RNAi-based screening identifies the Mms22L–Nfkbil2 complex as a novel regulator of DNA replication in human cells
2010; Springer Nature; Volume: 29; Issue: 24 Linguagem: Inglês
10.1038/emboj.2010.304
ISSN1460-2075
AutoresWojciech Piwko, Michael H. Olma, Michael Held, Julien N Bianco, Patrick G. A. Pedrioli, Kay Hofmann, Philippe Pasero, Daniel W. Gerlich, Matthias Peter,
Tópico(s)Mitochondrial Function and Pathology
ResumoArticle26 November 2010free access RNAi-based screening identifies the Mms22L–Nfkbil2 complex as a novel regulator of DNA replication in human cells Wojciech Piwko Corresponding Author Wojciech Piwko Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, Switzerland Search for more papers by this author Michael H Olma Michael H Olma Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, Switzerland Search for more papers by this author Michael Held Michael Held Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, Switzerland Search for more papers by this author Julien N Bianco Julien N Bianco Institute of Human Genetics, CNRS UPR 1142, Montpellier, France Search for more papers by this author Patrick G A Pedrioli Patrick G A Pedrioli Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, SwitzerlandPresent address: The Scottish Institute for Cell Signalling, The Sir James Black Centre, College of Life Sciences, University of Dundee, Dow Street, Dundee DD1 5EH, Scotland Search for more papers by this author Kay Hofmann Kay Hofmann Miltenyi Biotec GmbH, Bioinformatics Department, Bergisch-Gladbach, Germany Search for more papers by this author Philippe Pasero Philippe Pasero Institute of Human Genetics, CNRS UPR 1142, Montpellier, France Search for more papers by this author Daniel W Gerlich Daniel W Gerlich Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, Switzerland Search for more papers by this author Matthias Peter Corresponding Author Matthias Peter Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, Switzerland Search for more papers by this author Wojciech Piwko Corresponding Author Wojciech Piwko Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, Switzerland Search for more papers by this author Michael H Olma Michael H Olma Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, Switzerland Search for more papers by this author Michael Held Michael Held Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, Switzerland Search for more papers by this author Julien N Bianco Julien N Bianco Institute of Human Genetics, CNRS UPR 1142, Montpellier, France Search for more papers by this author Patrick G A Pedrioli Patrick G A Pedrioli Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, SwitzerlandPresent address: The Scottish Institute for Cell Signalling, The Sir James Black Centre, College of Life Sciences, University of Dundee, Dow Street, Dundee DD1 5EH, Scotland Search for more papers by this author Kay Hofmann Kay Hofmann Miltenyi Biotec GmbH, Bioinformatics Department, Bergisch-Gladbach, Germany Search for more papers by this author Philippe Pasero Philippe Pasero Institute of Human Genetics, CNRS UPR 1142, Montpellier, France Search for more papers by this author Daniel W Gerlich Daniel W Gerlich Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, Switzerland Search for more papers by this author Matthias Peter Corresponding Author Matthias Peter Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, Switzerland Search for more papers by this author Author Information Wojciech Piwko 1,‡, Michael H Olma1,‡, Michael Held1, Julien N Bianco2, Patrick G A Pedrioli1, Kay Hofmann3, Philippe Pasero2, Daniel W Gerlich1 and Matthias Peter 1 1Institute of Biochemistry, Department of Biology, ETH Zurich, Zurich, Switzerland 2Institute of Human Genetics, CNRS UPR 1142, Montpellier, France 3Miltenyi Biotec GmbH, Bioinformatics Department, Bergisch-Gladbach, Germany ‡These authors contributed equally to this work *Corresponding authors. Institute of Biochemistry, Department of Biology, ETH Zurich, 8093 Zurich, Switzerland. Tel.:+41 44 63 32 564; Fax: +41 44 63 21 298; E-mail: [email protected] or Tel.:+41 44 63 36 586; Fax: +41 44 63 21 298; E-mail: [email protected] The EMBO Journal (2010)29:4210-4222https://doi.org/10.1038/emboj.2010.304 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 Cullin 4 (Cul4)-based ubiquitin ligases emerged as critical regulators of DNA replication and repair. Over 50 Cul4-specific adaptors (DNA damage-binding 1 (Ddb1)–Cul4-associated factors; DCAFs) have been identified and are thought to assemble functionally distinct Cul4 complexes. Using a live-cell imaging-based RNAi screen, we analysed the function of DCAFs and Cul4-linked proteins, and identified specific subsets required for progression through G1 and S phase. We discovered C6orf167/Mms22-like protein (Mms22L) as a putative human orthologue of budding yeast Mms22, which, together with cullin Rtt101, regulates genome stability by promoting DNA replication through natural pause sites and damaged templates. Loss of Mms22L function in human cells results in S phase-dependent genomic instability characterised by spontaneous double-strand breaks and DNA damage checkpoint activation. Unlike yeast Mms22, human Mms22L does not stably bind to Cul4, but is degraded in a Cul4-dependent manner and upon replication stress. Mms22L physically and functionally interacts with the scaffold-like protein Nfkbil2 that co-purifies with histones, several chromatin remodelling and DNA replication/repair factors. Together, our results strongly suggest that the Mms22L–Nfkbil2 complex contributes to genome stability by regulating the chromatin state at stalled replication forks. Introduction Accurate and complete DNA duplication is fundamental to the maintenance of genomic integrity and is ensured by the cooperation of cell cycle checkpoints and DNA repair pathways. However, progressing replication forks encounter many obstacles, including tightly packed histone-associated heterochromatic regions, a myriad of non-nucleosomal protein impediments such as sites of active transcription, telomeres and centromeres, and damaged templates (Tourriere and Pasero, 2007). Dedicated mechanisms are required to slow down and modify active replication forks at these sites and require checkpoint-dependent stabilisation of the replisome. Occasionally, however, replication forks collapse and must be rescued by the firing of late origins or by specialised restart/repair processes that prevent the formation of double-strand breaks (DSBs), illegitimate recombination and gross chromosomal rearrangements. Although these mechanisms are of crucial importance to maintain genome integrity, little is known about the components and regulatory signals. The phosphatidylinositol 3 kinase-related protein kinases (PIKKs), ATM, ATR and DNA-dependent protein kinase (DNA-PK), emerged as key factors in several DNA damage response pathways. ATM and DNA-PK are activated primarily by the presence of DSBs, whereas stalled replication forks and the presence of replication protein A (RPA)-coated ssDNA stimulate ATR (Shiloh, 2003; Cimprich and Cortez, 2008). ATM and ATR and their major effector kinases Chk2 and Chk1 slow down or arrest cell cycle progression to allow time for efficient repair. In addition, PIKKs orchestrate DNA damage repair in a manner not only specific for the type of damage but also for the cell cycle stage at which the lesion occurs. DSBs, one of the most toxic DNA lesions, are repaired by two major mechanisms—non-homologous end joining (NHEJ) and homologous recombination (HR). NHEJ leads to ligation of two DNA ends without sequence homology and is predominantly used to repair breaks occurring during G1 or in specialised cases during S phase. In contrast, HR occurs preferentially in S and G2 and takes advantage of the presence of the sister chromatid, which is used as a template for error-free homology-driven repair (Branzei and Foiani, 2008). In addition to phosphorylation, DNA replication and repair pathways are regulated at multiple levels by ubiquitylation of key components, in particular by cullin 4 (Cul4)-RING E3 ubiquitin ligases (CRL4) (Lee and Zhou, 2007). All cullins interact via their C-terminal region with the RING finger protein Rbx1/Hrt1, which in turn recruits an E2 ubiquitin-conjugating enzyme. The N-terminal domain of cullins is specific for each subfamily, and interacts with distinct classes of substrate-specific adaptors (Pintard et al, 2004). The human genome encodes two Cul4-paralogues, Cul4A and Cul4B, both of which bind via their N-terminal regions to the DNA damage-binding 1 (Ddb1) protein. In turn, Ddb1 interacts with the various substrate-specific adaptor subunits, which belong to a large group of WD40 repeat (WDR)-containing proteins (Ddb1–Cul4-associated factors; DCAFs). However, the critical substrates and specific DCAFs involved in DNA replication and repair processes remain poorly understood. CRL4Cdt2 regulates several S phase-related processes by using the replicative polymerase sliding clamp PCNA as a cofactor to target the replication-licensing factor Cdt1 (Kim and Kipreos, 2007), the cyclin-dependent kinase (CDK) inhibitor p21 (Abbas et al, 2008) and the TLS polymerase eta Polη (Kim and Michael, 2008). Moreover, Cul4 complexes control two nucleotide excision repair (NER) pathways. For example, CRL4Ddb2 is recruited to UV-induced DNA lesions, where it mediates ubiquitylation of the repair protein XPC (Sugasawa et al, 2005), while the CRL4CSA complex targets CSB for degradation to remove stalled RNA polymerase II from damaged templates (Sugasawa et al, 2005; Fousteri et al, 2006). CRL4 complexes also support efficient DNA repair by altering nucleosome stability around the damaged site. For example, Cul4 ubiquitylates histone H2A, H3 and H4 near UV lesions, thereby causing histone eviction to expose the damaged DNA to repair proteins (Kapetanaki et al, 2006; Wang et al, 2006). Progression of DNA replication in Cul4-depleted cells is perturbed and accompanied by strong upregulation of the phosphorylated histone H2A variant H2AX (γH2AX), indicating the occurrence of spontaneous DSBs during the S phase (Olma et al, 2009). This is reminiscent of the function of Rtt101, the putative Cul4 orthologue in budding yeast, which, in complex with Mms1 and Mms22, has a crucial role in DNA replication through natural pause sites and damaged templates (Luke et al, 2006). Indeed, detailed studies revealed that rtt101Δ cells are genetically unstable because they are unable to promote HR at stalled replication forks during S phase (Duro et al, 2008). Mms22 binds to the N-terminal region of Rtt101 via Mms1, which shares significant similarities to human Ddb1, suggesting that Mms22 is a substrate-specific adaptor for the Rtt101-based ubiquitin ligase (Zaidi et al, 2008). Finally, genetic and biochemical experiments suggest that the Rtt101Mms22 E3 ligase functions downstream of Rtt109, a histone acetyltransferase required for H3-K56 acetylation, which marks newly replicated DNA and is required for genome stability maintenance in yeast and mammalian cells (Collins et al, 2007; Roberts et al, 2008). These studies suggest that a conserved Rtt101/Cul4-dependent pathway may be involved in replicating through chromosomal slow zones and promotes restart of stalled replication forks. In this study, we used automated live-cell microscopy and computational image analysis to quantify interphase timing in an RNAi screen, targeting all known and bioinformatically predicted substrate adaptors of Cul4. We identified several Cul4-associated proteins specifically regulating the duration of the G1, S and G2 phases of the cell cycle, and C6orf167/Mms22-like protein (Mms22L), a putative human homologue of yeast Mms22. Biochemical and cell biological analysis suggests that Mms22L, in complex with Nfkbil2, is required to ensure genome stability during DNA replication in mammalian cells. Results RNAi-based screening for Cul4-linked genes involved in cell cycle progression To investigate the functional importance of the different Cul4 substrate adaptors for DNA replication and genome integrity, we developed an automated RNAi-based screening protocol allowing quantification of replication and mitosis timing. We used a HeLa cell line stably expressing mCherry-tagged histone H2B as a chromatin marker to discriminate the mitotic stages, and EGFP-tagged PCNA to visualise the characteristic morphological changes of DNA replication foci during the S phase (Figure 1A). Automated time-lapse imaging data were annotated using the CellCognition computational framework (Held et al, 2010) to classify 11 different cell cycle stages by supervised machine learning (G1, early, mid and late S, and G2 phase based on EGFP-PCNA, and six mitotic stages based on H2B-mCherry; Figure 1A and B), which allowed to measure the duration of each single cell cycle stage. To validate the sensitivity and performance of this assay, we analysed the phenotype of HeLa cells that were RNAi-depleted for the known cell cycle regulators Skp2, Ddb1 and Rad51 (Figure 1C). In agreement with previous publications, downregulation of each of these genes resulted in the expected delays in G1, S and G2 phases, respectively (Sonoda et al, 1998; Sutterluty et al, 1999; Lovejoy et al, 2006). Figure 1.Automated high-content RNAi screening assay to measure the timing of different cell cycle phases. (A) HeLa cells stably expressing the chromatin marker H2B-mCherry (red) and the replication factory marker EGFP-PCNA (green) were imaged for 47 h using an automated wide-field epifluorescence microscope with a 10 × dry objective. Single cells were detected by local adaptive thresholding and distinct cellular morphologies were classified by supervised machine learning. Trajectories of single cells containing prometaphase stages were extracted and classification was improved with a hidden Markov model (HMM)-based method (Held et al, 2010). This yielded accurate annotations of all indicated cell cycle phases (lower panels) and apoptosis (not shown). (B) Overview of all detected cell trajectories for an RNAi control in the two channels sorted by occurrence in the movie and aligned at the prophase–prometaphase transition. (C) Cell trajectories detected in time-lapse movies of cells RNAi-depleted of Skp2, Ddb1 and Rad51 (oligos 11, 7 and 6), aligned at the mitosis-G1, G1-early S phase and late S phase–G2 transitions and sorted by G1, S and G2 phase length, respectively. Download figure Download PowerPoint To identify Cul4 adaptors involved in DNA replication, we designed an siRNA library targeting 147 genes associated with Cul4, including known and bioinformatically predicted DCAFs and other Cul4-interacting proteins (Figure 2A and Supplementary Table 1). Each gene was targeted by three to six distinct siRNA oligos (Supplementary Table 1), using reverse transfection in 96-well plates. Image acquisition was started 25 h after RNAi and cells were imaged for 47 h with a time resolution of approximately 6 min. Using the CellCognition software, we measured the duration of the respective cell cycle phases for at least 15 cells per siRNA, and used the calculated medians to classify each of the candidate genes. If more than half of all quantified siRNAs showed a z-score above 5, relative to the negative controls, we considered this gene as a potential hit. In all, 50 genes showed a significant delay in G1 phase (Supplementary Table 2), including 17 bona fide DCAFs characterised by WD40 domains (Figure 2B). As an example, Figure 2C and D illustrate the analysis of cells depleted of DCAF Wdr51B, which extended G1 phase to approximately 15 h, compared with 7 h for RNAi controls. Four DCAFs were not only necessary for timely G1 progression but also exhibited significant delays in S phase. Among them is the known DCAF, Wdr26, which slows down DNA replication by approximately 20%, as visualised by persistent PCNA foci (Supplementary Table 2, Figure 2E and F). In addition, we identified seven genes including four DCAFs specifically required for efficient progression through S phase, without significantly affecting G1 and G2 length. Taken together, this RNAi-based screen identified a subset of known or predicted CRL4 adaptors required for timely interphase progression, and suggested specific functions of CRL4-based E3 ligases in G1, entry into S phase and efficient progression through early and late stages of DNA replication. Figure 2.RNAi-based screen for Cul4-related genes involved in regulation of interphase in HeLa cells. (A, B) An RNAi library targeting Cul4-associated genes was based on MS analysis of purified Cul4A complexes and yeast two-hybrid screens of Cul4A and Cul4B. This list was complemented with published Cul4 interactors and detailed bioinformatic analysis of Cul4-related proteins. A total of 147 CRL4-related genes were then analysed by RNAi depletion for interphase delays using the automated screening assay depicted in Figure 1. Bona fide DCAFs showing specific delays at the indicated cell cycle phase are listed (B). The entire list including all analysed genes is provided in Supplementary Table 2. (C–H) The duration of the indicated cell cycle phase was measured for at least 20 cells depleted of selected candidates, and the mean, s.d. and the Student t-test P-value were calculated (C, E, G; oligos 2, 4, 5, 4, 7 and 8, respectively). Time-lapse microscopy images of a representative cell depleted for the indicated Cul4-related proteins, progressing through G1 (D, oligo 2), S (F, oligo 5) and G2 (H, oligo 7) phases. Download figure Download PowerPoint C6orf167/Mms22L is required for maintenance of genomic stability during S phase Downregulation of only two genes, PolR2E and C6orf167, caused a significant prolongation of G2 phase. The pronounced G2 delay of C6orf167-depleted cells classified with an average z-score of 20 and, in contrast to PolR2E, was not accompanied by changes in G1 or S phase duration (Figure 2G and H; Supplementary Table 3). Interestingly, C6orf167 codes for a yet uncharacterised human protein with low, but significant, similarity to yeast Mms22 (Supplementary Figure 1). Given the bioinformatic conservation and functional similarities in maintaining genome stability during DNA replication, we named C6orf167 as Mms22L. To test whether the prolonged G2 phase in Mms22L-depleted cells results from activation of the DNA damage checkpoint, we performed immunoblot analysis of extracts derived from HeLa cells treated with siRNA specifically targeting Mms22L, using phospho-specific antibodies against activated checkpoint proteins. Indeed, we observed increased phosphorylation of ATM on Ser 1981, Chk2 on Thr 68 and Chk1 on Ser 345 upon depletion of Mms22L (Figure 3A and Supplementary Figure 2), which suggests activation of both the ATM and ATR branches of the DNA damage checkpoint. This was accompanied by hyperphosphorylation of several PIKK substrates, including the histone H2A variant H2AX on Ser 139 (γH2AX) and RPA2, markers of DNA damage and replication stress, respectively (Anantha et al, 2007). Similar results were also observed for U2OS cells (Supplementary Figure 3; Supplementary data not shown), implying that the defects are not solely explained by the transformed phenotype of HeLa cells. The G2 arrest may be a direct consequence of DNA damage checkpoint activation, as the G2 phase duration of Mms22L-depleted HeLa cells treated with the ATM/ATR inhibitor caffeine was reduced from 8.8 to 4.7 h, which is comparable to RNAi controls (Figure 3B). Taken together, these results suggest that depletion of Mms22L induces a strong delay in G2 because of activation of the ATM/ATR-dependent checkpoint response pathway. Figure 3.C6orf167/Mms22L is required to maintain genome integrity during the S phase. (A) Extracts from HeLa cells treated with control or Mms22L (oligo 1) siRNAs for 72 h were immunoblotted with specific antibodies recognising the indicated markers. pRPA2 marks phosphorylated forms of RPA2; tubulin was included as loading control. (B) Control and Mms22L-depleted (pooled oligos 1–3) HeLa cells expressing EGFP-PCNA and H2B-mCherry were imaged for 2 days with a time resolution of 26 min. Cells were either treated (grey bars) or not treated (white bars) with 1 mM caffeine to inhibit ATM/ATR. Where indicated, control cells were treated with 0.02 μM CPT to induce checkpoint activation (last column). G2 duration was determined from at least 25 cells for each condition based on the PCNA and H2B markers as described in Figure 1. (C, D) Mms22L-depleted (oligo 1) HeLa cells accumulate nuclear γH2AX foci that colocalise with 53BP1 (C) and RPA2 (D), as visualised by immunofluorescence analysis with specific antibodies. DNA was stained with DAPI; the scale bar represents 20 μm. (E, F) HeLa cells were treated with control and Mms22L (oligo 1) siRNAs for 72 h, and ethanol-fixed γH2AX-positive cells were quantified by flow cytometry using specific antibodies. Propidium iodide (PI) was used to stain DNA. The average percentage of γH2AX-positive cells is indicated in the gated area (E). S phase was divided into four subphases using FlowJo software, and the percentage of γH2AX-positive cells was measured for each cell cycle phase (F). (G) Single-molecule analysis of DNA replication in Mms22L-depleted U2OS cells. Cells treated with control or Mms22L (pooled oligos 1–3) siRNAs for 72 h were pulse-labelled for 20 min with BrdU. Fibres were stretched by DNA combing and labelled DNA was visualised by specific antibodies against BrdU. The graph shows the distribution and median values of BrdU tract length. Representative images of DNA fibres are shown in Supplementary Figure 4. (H) Mms22L downregulation causes CPT hypersensitivity. HeLa cells treated with control siRNA or siRNAs downregulating Mms22L (pooled oligos 1–3) were exposed for 1 h to 1 μM CPT, and the number of surviving colonies was determined after 8–10 days. Cells depleted for the HR-component CtIP were included for positive control. Bars with s.d. represent the percentage (%) of surviving colonies compared with untreated controls averaged from four independent experiments. Download figure Download PowerPoint To corroborate these results, we used indirect immunofluorescence microscopy using antibodies specific for γH2AX to visualise DSBs. Indeed, the number of γH2AX foci strongly increased upon RNAi depletion of Mms22L in HeLa (Figure 3C and D, and Supplementary Figure 2) and U2OS cells (Supplementary Figure 3). Moreover, this signal colocalised with foci formed by 53BP1 (Figure 3C), an important mediator of ATM signalling (Schultz et al, 2000; Wang et al, 2002), and RPA2 (Figure 3D), which marks extended ssDNA regions (Binz et al, 2004). We also examined Mms22L-depleted cells by flow cytometry (Figure 3E). As expected, the cell cycle analysis showed dramatic accumulation of cells in G2/M, and a concomitant reduction of cells in both G1 and S phase. Staining with γH2AX-specific antibodies revealed that DSBs gradually accumulate during S phase and partially diminish during G2 (Figure 3E and F). In contrast, γH2AX was only weakly increased during G1, suggesting that, in the absence of Mms22L, accumulation of DSBs arising specifically during S phase triggers a checkpoint-dependent arrest in G2 to allow sufficient time for repair before entering mitosis. Consistent with a specific function during the S phase, DNA combing experiments revealed that replication forks moved significantly slower in U2OS cells depleted for Mms22L in comparison with the RNAi control (Figure 3G and Supplementary Figure 4). Collectively, these phenotypes are reminiscent of yeast cells lacking components of the Rtt101Mms22 E3 ligase, which regulates faithful DNA replication through damaged templates or replication slow zones (Duro et al, 2008; Zaidi et al, 2008). Because rtt101Δ and mms22Δ cells are strongly sensitive to genotoxic agents causing DNA damage during S phase (Luke et al, 2006; Collins et al, 2007; Zaidi et al, 2008), we next measured the clonogenic survival of Mms22L-depleted HeLa cells exposed to topoisomerase I (TopoI) poison camptothecin (CPT). As CPT-induced DSBs at replication forks are mainly repaired by HR-mediated mechanisms, depletion of the HR-promoting gene CtIP was included as a control (Sartori et al, 2007). Mms22L was required to protect HeLa cells from CPT treatment, although to a lesser extent than CtIP (Figure 3H). Together, we conclude that Mms22L identifies an evolutionarily conserved pathway required to repair DNA damage during S phase, possibly by promoting replication-associated HR. Mms22L is degraded upon DNA damage induced during S phase To characterise the Mms22L protein, we raised polyclonal antibodies against the N-terminal 325 amino acid fragment of Mms22L. Immunoblot analysis detected a single band of ∼130 kDa, which was strongly reduced upon depletion of Mms22L by RNAi (Figure 3A). To examine cell cycle expression of endogenous Mms22L, we synchronised HeLa cells in early S phase using a double thymidine block/release (DTB/R) protocol (Sumara et al, 2007). Mms22L was expressed throughout S and G2 phase, but its levels were reduced upon entering G1 (Figure 4A), indicating that Mms22L may be regulated in a cell cycle-dependent manner. Figure 4.Mms22L levels are regulated throughout the cell cycle and upon DNA damage induced during S phase. (A) HeLa cells were synchronised in early S phase using the DTB/R protocol and harvested at the indicated time points after release from the second thymidine block (in hours). Nocodazole (Noc)- or HU-arrested cells were included as controls. Protein extracts were immunoblotted with antibodies against Mms22L and cell cycle markers cyclin E (S phase), cyclin A (S/G2) and phospho-Ser10 H3 (mitosis). GAPDH controls for equal loading. (B) The localisation of HSS-Mms22L was analysed after transient transfection by indirect immunofluorescence using anti-HA antibodies. DAPI visualises DNA; scale bar: 20 μm. (C) Mms22L is associated with chromatin. HeLa whole-cell extracts (WCE) were fractionated into cytosol (C), nucleoplasm (N) and chromatin (Ch) (left panels). The chromatin fraction was subjected to DNA digestion using micrococcal nuclease, incubated in buffer containing 100 or 300 mM KCl, and separated into soluble (SN) and pellet (P) fractions (right panels). Samples were analysed by immunoblotting with antibodies against Mms22L, the nuclear marker histone H3 and cytoplasmic tubulin. (D) HSS-Mms22L is stabilised in Cul4- and Ddb1-depleted cells. HeLa cells stably expressing HSS-Mms22L were RNAi-depleted of Mms22L (oligo 2), Cul4A and Cul4B or Ddb1 for 24 h. Doxycycline was added for an additional 24 h to induce HSS-Mms22L expression, and protein extracts were immunoblotted for the indicated proteins. (E, F) HeLa cells were synchronised by DTB/R and treated 2, 4 and 5.5 h after release with 1 μM CPT for 3 h. Mms22L levels were analysed by immunoblotting (E). HeLa cells synchronised in S phase (3 h after DTB/R) were treated with either 1 μM CPT, 0.05% (v/v) MMS, 2 mM HU, 20 μM MG132 or 5 mM caffeine, and analysed by immunoblotting with the indicated antibodies (F). Note that caffeine does not inhibit RPA2 phosphorylation induced by HU treatment, as demonstrated previously (Cortez, 2003). Download figure Download PowerPoint To examine the subcellular localisation of Mms22L, we first analysed the localisation of an HA-2xStrep (HSS)-tagged version of Mms22L in transiently transfected HeLa cells. As shown in Figure 4B, HSS-Mms22L was primarily nuclear, and this localisation did not change during the cell cycle or after treatment of cells with CPT (data not shown). Consistent with these results, biochemical fractionation revealed that endogenous Mms22L was enriched in the chromatin fraction, but was solubilised by DNA digestion and high-salt treatment (Figure 4C). Together, these results suggest that a fraction of Mms22L is associated with chromosomes to support genome stability during S phase. We observed that the levels of HSS-Mms22L strongly increased in cells RNAi-depleted for Cul4A/B or Ddb1 (Figure 4D), suggesting that Mms22L is an unstable protein that is degraded in a Cul4-dependent manner. Therefore, we tested the stability of endogenous Mms22L in asynchronous and S phase-arrested HeLa cells by CHX chase and found that Mms22L was stable under these conditions (Supplementary Figure 5 and data not shown). However, Mms22L levels were specifically reduced in S phase cells exposed to CPT, and the half-life of Mms22L decreased to ∼2 h (Figure 4E and Supplementary Figure 5). A similar reduction was also observed after treating S phase cells with the DNA-methylating agent methyl methanesulphonate (MMS), but not hydroxyurea (HU), which stalls replication forks without inducing DSBs (Figure 4F). Although we were unable to detect ubiquitylated forms of Mms22L, its degradation was dependent on the activity of the ubiquitin-proteasome system (UPS) and DNA damage checkpoint signalling, as it was blocked by addition of the proteasome inhibitor MG132 and caffeine, respectively (Figure 4F and Supplementary Figure 5). Taken together, these results suggest that Mms22L activity upon replication stress is regulated by a Cul4- and ATM/ATR-dependent mechanism. Mms22L stably interacts with Nfkbil2 Prompted by the homology to yeast Mms22, we first examined whether Mms22L may stably interact with Cul4 and its linker Ddb1. However, we were unable to detect Cul4A or Ddb1 in Mms22L immunoprecipitates, even when the cells were treated with CPT to induce DNA damage (Figure 5A). Simil
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