Human CST complex protects stalled replication forks by directly blocking MRE11 degradation of nascent‐strand DNA
2020; Springer Nature; Volume: 40; Issue: 2 Linguagem: Inglês
10.15252/embj.2019103654
ISSN1460-2075
AutoresXinxing Lyu, Kai‐Hang Lei, Pau Biak Sang, Olga Shiva, Megan Chastain, Peter Chi, Weihang Chai,
Tópico(s)Genomics and Chromatin Dynamics
ResumoArticle19 November 2020free access Source DataTransparent process Human CST complex protects stalled replication forks by directly blocking MRE11 degradation of nascent-strand DNA Xinxing Lyu Xinxing Lyu Department of Cancer Biology, Cardinal Bernardin Cancer Center, Loyola University Chicago Stritch School of Medicine, Maywood, IL, USA Department of Biomedical Sciences, ESF College of Medicine, Washington State University, Spokane, WA, USA Search for more papers by this author Kai-Hang Lei Kai-Hang Lei orcid.org/0000-0002-8783-5806 Institute of Biochemical Sciences, National Taiwan University, Taipei, Taiwan Search for more papers by this author Pau Biak Sang Pau Biak Sang Department of Cancer Biology, Cardinal Bernardin Cancer Center, Loyola University Chicago Stritch School of Medicine, Maywood, IL, USA Search for more papers by this author Olga Shiva Olga Shiva Department of Biomedical Sciences, ESF College of Medicine, Washington State University, Spokane, WA, USA Search for more papers by this author Megan Chastain Megan Chastain Department of Biomedical Sciences, ESF College of Medicine, Washington State University, Spokane, WA, USA Search for more papers by this author Peter Chi Corresponding Author Peter Chi [email protected] orcid.org/0000-0001-9229-8729 Institute of Biochemical Sciences, National Taiwan University, Taipei, Taiwan Institute of Biological Chemistry, Academia Sinica, Taipei, Taiwan Search for more papers by this author Weihang Chai Corresponding Author Weihang Chai [email protected] orcid.org/0000-0003-1206-2324 Department of Cancer Biology, Cardinal Bernardin Cancer Center, Loyola University Chicago Stritch School of Medicine, Maywood, IL, USA Search for more papers by this author Xinxing Lyu Xinxing Lyu Department of Cancer Biology, Cardinal Bernardin Cancer Center, Loyola University Chicago Stritch School of Medicine, Maywood, IL, USA Department of Biomedical Sciences, ESF College of Medicine, Washington State University, Spokane, WA, USA Search for more papers by this author Kai-Hang Lei Kai-Hang Lei orcid.org/0000-0002-8783-5806 Institute of Biochemical Sciences, National Taiwan University, Taipei, Taiwan Search for more papers by this author Pau Biak Sang Pau Biak Sang Department of Cancer Biology, Cardinal Bernardin Cancer Center, Loyola University Chicago Stritch School of Medicine, Maywood, IL, USA Search for more papers by this author Olga Shiva Olga Shiva Department of Biomedical Sciences, ESF College of Medicine, Washington State University, Spokane, WA, USA Search for more papers by this author Megan Chastain Megan Chastain Department of Biomedical Sciences, ESF College of Medicine, Washington State University, Spokane, WA, USA Search for more papers by this author Peter Chi Corresponding Author Peter Chi [email protected] orcid.org/0000-0001-9229-8729 Institute of Biochemical Sciences, National Taiwan University, Taipei, Taiwan Institute of Biological Chemistry, Academia Sinica, Taipei, Taiwan Search for more papers by this author Weihang Chai Corresponding Author Weihang Chai [email protected] orcid.org/0000-0003-1206-2324 Department of Cancer Biology, Cardinal Bernardin Cancer Center, Loyola University Chicago Stritch School of Medicine, Maywood, IL, USA Search for more papers by this author Author Information Xinxing Lyu1,2, Kai-Hang Lei3, Pau Biak Sang1, Olga Shiva2, Megan Chastain2, Peter Chi *,3,4 and Weihang Chai *,1 1Department of Cancer Biology, Cardinal Bernardin Cancer Center, Loyola University Chicago Stritch School of Medicine, Maywood, IL, USA 2Department of Biomedical Sciences, ESF College of Medicine, Washington State University, Spokane, WA, USA 3Institute of Biochemical Sciences, National Taiwan University, Taipei, Taiwan 4Institute of Biological Chemistry, Academia Sinica, Taipei, Taiwan *Corresponding author. Tel: +886 2 33664066, Fax: +886 2 23635038; E-mail: [email protected] *Corresponding author. Tel: +708 327 3298; Fax: 708 327 3342; E-mail: [email protected] The EMBO Journal (2021)40:e103654https://doi.org/10.15252/embj.2019103654 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 Degradation and collapse of stalled replication forks are main sources of genomic instability, yet the molecular mechanisms for protecting forks from degradation/collapse are not well understood. Here, we report that human CST (CTC1-STN1-TEN1) proteins, which form a single-stranded DNA-binding complex, localize at stalled forks and protect stalled forks from degradation by the MRE11 nuclease. CST deficiency increases MRE11 binding to stalled forks, leading to nascent-strand degradation at reversed forks and ssDNA accumulation. In addition, purified CST complex binds to 5’ DNA overhangs and directly blocks MRE11 degradation in vitro, and the DNA-binding ability of CST is required for blocking MRE11-mediated nascent-strand degradation. Our results suggest that CST inhibits MRE11 binding to reversed forks, thus antagonizing excessive nascent-strand degradation. Finally, we uncover that CST complex inactivation exacerbates genome instability in BRCA2 deficient cells. Collectively, our findings identify the CST complex as an important fork protector that preserves genome integrity under replication perturbation. Synopsis The human CTC1-STN1-TEN1 (CST) complex binds to ssDNA and has been implicated in protecting genomic stability under replication stress. This study shows that the CST complex localizes to replication forks in response to their stalling, and protects them from degradation by MRE11 nuclease. Human CST proteins localizes at stalled replication forks. CST deficiency increases MRE11 association with stalled forks and MRE11-mediated degradation of nascent-strand DNA at reversed forks, leading to genome instability. The CST complex binds to ds/ssDNA substrates and directly blocks MRE11 degradation of DNA in a sequence-independent manner. The DNA binding activity of CST is a prerequisite for blocking MRE11 degradation at reversed forks. CST inactivation exacerbates genome instability in BRCA2-deficient cells. Introduction Faithful DNA replication is fundamental to genome integrity maintenance. During the process of DNA replication, replication machinery may encounter obstacles that cause replication forks slowing or stalling, leading to replication stress. Defects in stalled fork restart or stabilization induce genome instability, which is the main source of the pathology of human genetic diseases including cancer, aging, neurological diseases, and developmental defects (Zeman & Cimprich, 2013; Mazouzi et al, 2014). A panoply of mechanisms protects genome stability in response to fork stalling to guard normal cell proliferation. Slowing or stalled replication generates extensive ssDNA stretches at forks, which are bound by the replication protein A (RPA) complex. RPA binding to ssDNA activates the ataxia telangiectasia and Rad3-related (ATR) kinase and recruits numerous proteins to facilitate the protection and restart of stalled forks (Flynn & Zou, 2011; Marechal & Zou, 2015). Multiple pathways, including dormant origin firing, replication repriming, translesion synthesis, template switching, and fork reversal, act to facilitate the protection and restart of the stalled forks (Zeman & Cimprich, 2013; Bhat & Cortez, 2018). Electron microscopy analysis has revealed that various replication stressors can trigger fork remodeling to form a chicken foot-like physical structure through fork regression (Zellweger et al., 2015; Kolinjivadi et al., 2017). This fork reversal mechanism has emerged as an important means for stabilizing stalled forks and resuming replication following replication perturbation. For this reason, the dynamics of reversed forks has been extensively studied (Neelsen & Lopes, 2015; Quinet et al., 2017b; Rickman & Smogorzewska, 2019). The formation of reversed forks is catalyzed by SMARCAL1, ZRANB3, FANCM, HLTF, FBH1 (Gari et al., 2008; Betous et al., 2012; Kile et al., 2015; Kolinjivadi et al., 2017; Taglialatela et al., 2017; Vujanovic et al., 2017) and also involves RAD51 (Zellweger et al., 2015). While fork reversal is a protective mechanism to preserve fork stability under replication stress, the regressed arm also provides a target for nuclease-dependent end resection. Without fork protectors such as BRCA1/2, RAD51, BARD1, and Abro1, reversed forks become vulnerable to nucleases MRE11, EXO1, DNA2, CtIP (Ying et al., 2012; Thangavel et al., 2015; Kolinjivadi et al., 2017; Lemacon et al., 2017; Mijic et al., 2017; Xu et al., 2017; Billing et al., 2018). Uncontrolled fork degradation by these nucleases is detrimental to genome stability and has been linked to both the lethality of BRCA2-defective embryonic stem cells and the sensitivity of BRCA-defective cells to certain chemotherapeutic treatments (Schlacher et al., 2011; Ying et al., 2012; Ray Chaudhuri et al., 2016). To date, the molecular mechanism underlying the balance between the protection and resection of regressed arms remains unclear. The human CST complex, comprising of CTC1, STN1, and TEN1, contains multiple OB- fold domains and resembles the RPA70-RPA32-RPA14 complex structure (Miyake et al., 2009). CST prefers binding to G-rich sequences in vitro in a manner dependent on the sequence length (Miyake et al., 2009; Chen et al., 2012). It is capable of binding to an 18-nt G-rich ssDNA with high affinity, but the sequence specificity is lost if the oligonucleotide becomes longer (Miyake et al., 2009; Hom & Wuttke, 2017). Interestingly, although CST is unable to bind to dsDNA, ss/dsDNA junctions stabilize CST binding and decrease minimal ssDNA length requirement for CST binding to 10 nt (Bhattacharjee et al., 2017), indicating that CST may bind to special DNA structures with ss/dsDNA junctions in vivo. A set of missense mutations of CTC1 and STN1 genes have been identified in patients with the Coats plus disease, a rare autosomal recessive disorder characterized by bilateral exudative retinopathy, retinal telangiectasias, growth retardation, intracranial calcifications, bone abnormalities, gastrointestinal vascular ectasias, accompanied by common early-aging pathological features (Anderson et al., 2012; Keller et al., 2012; Simon et al., 2016). Like its homolog in budding yeast (Cdc13-Stn1-Ten1), the well-characterized and conserved roles of human/mammalian CST are in preserving telomere integrity. CST promotes efficient and complete replication of telomeric DNA to prevent sudden loss of telomeres (Huang et al., 2012; Gu & Chang, 2013). In addition, CST stimulates the priming activity of DNA polymerase α (POLα)-primase (Lue et al., 2014; Ganduri & Lue, 2017) and mediates C-strand fill-in at telomere ends to replenish resected C-strands (Huang et al., 2012). CST also inhibits telomerase access to telomeres and coordinates G- and C-strand synthesis (Chen et al., 2012). Recently, a number of telomere-independent roles of CST have been uncovered. As a downstream effector in the 53BP1 pathway, CST participates in double-strand break (DSB) repair in a Shieldin-dependent manner (Barazas et al., 2018; Mirman et al., 2018). CST also plays a role in genome-wide replication. During normal replication, CST directly associates with the MCM complex and blocks MCM binding to CDT1 to decrease origin licensing in G1-phase. In addition, CST facilitates origin firing by promoting AND-1 and POLα chromatin association in S-phase (Wang et al., 2019). Previously, we have shown that CST associates with G- or C-rich repetitive non-telomeric sequences in response to fork stalling, probably by binding to ssDNA accumulated at stalled forks (Chastain et al., 2016). CST facilitates fork recovery and its deficiency induces instabilities of these sequences and chromosome fragmentation, indicating that CST is an important player for protecting genome stability under replication stress (Stewart et al., 2012; Chastain et al., 2016). However, the precise molecular mechanism underlying how CST participates in rescuing stalled replication remains unclear. In this study, we report that CST is located at stalled forks upon hydroxyurea (HU) treatment, and CST deficiency leads to ssDNA accumulation under replication stress. DNA fiber analysis demonstrates that CST depletion induces MRE11 degradation of nascent-strand DNA that is dependent on fork reversal, suggesting that CST antagonizes nascent DNA degradation at reversed forks. Using purified human CST and human MRE11 proteins, we find that CST binds to the ss/dsDNA structure that mimics the reversed fork, directly blocking MRE11 degradation in vitro. Consistent with our in vitro data, we observe that MRE11 binding to stalled forks is increased upon CST depletion. The fork protection function of CST requires its binding to DNA, and loss of CST binding to DNA leads to nascent-strand degradation in cells and in vitro. In addition, we uncover that CST inactivation exacerbates genome instability in BRCA2 deficient cells. Collectively, our results suggest that CST binds to reversed forks to inhibit MRE11 access to nascent-strand DNA, thus acting as a direct fork protector to maintain genome stability in response to replication stress. Results CST localizes at stalled forks We have previously shown that in addition to promoting efficient replication of telomeric DNA, CST is also enriched at G- or C-rich repetitive sequences genome-wide after replication fork stalling to protect the stability of these sequences (Chastain et al., 2016). However, it is unknown whether such protective function is direct or indirect due to the lack of explicit evidence supporting CST localization at stalled forks. To gain insight into the function of CST in protecting genome stability, we examined the localization of endogenous CST proteins at stalled forks in response to replication stress using the in situ protein interactions at nascent and stalled replication forks (SIRF) assay, which offers sensitive visualization of protein localization at forks at a single-cell level if the protein-of-interest is in close proximity to EdU-labeled nascent strands (Roy et al., 2018). Briefly, nascent DNA in proliferating cells is labeled with EdU, and the incorporated EdU is then biotinylated by the click reaction. Subsequently, the proximity ligation assay (PLA) is used to visualize the colocalization of the target protein with biotinylated EdU (Fig 1A). To ensure the results were not cell line specific, we performed SIRF in two cell lines U2OS and HCT116. We pulse-labeled cells with EdU for 8 min, treated them with HU for 3 h and then carried out SIRF analysis using anti-CTC1 or anti-STN1 antibody (Fig 1B and C). Since EdU incorporation efficiency might be affected by HU treatment (Roy et al., 2018), we performed the EdU SIRF as previously described (Roy et al., 2018). In agreement with previous report (Roy et al., 2018), we observed slight increase of EdU-SIRF signal after HU treatment (Appendix Fig S1A). Thus, we normalized CTC1/STN1-SIRF signals to mean EdU-SIRF signals before statistical analysis (Fig 1B and C, and Appendix Fig S1A). Very few SIRF positive cells were observed without of EdU, suggesting that SIRF assay was specific for determining protein localization at forks (Fig 1B and C and Appendix Fig S1B). To validate that the antibody specifically recognized the endogenous CTC1 protein, we performed CTC1 SIRF in knockdown cells. Depletion of CTC1 resulted in the dramatic reduction of CTC1 SIRF foci (Appendix Fig S1F), suggesting the observed SIRF signal primarily resulted from antibody specific recognition of the CTC1 protein. As control, we performed SIRF against the DNA polymerase processivity factor proliferating cell nuclear antigen (PCNA). As expected, PCNA localized at active forks and formed distinct SIRF signals in untreated replicating cells (Fig 1B and C, Appendix Fig S1B). We also observed decreased PCNA SIRF foci after HU-induced fork stalling (Fig 1B and C), consistent with previous findings from iPOND and SIRF assays that the amount of PCNA at stalled forks reduces when compared to that at elongating forks (Sirbu et al., 2011; Roy et al., 2018). Figure 1. CST localizes at stalled forks A. Scheme of SIRF assay. Nascent DNA was pulse-labeled with EdU. Upon fork stalling, proteins located at stalled forks are in close proximity to EdU. EdU was then biotinylated using click reaction, followed by incubating with biotin and target protein antibodies. PLA amplification was used to visualize the localization of target proteins at EdU-labeled stalled forks. B, C. Detection of CST at stalled and active forks with SIRF assay in U2OS (B) and HCT116 (C) cells. To detect CST at stalled forks, cells were pulse-labeled with EdU for 8 min, followed by 4 mM HU treatment for 3 h. To detect CST at active forks, SIRF was performed without HU treatment. Representative SIRF images of CTC1, STN1, and PCNA on normal or stalled forks are shown. Scale bars: 10 µm. Three independent experiments were performed and scatter plots from one experiment are shown in the main figure. Scatter plots of two additional biological replicates are included in Appendix Fig S1C (U2OS cells) and Appendix Fig S1D and E (HCT116 cells). Relative SIRF fluorescence intensity was calculated by normalizing to untreated samples and then to their respective EdU-SIRF signals. Mean value in each sample is labeled in the graph. N: the number of cells analyzed in each sample. P values were calculated by Mann–Whitney test. ***P < 0.001. Red lines indicate the mean values. Error bars: SEM. Download figure Download PowerPoint In both cell lines, we detected SIRF signals of CTC1 and STN1 in untreated replicating cells, indicating that CST also localizes at unperturbed forks (Fig 1B and C). This is in agreement with previous reports that CST interacts with POLα and facilitates DNA replication (Casteel et al., 2009; Chen et al., 2013; Ganduri & Lue, 2017). SIRF amplification signals from CTC1 and STN1 were increased by HU treatment (Fig 1B and C), suggesting an increased CTC1 and STN1 localization at stalled forks in response to HU treatment. CST deficiency leads to the accumulation of ssDNA and increases MRE11 degradation of nascent-strand DNA at stalled forks The initial outcome of replication stalling is the aberrant ssDNA production at stalled forks, likely caused by the uncoupling of DNA helicase from DNA polymerase machinery. CST is a ssDNA-binding protein complex and previously has been reported to suppress the formation of aberrant ssDNA at telomeres (Huang et al., 2012; Chen et al., 2013). We therefore examined the effect of CST depletion on genomic ssDNA production after fork stalling. HeLa cells depleted of STN1 (Fig 2A) were treated with HU for 3 h, a condition that induced fork stalling and produced ssDNA (Ercilla et al., 2019). Subsequently, BrdU staining was performed under the non-denaturing condition to measure ssDNA amount. As shown in Fig 2B, STN1 depletion increased the amount of ssDNA after HU treatment, suggesting that CST loss leads to aberrant ssDNA accumulation (Fig 2B and Appendix Fig S2A). Figure 2. CST suppresses the accumulation of ssDNA at stalled forks and protects nascent-strand DNA from MRE11 degradation Western blot showing STN1 knock-down. shLUC (siRNA targeting luciferase) was used as the control. CST deficiency causes ssDNA accumulation upon fork stalling. HeLa cells with STN1 knock-down were incubated with 10 µM BrdU for 48 h, followed by 2 mM HU treatment for 3 h, and subsequently stained with BrdU antibody under a non-denaturing condition to detect ssDNA. N, the number of cells analyzed in each condition. Scale bars: 20 µm. Relative BrdU fluorescence intensity was quantitated by Image J. P values were calculated by one-way ANOVA analysis with post hoc Tukey. Error bars, SEM, ***P < 0.001, **P < 0.01. Three independent experiments were performed. Results from two additional biological replicates are included in Appendix Fig S2A. Scheme of DNA fiber analysis and representative fiber images. DNA fiber analysis of U2OS cells with STN1 knock-down and its RNAi-resistant Flag-STN1 WT re-expression with and without mirin treatment. Results from biological replicates and corresponding representative fiber images are included in Appendix Fig S2B. DNA fiber analysis of BJ cells with STN1 knock-down by two siRNA sequence with and without mirin treatment. Results from biological replicates are included in Appendix Fig S2C. DNA fiber analysis of HCT116 cells with STN1 knock-down with and without mirin treatment. Results from biological replicates are included in Appendix Fig S2D. DNA fiber analysis of U2OS cells showing that SMARCAL1-mediated fork reversal is needed for nascent-stand degradation observed in CST-deficient cells. Results from biological replicates are included in Appendix Fig S2E. DNA fiber analysis of U2OS cells showing that ZRANB3-mediated fork reversal is needed for nascent-strand degradation observed in CST-deficient cells. Results from biological replicates are included in Appendix Fig S2F. Data information: For all DNA fiber experiments in this study, at least two independent treatments and fiber experiments were performed to ensure reproducibility. Results from one experiment are shown in the main figure. Scatter plots indicate IdU/CIdU tract length ratios for individual replication forks. N, the number of DNA fibers analyzed in each condition. P values were calculated by one-way ANOVA analysis with post hoc Tukey. ***P < 0.001, **P < 0.01, *P < 0.05. Red lines indicate mean values. Error bars: SEM. Western blots showing protein knock-down in DNA fiber assays are included. Source data are available online for this figure. Source Data for Figure 2 [embj2019103654-sup-0002-SDataFig2.pdf] Download figure Download PowerPoint In response to replication stalling, forks may reverse to stabilize stalled forks and promote restart (Quinet et al., 2017b). Reversed forks, if not protected properly, are attacked by nucleases such as MRE11, DNA2 and EXOI, causing excessive degradation of nascent-strand DNA (Thangavel et al., 2015; Lemacon et al., 2017). The accumulation of ssDNA in CST deficiency cells led us to hypothesize that CST might play a role in protecting nascent-strand DNA from nucleolytic degradation. To test this hypothesis, we analyzed fork stability using DNA fiber assays (Nieminuszczy et al., 2016; Quinet et al., 2017a). Replication tracks were sequentially labeled with chlorodeoxyuridine (CIdU) and iododeoxyuridine (IdU) for 30 min, followed by 4 mM HU treatment for 5 h to induce fork stalling (Fig 2C). Only bi-colored tracks were included in analysis. We found that STN1 depletion in U2OS cells caused increased fork degradation, and RNAi-resistant wild-type STN1 rescued such degradation in knockdown cells (Fig 2D and Appendix Fig S2B). The presence of the MRE11 inhibitor mirin suppressed such shortening, indicating that STN1 depletion leads to unscheduled MRE11-mediated degradation of nascent-strand DNA at stalled forks (Fig 2D). The same phenotype was recapitulated in two other cell lines normal foreskin fibroblast cell line BJ and colon cancer cell line HCT116 (Figs 2E and F, and Appendix Figs S2C and D). Likewise, CTC1 depletion with shRNA caused similar fork degradation (see Fig 5D below). Expression of the RNAi-resistant CTC1 (res-CTC1) cDNA fully rescued nascent-strand shortening in knock-down cells, confirming that the phenotype was caused specifically by knock-down (see Fig 5D below). Figure 5. The ssDNA-binding ability of CST is required for protecting nascent strands from degradation Deletion of the N-terminal 700 aa of CTC1 abolishes CTC1 localization at telomeres. U2OS stably expressing vector (V), WT Myc-CTC1 and Myc-Δ700N were treated with or without HU and co-stained with Myc (red) and TRF2 (green) antibodies. Boxed areas are amplified in inserts to indicate CTC1/TRF2 colocalization (yellow).Scale bars: 10 μm. Quantification of percent of cells with > 3 CTC1/TRF2 colocalization foci was from three independent experiments. In each experiment, > 150 cells from each sample were analyzed. P values were calculated by one-way ANOVA analysis with post hoc Tukey from three independent experiments, ***P < 0.001. Error bars: SEM. Δ700N does not affect CST complex formation. HEK293T cells were co-transfected with Myc- CTC1 or Myc-Δ700N, His6-STN1, and HA-TEN1. Co-IP was performed with Myc antibody to pulldown His6- STN1 and HA-TEN1. Full-length Myc-CTC1 was prone to degradation during immunoprecipitation. Δ700N retains RAD51 interaction. HEK293T cells were co-transfected with Myc-CTC1 or Myc-Δ700N, His6- STN1, HA-TEN1, and Flag-RAD51 or vector control (V) and treated with HU (2 mM, 16 h). Co-IP was performed with Flag antibody to pulldown Myc-CTC1. DNA fiber analysis of CTC1 depleted U2OS cells with RNAi-resistant WT or Δ700N re-expression. Two independent experiments were performed. Results from the biological replicate are included in Appendix Fig S4B. Mean values in each sample are listed at the top of the graph. P values were calculated by one-way ANOVA analysis with post hoc Tukey. N, the number of cells analyzed in each condition, ***P < 0.001. Red lines indicate the mean values. Error bars: SEM. Source data are available online for this figure. Source Data for Figure 5 [embj2019103654-sup-0004-SDataFig5.pdf] Download figure Download PowerPoint Nascent-strand degradation caused by CST deficiency requires fork reversal Fork reversal through remodeling of stalled forks into four-way junction structures is catalyzed by SMARCAL1, ZRANB3, and others (Kolinjivadi et al., 2017; Taglialatela et al., 2017; Vujanovic et al., 2017). To determine whether the fork protection function of CST relied on fork reversal, we knocked down SMARCAL1 or ZRANB3 in STN1-depleted cells and measured nascent-strand degradation. Our results revealed that depletion of SMARCAL1 or ZRANB3 rescued nascent-strand degradation in STN1-deficient cells (Figs 2G and H, and Appendix Figs 2E and F). Consistent with previous reports (Coquel et al., 2018; Tonzi & Yin, 2018), depletion of SMARCAL1 slightly increased IdU/CldU ratio in control knock-down cells (Fig 2G), though the underlying reason is unclear. Regardless, our results suggest that fork reversal is a prerequisite for fork degradation in CST-deficient cells. CST deficiency increases MRE11 association to stalled forks, leading to ssDNA accumulation To determine whether nascent-strand DNA degradation in CST-deficient cells was caused by the accumulation of MRE11 at stalled forks, we then performed MRE11 SIRF. SIRF results revealed that STN1 depletion increased the accumulation of MRE11 at stalled forks (Fig 3A and Appendix Fig S3A). Treating STN1 knock-down cells with mirin rescued ssDNA accumulation (Fig 3B and Appendix Fig S3B), suggesting that the elevated genome instability in CST-deficient cells was likely caused by the unscheduled MRE11 nucleolytic resection of the nascent-strand DNA. Figure 3. CST deficiency increases MRE11 localization at stalled forks, leading to ssDNA accumulation SIRF detection of MRE11 at stalled forks in STN1-deficient U2OS cells. Representative SIRF images of MRE11 at normal or stalled replication forks are shown. Scale bars: 10 µm. Two independent experiments were performed. Results from the biological replicate are shown in Appendix Fig S3A. P values were calculated by one-way ANOVA analysis with post hoc Tukey, ***P < 0.001. Red lines indicate mean values. Error bars: SEM. ssDNA accumulation analysis in STN1-deficient U2OS cells with and without mirin treatment. Representative images of native BrdU staining are shown. Scale bars: 20 µm. Three independent experiments were performed. Results from two additional biological replicates are shown in Appendix Fig S3C. P values were calculated by one-way ANOVA analysis with post hoc Tukey, ***P < 0.001. Red lines indicate mean values. Error bars, SEM. Download figure Download PowerPoint Purified CST binds to 5’ overhang and directly blocks MRE11 degradation of DNA The above observations indicated that CST could protect reversal forks from MRE11 degradation to maintain genome stability. MRE11 possesses the 3’–5’ exonuclease activity and can extensively degrade ss/dsDNA junction structures with 5’ overhangs formed by regressed forks (Kolinjivadi et al., 2017). It has been reported that CST prefers binding to ss/dsDNA junctions in vitro (Bhattacharjee et al., 2017). Hence, we hypothesized that CST might bind to 5’ overhangs of regressed arms to protect nascent-strand DNA from MRE11 degradation. To test this, we purified the human MRE11 protein (Fig 4A), generated a ss/dsDNA junction structure with a 5’ overhang that mimicked the regressed arm (Fig 4B i), and used the purified human heterotrimeric CST complex and MRE11 proteins to reconstitute nascent-strand DNA protection in vitro. Recombinant human MRE11 was purified from Expi293F cells as described in Methods (Fig 4A) and then incubated with the ss/dsDNA substrate. In agreement with the previous report, MRE11-dependent degradation in the 3’ to 5’ direction was observed (Fig 4B ii), and the MRE11 nuclease activity required cofactor Mn2+ (Fig 4B iii, and Appendix Fig S4F) (Paull & Gellert, 1998). Next, we purified the human CST compl
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