the time required for a reduction in foci to non-irradiated levels, was similar in the two cell types (Fig 4D, from NoIR to IR-8 h). Eight hours after IR, SIRT7-depleted cells showed increased number of γH2AX foci per nucleus compared with WT, to a similar extent as was observed before IR. We next explored whether DNA damage was associated with a specific type of chromatin by analyzing the nuclear localization of γH2AX foci. We enumerated the number of γH2AX foci overlapping or within the periphery of pericentric heterochromatin (HC) assessed by DAPI staining (Fig 4E and F, G2 is shown; Appendix Fig S4B). Euchromatic numbers were estimated by subtracting the heterochromatic from the total foci number. In agreement with previous reports (Goodarzi et al, 2010), euchromatin- and heterochromatin-associated DSBs were repaired with fast and slow kinetics, respectively (Fig 4E). The number of DSBs associated with pericentric heterochromatin regions was similar between WT and SirT7−/− cells before and at all time points after IR. Figure 4. SIRT7 protects cells from endogenous and induced DNA damage A. Quantitation of neutral comet assays, using passage 3 WT and SirT7−/− primary MEFs, showing (left) the amount (%) of DNA in the tail and (right) the tail moment (see Fig S3A for representative images). B. Western blot showing ATM and KAP1 phosphorylation and total protein levels in WT and SirT7−/− primary fibroblasts after IR (8 Gy). ATM inhibitor (ATMi) KU-55933 was added 30 min prior to irradiation where indicated. One representative blot from four independent experiments is shown. C–F. IF analysis of WT and SirT7−/− primary fibroblasts showing γH2AX dynamics after DNA damage induction. Cells were untreated (NoIR) or treated with 1 Gy of X-rays (IR) and fixed at the indicated times postinsult. Cells were pulsed with EdU (green) 30 min prior to fixation, then stained for γH2AX (red), and counterstained with DAPI (blue) (n > 30 cells per group/mice; 3 mice per genotype). (C) Representative images of untreated S-phase (NoIR, EdU positive) WT and SirT7−/− nuclei (scale bar 5 μm). (D) Quantitation of the number of γH2AX foci per nucleus at the indicated time points post-IR (1 Gy) and indicated cell cycle phases (mean ± SEM from three independent experiments). (E) Quantitation of experiment described in (C) showing mean number of γH2AX foci per nucleus in euchromatic and heterochromatic regions at the indicated time points before and after IR in G2 (mean ± SEM from three independent experiments). Total γH2AX foci and γH2AX foci overlapping with or at the periphery of heterochromatic regions were enumerated. Nuclei and pericentric heterochromatin (chromocenters) were segmented by DAPI staining. Euchromatic numbers were estimated by subtracting the heterochromatic number of foci from the total foci number. (F) Representative images of WT and SirT7−/− primary fibroblasts in G2-phase showing γH2AX (red) and DAPI (blue) at the indicated period of time after IR. (Right) 3D rendering of the IF segmentation depicting nuclei (pale blue), chromocenters (darker blue), and γH2AX (yellow denotes foci associated with pericentric heterochromatin, otherwise foci are red). Scale bar 2 μm. G. FACS quantitation of WT and SirT7−/− MEF cells in S-phase after insult with 10 mM hydroxyurea (HU) for 24 h. Cells were fixed and stained with 7AAD and cell cycle was monitored by FACS (mean ± SEM; five samples per genotype). H, I. DNA fiber labeling analysis was used to assess DNA replication fork progression in passage 3 and passage 6 primary WT and SirT7−/− MEFs. (H) Representative images from cells labeled for 20 min with IdU (green) followed by 20 min of CldU (red). (I) Quantitation of fork velocity (fiber length/labeling time; mean ± SEM; three samples per genotype per condition). Scale bar 10 μm. Data information: *P < 0.05; **P < 0.01; ***P < 0.001 by ANOVA single factor. Source data are available online for this figure. Source Data for Figure 4 [embj201593499-sup-0004-SDataFig4.pdf] Download figure Download PowerPoint We next questioned whether SIRT7 deletion leads to replication-associated defects. To test this possibility, WT and SirT7−/− MEFs were treated with hydroxyurea (HU) to promote replication fork stalling and DSB formation during S-phase. At 24 h after HU treatment, we observed in both cell types that HU led to S-phase arrest. However, this was more prominent in SIRT7-deficient cells compared with WT cells (Fig 4G, HU), suggesting that SIRT7-deficient cells are more prone to replication-associated stress. To directly examine this possibility, DNA fiber labeling analysis was used to assess DNA replication fork progression in WT and SirT7−/− primary MEFs. Loss of SIRT7 resulted in a significant reduction in DNA replication fork velocity in SirT7−/− primary MEFs as compared with WT, which became exacerbated as cells were kept in culture (Fig 4H and I). Reduced replication rate often correlates with increased activation of dormant origins of replication, which is believe to be an attempt to rescue global replication rate after forks collapsed (Zhong et al, 2013). Consistently, we observed a remarkable increase in the presence of stalled replication forks and in the firing of new origins of replication upon replication block with HU in SirT7−/− primary MEFs as compared with WT (Fig EV2A–C). Click here to expand this figure. Figure EV2. Increased replication fork stall and new origin firing in SirT7−/− MEFs Schematic representation of labeling protocol used in (B, C). Cells were pulse-labeled with CldU (red) for 20 min, treated with 2 mM HU for 2 h, and released into media containing IdU (green) for 1 h. Quantitation of the experiment depicted in (A, C), using WT and SirT7−/− primary MEFs in passage 3. Data represent the relative number of stalled forks (CIdU only, red), and new origins (IdU only, green) from the total number of replication tracks labeled with CldU (mean ± SEM; three samples per genotype). *P < 0.05; **P < 0.01; ***P < 0.001 by ANOVA single factor. Representative images from the experiment described in (A) and quantified in (B). Scale bar 10 μm. Download figure Download PowerPoint Overall, our results show that SIRT7 deficiency leads to replication stress, which has an important impact on genome stability and could contribute to explain the observed progeroid phenotype. Impaired double-strand break repair in the absence of SIRT7 Sirtuins participate in the DNA damage response (DDR) by regulating cell cycle progression and DNA repair, particularly non-homologous end joining (NHEJ) and homologous recombination (HR), which repair DNA DSBs (Jeong et al, 2007; Yuan et al, 2007; Li et al, 2008; McCord et al, 2009; Kaidi et al, 2010; Serrano et al, 2013; Toiber et al, 2013). Similarly, SIRT7 may participate in the maintenance of genome integrity by modulating DSB repair. To test this, we analyzed 53BP1 chromatin focus formation by IF (Fig 5A). 53BP1 is a repair protein that promotes NHEJ by protecting DNA from end resection (Panier & Boulton, 2014). We observed a remarkably reduced number of 53BP1 foci per nucleus in the absence of SIRT7 before (Fig EV3A) and after inducing DNA damage by IR (Figs 5B and EV3B). Total 53BP1 protein levels were similar between WT and SirT7−/− cells (Appendix Fig S3B), suggesting that SIRT7 depletion specifically impacts 53BP1 binding to chromatin. Noticeably, the mean volume of 53BP1 foci per nucleus was reduced in cells from SirT7−/− mice compared with WT (Fig 5C). Although 53BP1 facilitation of NHEJ, to the detriment of HR, is plausibly being exerted throughout the cell cycle, 53BP1 also participates in HR-mediated repair of heterochromatin, which is repaired with slow kinetics as compared with euchromatin (Murray et al, 2012). However, our data do not support the notion that SIRT7 regulation of 53BP1 recruitment to damaged chromatin affects heterochromatin repair. We did not observe slower repair kinetics of IR-induced DSBs in SIRT7-depleted cells (Fig 4D), or an accumulation of DNA damage at pericentric heterochromatin (Fig 4E and F), as we would expect for a heterochromatin repair impairment. However, we cannot discard that SIRT7 participates in the repair of heterochromatin out of the pericentric regio