Human CST promotes telomere duplex replication and general replication restart after fork stalling
2012; Springer Nature; Volume: 31; Issue: 17 Linguagem: Inglês
10.1038/emboj.2012.215
ISSN1460-2075
AutoresJason A. Stewart, Feng Wang, Mary F. Chaiken, Christopher Kasbek, Paul D. Chastain, Woodring E. Wright, Carolyn M. Price,
Tópico(s)CRISPR and Genetic Engineering
ResumoArticle3 August 2012free access Human CST promotes telomere duplex replication and general replication restart after fork stalling Jason A Stewart Jason A Stewart Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA Search for more papers by this author Feng Wang Feng Wang Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA Search for more papers by this author Mary F Chaiken Mary F Chaiken Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA Search for more papers by this author Christopher Kasbek Christopher Kasbek Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA Search for more papers by this author Paul D Chastain II Paul D Chastain II Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA Search for more papers by this author Woodring E Wright Woodring E Wright Department of Cell Biology, University of Texas Southwestern Medical Center, Dallas, TX, USA Search for more papers by this author Carolyn M Price Corresponding Author Carolyn M Price Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA Search for more papers by this author Jason A Stewart Jason A Stewart Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA Search for more papers by this author Feng Wang Feng Wang Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA Search for more papers by this author Mary F Chaiken Mary F Chaiken Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA Search for more papers by this author Christopher Kasbek Christopher Kasbek Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA Search for more papers by this author Paul D Chastain II Paul D Chastain II Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA Search for more papers by this author Woodring E Wright Woodring E Wright Department of Cell Biology, University of Texas Southwestern Medical Center, Dallas, TX, USA Search for more papers by this author Carolyn M Price Corresponding Author Carolyn M Price Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA Search for more papers by this author Author Information Jason A Stewart1,‡, Feng Wang1,‡, Mary F Chaiken1, Christopher Kasbek1, Paul D Chastain2, Woodring E Wright3 and Carolyn M Price 1 1Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA 2Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA 3Department of Cell Biology, University of Texas Southwestern Medical Center, Dallas, TX, USA ‡These authors contributed equally to this work *Corresponding author. Department of Cancer and Cell Biology, University of Cincinnati, 3125 Eden Avenue, Cincinnati, OH 45267, USA. Tel.:+1 513 558 0450; Fax:+1 513 558 8474; E-mail: [email protected] The EMBO Journal (2012)31:3537-3549https://doi.org/10.1038/emboj.2012.215 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 Mammalian CST (CTC1-STN1-TEN1) associates with telomeres and depletion of CTC1 or STN1 causes telomere defects. However, the function of mammalian CST remains poorly understood. We show here that depletion of CST subunits leads to both telomeric and non-telomeric phenotypes associated with DNA replication defects. Stable knockdown of CTC1 or STN1 increases the incidence of anaphase bridges and multi-telomeric signals, indicating genomic and telomeric instability. STN1 knockdown also delays replication through the telomere indicating a role in replication fork passage through this natural barrier. Furthermore, we find that STN1 plays a novel role in genome-wide replication restart after hydroxyurea (HU)-induced replication fork stalling. STN1 depletion leads to reduced EdU incorporation after HU release. However, most forks rapidly resume replication, indicating replisome integrity is largely intact and STN1 depletion has little effect on fork restart. Instead, STN1 depletion leads to a decrease in new origin firing. Our findings suggest that CST rescues stalled replication forks during conditions of replication stress, such as those found at natural replication barriers, likely by facilitating dormant origin firing. Introduction Mammalian telomeres consist of kilobases of duplex T2AG3/C3TA2 repeats bound by a six-protein complex called shelterin (Palm and de Lange, 2008; Stewart et al, 2012). The G-rich strand ends in a 12–400 nt ssDNA 3′ overhang which is proposed to invade the duplex region to create a telomeric loop (t-loop) that caps the chromosome terminus. Telomeres pose a unique problem for the replication machinery due to their repetitive nature and unusual terminal structure (Gilson and Geli, 2007; Stewart et al, 2012). The duplex region is replicated by the conventional DNA replication machinery. However, the heterochromatic nature of this region and its potential to form secondary structures appear to impede passage of the replication fork (Paeschke et al, 2010). The t-loop may provide an additional barrier to the fork. In human cells that express telomerase, the 3′ overhang is elongated by telomerase soon after passage of the replication fork (Zhao et al, 2009). The extended overhang is then partially filled at the end of S-phase. This C-strand fill-in is a stepwise process and is thought to occur through the recruitment of DNA polymerase α-primase (pol α) (Zhao et al, 2009). Recent findings indicate that a number of extra proteins are needed in addition to the standard replication machinery to properly replicate the telomeric duplex. These include TRF1, FEN1, BLM, RTEL, RECQL4, BRCA2 and RAD51 (Sfeir et al, 2009; Badie et al, 2010; Saharia et al, 2010; Ghosh et al, 2011). Depletion of these proteins causes the appearance of multi-telomeric signals (MTS) during telomere FISH analysis on metaphase chromosomes. MTS have also been called fragile telomeres because they share features of common fragile sites which are observed cytogenetically as gaps or breaks in chromosomes (Durkin and Glover, 2007). While the actual nature of the MTS is poorly understood, like fragile sites, they can form under conditions of replication stress and replication fork stalling (Durkin and Glover, 2007; Chan et al, 2009; Sfeir et al, 2009). Fork stalling can be induced by a number of factors, such as repetitive or complex DNA sequences, depletion of nucleotide pools and DNA damage (Durkin and Glover, 2007; Branzei and Foiani, 2010; Petermann and Helleday, 2010). Once stalled, replication must be rapidly restarted to maintain genome stability. In the absence of restart, replication forks collapse leading to regions of ssDNA, DNA double-strand breaks and unwanted recombination events. The mechanisms underlying replication restart at telomeres, fragile sites and other sites of difficult-to-replicate DNA are not fully understood. The work described here indicates that the recently identified mammalian CTC1-STN1-TEN1 (CST) complex promotes DNA replication restart at both telomeric and non-telomeric sites. Mammalian CST resembles the Cdc13-Stn1-Ten1 complex (ScCST) from Saccharomyces cerevisiae in that the STN1 and TEN1 subunits are conserved, both complexes share a structural similarity to Replication Protein A (RPA) and both bind tightly to ssDNA (Giraud-Panis et al, 2010; Price et al, 2010). ScCST is responsible for protecting yeast telomeres through sequence-specific binding to the G-strand overhang (Pennock et al, 2001; Shore and Bianchi, 2009) and for coordinating G- and C-strand synthesis during telomere replication via interactions with telomerase and pol α (Qi and Zakian, 2000; Chandra et al, 2001; Puglisi et al, 2008). Mammalian CST also localizes to telomeres and its depletion causes changes in telomere structure (see below) indicating a role in telomere maintenance (Miyake et al, 2009; Surovtseva et al, 2009). Unlike its yeast counterpart, mammalian CST does not appear to be required for telomere protection but it may play a similar role in telomere replication. The proposed role for mammalian CST is in the C-strand fill-in that occurs following extension of the 3′ overhang by telomerase (Price et al, 2010). Recent studies show that this fill-in occurs hours after replication of the telomere duplex (Zhao et al, 2009). Since a replisome is unlikely to be present hours after telomeres have replicated and telomerase has acted, CST is proposed to recruit pol α to initiate the fill-in reaction (Zhao et al, 2009). However, several lines of evidence suggested that CST might also have non-telomeric functions. First, CST binding to ssDNA is sequence independent and only ∼20% of STN1 foci localize to telomeres (Miyake et al, 2009). Second, depletion of CTC1 results in increased γH2AX foci, which do not colocalize with telomeres (Miyake et al, 2009; Surovtseva et al, 2009). Finally, CTC1 and STN1 were initially isolated as subunits of a pol α accessory factor (AAF) that increases pol α processivity and affinity for an ssDNA template (Goulian et al, 1990; Casteel et al, 2009). More recent work indicates that Xenopus CST shares many of these same properties (Nakaoka et al, 2012). Interestingly, a new series of clinical studies has identified mutations in the CTC1 subunit of CST as the cause of human disease (Anderson et al, 2012; Keller et al, 2012; Polvi et al, 2012). Initially, CTC1 mutations were shown to underlie Coats plus, a severe pleiotropic disorder with many clinical manifestations ranging from retinal telangiectasia, intracranial calcification with leukodystrophy and brain cysts, to predisposition to fractures and gastrointestinal bleeding. A subset of the clinical features of Coats plus overlap those found in disorders caused by deficiencies in telomere maintenance, the so-called telomere syndromes. Moreover, analysis of telomere length revealed that cells from some, but not all, patients had shortened telomeres (Anderson et al, 2012; Polvi et al, 2012). Most recently, a CTC1 mutation has been found to manifest as dyskeratosis congenita with the classical features of a defect in telomere maintenance (Keller et al, 2012). Thus, the clinical manifestations and cellular phenotypes of CTC1 mutations support both telomeric and non-telomeric roles for CST in human biology. Here, we provide evidence that CST has both telomeric and non-telomeric roles in resolving problems associated with DNA replication. At the telomere, stable knockdown of either CTC1 or STN1 caused MTS and delayed telomere replication. STN1 depletion also leads to a general decrease in both new origin firing and the resumption of DNA replication following treatment with hydroxyurea (HU) to induce genome-wide fork stalling. Together, our findings suggest that human CST plays a key role in replication restart as a specialized replication factor, which promotes DNA replication under conditions of replication stress or at natural replication barriers. Results Knockdown of CTC1 or STN1 promotes genome instability and MTS We previously showed that acute depletion of human CTC1 with siRNA leads to an increase in γH2AX staining, chromatin bridges and a variety of telomere defects including increased telomere loss and G-overhang elongation (Surovtseva et al, 2009). To determine whether STN1 knockdown causes similar defects, we created cell lines with stable knockdown of either STN1 or CTC1 by infecting HeLa 1.2.11 cells with shRNA-encoding lentivirus. Single cell clones were established and the level of knockdown was assessed by immunoblotting and RT–qPCR for STN1 and RT-qPCR alone for CTC1 (due to the lack of a suitable antibody). STN1 mRNA levels were decreased by 60–80% and the protein was barely detectable (Figure 1A and B). CTC1 mRNA levels were decreased by ∼70% (Figure 1B). One of the shSTN1 clones (shSTN1-7) was subsequently transfected with a construct expressing sh-resistant Flag-tagged STN1 (shSTN1-7 Res; Figure 1A) and the resulting cells were used to verify that phenotypes were specific to STN1 depletion and not off-target effects. Figure 1.Depletion of CTC1 or STN1 causes genomic instability in HeLa 1.2.11 cells. (A) Western blot showing knockdown of STN1 (42 kDa) in shSTN1 clones and re-expression of an sh-resistant Flag-STN1 (shSTN1-7 Res). Loading control is α-Actinin (100 kDa). Lanes 1–3 contain 25 μg of protein and lane 4 contains 10 μg. Numbers below gel indicate the level of STN1 relative to non-target control (shNT) after normalization to α-Actinin. (B) RT–qPCR of STN1 and CTC1 mRNA in different clones. Levels are relative to shNT with normalization to GAPDH (mean±s.e.m., n=3 independent experiments). (C, D) Anaphase bridges observed after release of control, shSTN1 or shCTC1 clones from nocadozole block (mean±s.e.m., n⩾3 independent experiments). NT, non-target; WT, wild type. Download figure Download PowerPoint The shSTN1 clones divided at a normal rate and showed no significant growth defects (Supplementary Figure 1A and see below). These results contrast to previous findings with HeLa cells in which acute siRNA knockdown of STN1 caused cell death (Dai et al, 2010). The robust growth of the shRNA clones probably reflects the lower level of knockdown. Our previous studies showed that complete loss of CTC1 causes anaphase bridge formation in Arabidopsis (Surovtseva et al, 2009). Whether lack of CTC1 has a similar effect in human cells was not established because very few cells entered metaphase after the acute CTC1 depletion (Surovtseva et al, 2009). When we examined the HeLa 1.2.11 stable CTC1 and STN1 knockdown cell lines we found they had normal levels of metaphase and anaphase cells and we observed that depletion of either protein caused a significant increase in the frequency of anaphase bridges (Figure 1C and D). The increase due to STN1 depletion was largely prevented by expression of the sh-resistant Flag-STN1. The STN1 knockdown clones also exhibited other phenotypes previously observed after siRNA knockdown of CTC1 including an increase in the number of micronuclei and an increase in the average length of the telomeric G-strand overhang (Supplementary Figure 1B and C), suggesting that, like CTC1, STN1 prevents genome instability and helps maintain G-overhang length. One difference in the phenotype seen after acute CTC1 knockdown versus stable CTC1 or STN1 depletion was the nature of the telomere defects visible by fluorescence in-situ hybridization (FISH). While acute knockdown of CTC1 in HeLa S3 cells (a strain with telomeres of 3–7 kb) yielded an increase in chromosomes lacking telomeric FISH signals (Surovtseva et al, 2009), a similar increase in signal-free ends was not observed in the shCTC1 and shSTN1 HeLa 1.2.11 clones (telomeres of 10–20 kb). Instead, and as previously reported, we observed an increase in the number of chromosome ends with MTS (Figure 2A and B; Supplementary Figure 2A; Price et al, 2010). To further explore this finding, we examined MTS occurrence in two separate shSTN1 and shCTC1 clones and after rescue of the STN1 knockdown with the sh-resistant Flag-STN1. With each knockdown clone, we observed an approximately two-fold increase in MTS that was rescued in the sh-resistant Flag-STN1 cell line (Figure 2B; Supplementary Table 1). Figure 2.CTC1 or STN1 depletion cause multi-telomeric signals (MTS). (A, C) Telomere FISH of HeLa 1.2.11 shCTC1 or shSTN1 clones (A) or U2OS shSTN1 clone (C) showing examples of MTS (white arrows). Green, FITC-telomere probe; blue, DAPI. (B) Quantification of MTS in HeLa 1.2.11 cell lines. Metaphase spreads were made from cells grown±0.25 μg/ml aphidicolin for 16 h prior to the addition of colchicine or colcemid (mean±s.e.m., n⩾3 independent experiments). (D) Quantification of MTS from a single experiment with a U2OS STN1 knockdown clone. NT, non-target; WT, wild type. Download figure Download PowerPoint To determine whether our ability to detect signal-free ends in HeLa S3 after CTC1 siRNA knockdown but not in HeLa 1.2.11 after stable CTC1 knockdown reflected use of different HeLa strains or the different knockdown approach, we used siRNA to deplete CTC1 in HeLa 1.2.11 cells (Supplementary Figure 2B; Supplementary Table 1). The siRNA depletion caused a large increase in signal-free ends and a smaller, but consistent, increase in MTS (Supplementary Figure 2C). Thus, acute CTC1 knockdown appears to favour telomere loss over MTS formation while stable knockdown of CTC1 or STN1 causes MTS alone. It is likely that acute knockdown of CTC1 in the HeLa S3 cells also caused some MTS but the short telomeres made them difficult to detect. We next examined whether stable STN1 knockdown caused MTS or loss of telomere signal in U2OS cells, an ALT cell line with long heterogeneous telomeres (>20 kb). As with the HeLa 1.2.11 cells, we observed an increase in MTS but not in signal-free ends (Figure 2C and D; Supplementary Figure 3C and D; Supplementary Table 1). Interestingly, STN1 knockdown caused a significant growth defect in U2OS cells. Although the level of knockdown was similar to that observed in the HeLa 1.2.11 cells, the STN1-depleted U2OS cells grew more slowly than control U2OS cells (Supplementary Figure 3A and B) and single cell clones only survived for a few weeks, causing us to examine a knockdown pool to verify the MTS phenotype (Supplementary Figure 3C and D). Overall, we conclude that both STN1 and CTC1 are needed for genome stability and telomere maintenance. CST depletion slows replication through the telomeric tract Given that the appearance of MTS can reflect problems associated with replication through the duplex region of the telomere (Sfeir et al, 2009; Saharia et al, 2010), we suspected that the MTS caused by STN1 or CTC1 depletion might stem from a similar cause. To explore this possibility, we examined the effect of combining STN1 or CTC1 knockdown with aphidicolin treatment. Aphidicolin causes replication stress by inhibiting DNA polymerase α, δ and ε (Cheng and Kuchta, 1993) and is known to induce MTS (Sfeir et al, 2009). As expected, aphidicolin treatment caused an increase in MTS levels in both the WT and non-target (shNT) control cell lines (Figure 2B; Supplementary Table 1). However, treatment with aphidicolin in the context of STN1 or CTC1 depletion resulted in an epistatic-like interaction in which the levels of MTS remained similar or slightly decreased relative to those observed without aphidicolin treatment. These findings suggest that CST and DNA polymerase α, δ and/or ε act within a common pathway to prevent MTS formation. While the experiment does not address the exact step at which CST and aphidicolin interface, it supports a role for CST in replication of the telomeric duplex. To test more directly for the role of CST in telomere replication, we next asked whether STN1 depletion delays the overall rate of replication through either the telomere duplex or the bulk of the genome. For this experiment, the HeLa 1.2.11 shSTN1-7, shSTN1-7Res and shNT clones were synchronized at G1/S with a double-thymidine block, released into fresh media and allowed to enter S-phase (Supplementary Figures 4E and 5E). Cells were then pulsed labelled with either BrdU or EdU for consecutive 1.5 h intervals (Figure 3A) and harvested at the end of each time point. First, we examined the overall rate of genome replication. The EdU-labelled cells were fixed, the EdU was reacted with fluorophore and the relative amount of EdU uptake measured by FACS. As shown in Figure 3B and Supplementary Figures 4A, 4B, 5A and 5B, the rate of EdU uptake in the shSTN1, shSTN1-7-Res and control shNT cells was essentially identical throughout S-phase indicating that, at this level of knockdown, STN1 depletion does not affect the rate of whole genome replication. Figure 3.STN1 depletion delays telomere replication but does not affect the rate of bulk genomic DNA replication. (A) Experimental timeline. HeLa 1.2.11 cells were released from a double-thymidine block into S-phase and incubated with BrdU or EdU for consecutive 1.5 h intervals. (B) Rates of bulk genomic DNA replication were determined by EdU uptake. Graph shows EdU incorporated at consecutive time periods (Mean EdU staining X % EdU-positive cells). (C–E) Rates of telomere replication throughout S-phase. (C) BrdU-labelled DNA from 4.5 and 6 h time points was subject to CsCl sedimentation to separate leading and lagging strand telomeres. Telomeric DNA from each gradient fraction was quantified by slot blot hybridization. (D) Per cent of newly replicated leading strand telomere signals relative to the total telomere signal for each time period throughout S-phase. (E) Examples of slot blot used to obtain data in (C) and (D). Data are representative of three independent experiments. NT, non-target. Download figure Download PowerPoint Next, we quantified the amount of telomeric DNA replicated at each time point throughout S-phase. DNA from the BrdU-labelled cells was isolated, restriction digested and subjected to CsCl density gradient centrifugation to separate unreplicated and replicated telomeres (Chai et al, 2006). The gradient was fractionated and the relative amount of telomeric DNA in each fraction was quantified by slot blot hybridization using a telomeric DNA probe (Figure 3C–E; Supplementary Figures 4C, 4D, 5C and 5D). Telomeres replicated by leading strand synthesis incorporate multiple BrdU molecules per telomeric repeat (TTAGGG) and hence sediment at a higher density than telomeres replicated by lagging strand synthesis (CCCTAA) and both are separated from any unreplicated telomeric DNA (Chai et al, 2006; Zhao et al, 2011). Quantification of replicated telomeric DNA from the shNT, shSTN1-7 and shSTN1-7 Res cells revealed a considerable difference in the timing of telomere replication in the control versus the STN1-depleted cells (Figure 3C–E; Supplementary Figures 4C, 4D, 5C and 5D). Although all three cell types initiated telomere replication in a similar manner, the STN1 knockdown cells completed replication more slowly such that telomere replication reached a maximum and then declined 1.5–3 h earlier in the control cells (Figure 3D; Supplementary Figures 4D and 5D). While there was some experiment-to-experiment variation in the timing of maximal telomere replication, the delay in the STN1 knockdown cells was very consistent and was readily apparent regardless of whether we quantified the total amount of replicated telomere DNA (Supplementary Figure 4D) or the amount of the leading strand peak (which was more visible at early time points) (Figure 3D; Supplementary Figure 5D). Thus, our results indicate that STN1 depletion slows replication of the telomere duplex without affecting the rate of bulk genomic DNA replication. We therefore conclude that components of the CST complex play a specific role in promoting efficient replication of the telomeric tract. The above findings also provide strong support for our proposal that the MTS observed after STN1 or CTC1 depletion result from problems associated with telomere replication. STN1 and TRF1 promote telomere replication via different pathways Since CST does not appear to be a general replication factor, possible functions for STN1 in replication of the telomere duplex DNA could include helping prevent replication fork stalling or promoting subsequent replication restart. To learn more about the role of CST in these processes, we next examined the effect of STN1 and TRF1 co-depletion on MTS frequency. TRF1 helps prevent fork stalling during telomere replication and the increased stalling caused by TRF1 depletion results in elevated MTS (Martinez et al, 2009; Sfeir et al, 2009). Thus, analysis of MTS levels after co-depletion of STN1 and TRF1 should indicate whether STN1 affects fork stalling via the same or different pathways. TRF1 was depleted in the HeLa 1.2.11 shSTN1-7 clone by transfecting cells twice, 24 h apart, with a previously characterized siRNA (Ohishi et al, 2010). Forty-eight hours after the second transfection, RNA was extracted and the level of knockdown measured by RT–qPCR (Figure 4A). Telomere FISH was performed 48 h after the second transfection and the number of MTS determined. As expected, MTS levels were increased approximately two-fold above background with either TRF1 or STN1 single knockdown (Figure 4B). The overall background levels of MTS were higher than observed in previous experiments with HeLa 1.2.11 (compare Figure 2B and Figure 4B). This was most likely due to the siRNA transfection as the fold increase with STN1 knockdown was similar in both experiments. When we compared MTS levels caused by co-depletion of STN1 and TRF1 relative to those observed after STN1 or TRF1 single knockdown, we consistently observed a greater than additive increase in MTS (Figure 4B; Supplementary Table 1). This result indicates that STN1 and TRF1 affect different processes during telomere replication. Given that CTC1 and STN1 appear to function in conjunction with DNA polymerase to prevent MTS formation (Figure 2B) and CST acts as a DNA pol α affinity factor (Goulian and Heard, 1990; Goulian et al, 1990), the result also suggests that the increase in MTS after STN1 depletion might reflect a role for STN1 in replication restart rather than in the prevention of fork stalling. Figure 4.Co-depletion of TRF1 and STN1 causes an additive increase in MTS. (A) Relative level of TRF1 mRNA 48 h after siRNA transfection as measured by RT–qPCR with normalization to GAPDH (mean±s.e.m., n=3 independent experiments). (B) Quantification of MTS (mean±s.e.m., n=3 independent experiments). Dashed line indicates the background level of MTS. NT, non-target. Download figure Download PowerPoint STN1 promotes genome-wide replication restart after HU-induced fork stalling Since CST appears to have both telomeric and non-telomeric functions, we hypothesized that CST might promote recovery from replication fork stalling at non-telomeric locations. To test this possibility, we examined whether STN1 promotes DNA replication restart after fork stalling across the genome. Replication fork stalling was induced by treatment with HU, a ribonucleotide reductase inhibitor, which stalls DNA polymerization by depleting nucleotide pools (Koc et al, 2004). Cells were treated with moderately high levels of HU (2 mM) over a short time frame (2 h) to avoid inducing fork collapse (Petermann and Helleday, 2010; Petermann et al, 2010). After HU treatment, cells were released into media containing EdU for 30 min to label cells that resumed replication. Cells were then fixed, the EdU was reacted with fluorophore and actively replicating cells identified by immunofluorescence (Figure 5A). Following image capture, the mean fluorescence intensity of the EdU signal was quantified. The results are presented as both the per cent of nuclei at different Arbitrary Fluorescence Units (AFU) (Figure 5B) and the average fluorescence intensity of all nuclei counted (Figure 5C). HU-induced fork stalling was verified by the lack of EdU incorporation during HU treatment. Figure 5.Replication restart after HU treatment is inhibited by STN1 depletion. (A–C) Cells were treated for 2 h with 2 mM HU and released into medium containing EdU for 30 min. (A) EdU incorporation by HeLa 1.2.11 clones after release from HU. Blue, DAPI; green, EdU. (B) Quantification of the levels of EdU uptake after release from HU, as measured by mean fluorescence intensity (mean±s.e.m., n⩾3 independent experiments). Each bar indicates the total number of nuclei above or below the AFU given below. AFU, arbitrary fluorescence units. Nuclei below 10 AFU are considered as EdU negative, those above 10 AFU are EdU positive. (C) Average AFU values of all nuclei following HU removal for both HeLa1.2.11 knockdown clones and pools of U2OS knockdown cells (mean±s.e.m., n⩾3 independent experiments). NT, non-target. Download figure Download PowerPoint As anticipated, recovery from HU treatment resulted in a significant decrease in EdU uptake relative to untreated cells, reflecting gradual recovery from fork stalling. Interestingly, the HU caused a slower recovery (less EdU uptake) in the shSTN1 HeLa 1.2.11 clones than in the shNT control cells (Figure 5C). Furthermore, STN1 depletion caused the per cent of EdU-negative nuclei (mean AFU ⩽10) to increase and the per cent of nuclei with higher levels of EdU incorporation (AFU >20) to decrease (Figure 5B). These effects were largely rescued by expression of the sh-resistant Flag-STN1 allele (Figure 5B and C). Importantly, without HU treatment, the per cent of EdU-positive cells and the levels of EdU incorporation were similar in the shSTN1 and control HeLa 1.2.11 cells, indicating that the decrease in EdU incorporation after HU treatment was not due to inherent differences in the number of cells in S-phase or rates of replication (Figure 5C). We therefore conclude that STN1 depletion delays replication restart after fork stalling in HeLa 1.2.11 cells. To examine whether the deficiency in replication restart was a general phenomenon, we also examined EdU incorporation in STN1-depleted U2OS cells (Figure 5C; Supplementary Figure 6). As observed with
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