Title: Mouse Rif1 is a key regulator of the replication-timing programme in mammalian cells
Abstract: Article31 July 2012free access Mouse Rif1 is a key regulator of the replication-timing programme in mammalian cells Daniela Cornacchia Daniela Cornacchia EMBL Mouse Biology Unit, Monterotondo, Rome, Italy Search for more papers by this author Vishnu Dileep Vishnu Dileep Department of Biological Science, Florida State University, Tallahassee, FL, USA Search for more papers by this author Jean-Pierre Quivy Jean-Pierre Quivy Institut Curie, Centre de recherche, Paris, France Centre National de la Recherche Scientifique (CNRS), Unité Mixte de Recherche UMR218, Laboratory of Nuclear Dynamics and Genome Plasticity, Paris, France Search for more papers by this author Rossana Foti Rossana Foti EMBL Mouse Biology Unit, Monterotondo, Rome, Italy Search for more papers by this author Federico Tili Federico Tili EMBL Mouse Biology Unit, Monterotondo, Rome, Italy Search for more papers by this author Rachel Santarella-Mellwig Rachel Santarella-Mellwig EMBL Meyerhof Str., Heidelberg, Germany Search for more papers by this author Claude Antony Claude Antony EMBL Meyerhof Str., Heidelberg, Germany Search for more papers by this author Geneviève Almouzni Geneviève Almouzni Institut Curie, Centre de recherche, Paris, France Centre National de la Recherche Scientifique (CNRS), Unité Mixte de Recherche UMR218, Laboratory of Nuclear Dynamics and Genome Plasticity, Paris, France Search for more papers by this author David M Gilbert David M Gilbert Department of Biological Science, Florida State University, Tallahassee, FL, USA Search for more papers by this author Sara B C Buonomo Corresponding Author Sara B C Buonomo EMBL Mouse Biology Unit, Monterotondo, Rome, Italy Search for more papers by this author Daniela Cornacchia Daniela Cornacchia EMBL Mouse Biology Unit, Monterotondo, Rome, Italy Search for more papers by this author Vishnu Dileep Vishnu Dileep Department of Biological Science, Florida State University, Tallahassee, FL, USA Search for more papers by this author Jean-Pierre Quivy Jean-Pierre Quivy Institut Curie, Centre de recherche, Paris, France Centre National de la Recherche Scientifique (CNRS), Unité Mixte de Recherche UMR218, Laboratory of Nuclear Dynamics and Genome Plasticity, Paris, France Search for more papers by this author Rossana Foti Rossana Foti EMBL Mouse Biology Unit, Monterotondo, Rome, Italy Search for more papers by this author Federico Tili Federico Tili EMBL Mouse Biology Unit, Monterotondo, Rome, Italy Search for more papers by this author Rachel Santarella-Mellwig Rachel Santarella-Mellwig EMBL Meyerhof Str., Heidelberg, Germany Search for more papers by this author Claude Antony Claude Antony EMBL Meyerhof Str., Heidelberg, Germany Search for more papers by this author Geneviève Almouzni Geneviève Almouzni Institut Curie, Centre de recherche, Paris, France Centre National de la Recherche Scientifique (CNRS), Unité Mixte de Recherche UMR218, Laboratory of Nuclear Dynamics and Genome Plasticity, Paris, France Search for more papers by this author David M Gilbert David M Gilbert Department of Biological Science, Florida State University, Tallahassee, FL, USA Search for more papers by this author Sara B C Buonomo Corresponding Author Sara B C Buonomo EMBL Mouse Biology Unit, Monterotondo, Rome, Italy Search for more papers by this author Author Information Daniela Cornacchia1, Vishnu Dileep2,‡, Jean-Pierre Quivy3,4,‡, Rossana Foti1, Federico Tili1, Rachel Santarella-Mellwig5, Claude Antony5, Geneviève Almouzni3,4, David M Gilbert2 and Sara B C Buonomo 1 1EMBL Mouse Biology Unit, Monterotondo, Rome, Italy 2Department of Biological Science, Florida State University, Tallahassee, FL, USA 3Institut Curie, Centre de recherche, Paris, France 4Centre National de la Recherche Scientifique (CNRS), Unité Mixte de Recherche UMR218, Laboratory of Nuclear Dynamics and Genome Plasticity, Paris, France 5EMBL Meyerhof Str., Heidelberg, Germany ‡These authors contributed equally to this work *Corresponding author. EMBL Mouse Biology Unit Monterotondo, Campus Buzzati Traverso, Via Ramarini 32, Monterotondo (RM) 00015, Italy. Tel.:+39 06 90091348; Fax:+39 06 90091406; E-mail: [email protected] The EMBO Journal (2012)31:3678-3690https://doi.org/10.1038/emboj.2012.214 There is a Have you seen? (September 2012) associated with this Article. 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 The eukaryotic genome is replicated according to a specific spatio-temporal programme. However, little is known about both its molecular control and biological significance. Here, we identify mouse Rif1 as a key player in the regulation of DNA replication timing. We show that Rif1 deficiency in primary cells results in an unprecedented global alteration of the temporal order of replication. This effect takes place already in the first S-phase after Rif1 deletion and is neither accompanied by alterations in the transcriptional landscape nor by major changes in the biochemical identity of constitutive heterochromatin. In addition, Rif1 deficiency leads to both defective G1/S transition and chromatin re-organization after DNA replication. Together, these data offer a novel insight into the global regulation and biological significance of the replication-timing programme in mammalian cells. Introduction Replication of the mammalian genome is organized in both space and time. Large segments of the genome, called replication domains, are coordinately replicated through the nearly synchronous firing of clusters of replication origins. Replication domains are visualized as foci and their position in the nucleus as well as temporal order of activation are inherited throughout cell cycles (Jackson and Pombo, 1998; Ma et al, 1998; Dimitrova and Gilbert, 1999b; Sadoni et al, 2004). The molecular and genetic mechanisms underlying such highly orchestrated coordination and inheritance processes are still largely unknown. Transcriptional activity, epigenetic marks and tri-dimensional (3D) chromatin organization have all been proposed to play a role in defining the identity of replication domains and their order of activation (Gilbert and Gasser, 2006; Gondor and Ohlsson, 2009; Hiratani et al, 2009). Yet, the relationships between these inter-dependent processes are poorly understood. A particularly intriguing relationship is between replication timing and transcription, because of the strong correlation between early replication and active gene expression. A long-standing question has been whether transcriptional activity and replication timing are inter-dependently regulated. In some cases, changes in transcriptional activity, for example, induced by developmental transitions, have been shown to correlate with changes in replication timing (Hiratani et al, 2008) but recent genome-wide techniques have revealed that their relationship is not direct (Hiratani et al, 2009). Moreover, interfering with the activity of chromatin modifying enzymes can affect replication timing, but the effect is generally of modest extent and local (Aparicio et al, 2004; Li et al, 2005; Wu et al, 2006; Jorgensen et al, 2007; Goren et al, 2008; Yokochi et al, 2009). To date, no global determinants of the spatio-temporal organization of mammalian DNA replication have been identified, since all the mutations analysed influence either few loci or repetitive sequences such as rDNA or pericentric regions. The most dramatic rearrangements of replication timing thus far reported take place during embryonic stem cells (ESCs) differentiation (Hiratani et al, 2008). The strongest correlation of chromatin properties to replication timing so far observed is to 3D chromatin organization. Indeed, both chromatin interactions and subnuclear position of different replication domains display a high degree of correspondence with their timing of replication (Gilbert et al, 2010; Ryba et al, 2010; Yaffe et al, 2010). After metaphase, chromatin is highly mobile in the nucleus. During G1, upon re-assembly of the nuclear envelope and lamina, chromatin is anchored and assumes defined positions. This moment coincides with the identification and segregation of the replication domains whose origins fire at different times and is therefore called ‘timing decision point’ (TDP) (Dimitrova and Gilbert, 1999a; Leonhardt et al, 2000; Chubb et al, 2002). One plausible theory about the mechanism of timing control proposes the existence of a protein or protein complex that is rate limiting for origin firing as recently identified in budding yeast (Mantiero et al, 2011). In this scheme, at the beginning of S-phase, origins with the highest probability to fire would be the ones with the highest affinity/accessibility for the limiting factor(s). Only upon release of the limiting factor(s) from the early domains, origins with lower affinity could then fire (Rhind, 2006). In line with the idea of regulated access to limiting initiation factors being a key mechanism of replication-timing regulation, it was recently shown that the clustering of origins in subnuclear domains mediated by Fkh1 and 2 is indeed essential for the origin-firing programme in budding yeast (Knott et al, 2012). The anchorage of chromosome domains to the nuclear periphery during G1 could be a means to spatially segregate different genomic compartments either with different protein content and/or to spatially define the accessibility of a limiting factor required for origin firing to a given replication domain. Thus, by regulating interaction with the limiting factor(s), chromatin anchoring could control the temporal order of origin activation. The biological significance of spatial compartmentalization of DNA replication also remains unclear. It may be that this organization reflects the necessity of re-assembling different types of chromatin structures in the wake of the replication fork, creating a temporal and local concentration of specific factors for each chromatin type (Gilbert, 2001). For example, temporal segregation of replication of pericentric heterochromatin (pHC) to mid S-phase could serve the purpose of guaranteeing the availability/concentration of specialized chromatin remodelers and histone chaperones, especially required for assisting fork progression through highly compacted chromatin and for re-establishing the appropriate epigenetic features (Quivy et al, 2008; Loyola et al, 2009; Maison et al, 2010). In this work, we identify Rif1 as the first key component of the molecular machinery that determines the replication-timing programme in mammalian cells. Rif1 was originally identified in budding yeast as a Rap1 interacting protein (Hardy et al, 1992). As for Rap1, Rif1 deletion causes telomere hyper-elongation, suggesting that Rif1 is a telomerase negative regulator. The mechanism by which Rif1 participates in telomere homoeostasis remains unclear. However, it was recently published that budding yeast Rif1 regulates telomere replication timing by temporally restricting telomerase access (Gallardo et al, 2011; Lian et al, 2011). In fission yeast, Rif1 does not bind Rap1, but still interacts with telomeres and participates in their length regulation (Kanoh and Ishikawa, 2001). In mammalian cells, others and we identified Rif1 as a double-strand break (DSBs) response factor (Silverman et al, 2004; Xu and Blackburn, 2004). In addition, we found that Rif1 had a very important role during fork re-start downstream of ATR (Buonomo et al, 2009; Xu et al, 2010). We therefore attributed the effects of Rif1 deletion on S-phase progression to its activity during fork re-start (Buonomo et al, 2009). Both during DSBs response and upon fork stalling, localization of Rif1 at the sites of DNA damage required 53BP1 (Silverman et al, 2004; Buonomo et al, 2009). However, Rif1 is an essential gene (Buonomo et al, 2009), while 53BP1 knockout mice are viable (Ward et al, 2003), prompting our investigation of non-DNA-damage related, 53BP1-independent functions of Rif1. Our work presented here shows that the most rapid and widespread effect of deleting Rif1 is a genome-wide alteration of the temporal order of origin firing. We also describe the consequences of Rif1 deficiency on chromatin organization and cell-cycle progression. Through studying this key regulator, we can therefore provide an insight into the links between replication-timing control, chromatin organization and genome stability. Results Rif1 shows a dynamic S-phase-specific pattern To gain molecular insight into Rif1 function during S-phase, we have generated a knockin mouse replacing the starting Rif1 ATG codon with the coding sequence of the FLAG–HA2 tag, resulting in the Rif1FH allele (Supplementary Figure S1A and B). The functionality of this allele was confirmed by the birth of Mendelian numbers of Rif1FH/FH mice (Supplementary Figure S1C) and their normal health status. Immunofluorescence using an anti-Rif1 antibody in heterozygous Rif1FH/+ mouse embryonic fibroblasts (MEFs) confirmed that the tagged protein fully recapitulated the dynamics of the untagged Rif1 (Supplementary Figure S1D). We therefore employed this endogenously epitope-tagged Rif1 protein to study in detail Rif1 localization during cell cycle by immunofluorescence. We performed our immunofluorescence studies in cells that were pre-extracted with a buffer containing 0.5% Triton X-100. This allowed us to concentrate on the fraction of the protein that is either chromatin-bound or present in the insoluble nuclear fraction. During G1 as well as G2, apart from a variable number of unclassified brighter foci, Rif1 was distributed diffusely through the nucleus (Figure 1A and B). S-phase can be divided in six different stages, each characterized by a specific spatial EdU pattern (Supplementary Figure S2) (Dimitrova and Berezney, 2002; Quivy et al, 2004). S1–3, S4–5 and S6 represent early, mid and late S-phase, respectively. In S1/2, Rif1 was present throughout the nucleus, although its distribution was not homogeneous and it never co-localized with the replication fork (Figure 2A, S1/2) or with the replicative helicase MCM3 (Figure 1A, S1). In S3, the latter part of early S and prior to the appearance of the replication forks within pHC, Rif1 was specifically enriched at chromocenters (Figure 2A), where it co-localized with MCM3 (Figure 1A, S3). As previously described (Quivy et al, 2004), replication of pHC occurs at the outer edges of the chromocenters and once replicated, the DNA moves to the interior of the chromocenter. During the initial steps of pHC replication (S4), Rif1 localization was restricted to the innermost of this structure, which is yet to be replicated, appearing always closely juxtaposed to the EdU but never overlapping with it (Figure 2B). In S5, Rif1 no longer co-localized with the fully replicated chromocenters (Figure 2A and B). In S6, Rif1 signal had again become more diffuse. These data show that Rif1 displays a highly dynamic and specific S-phase behaviour. Interestingly, Rif1 never co-localizes with the replication fork, but on the contrary, clearly precedes it, at least at pHC during mid S-phase. Figure 1.Rif1 localization during G1 and G2. (A) Confocal microscope images of cells in different stages of G1 and early S identified by the intensity and distribution of MCM3 signal and EdU pattern. During G1, Rif1FH (anti-HA, green) shows a diffuse nuclear staining (DAPI, blue). It co-localizes with MCM3 (red) only during the latter part of early S (S3). EdU is shown in Cyan. Scale bar: 10 μm. (B) Cells in late S-phase and different G2/M stages were identified by staining for phosphorylated Ser10 histone H3 (H3S10ph, red). Rif1FH (anti-HA, green) returns to its diffuse localization during G2 (DAPI, blue). EdU is shown in Cyan. Scale bar: 10 μm. Download figure Download PowerPoint Figure 2.Dynamic Rif1 localization during S-phase. (A) Confocal microscope images of fluorescently stained newly replicated DNA (EdU, red), immunofluorescence for Rif1FH (anti-HA, green) and DAPI (blue) in Rif1FH/FH MEFs. Scale bar: 10 μm. (B) Rif1 precedes replication at chromocenters. Magnifications of insets highlighted from (A). Scale bar: 2 μm. Download figure Download PowerPoint Rif1 is required for the regulation of replication timing Rif1's localization ahead of the replication fork, marking domains whose origins have yet to be activated, prompted us to investigate spatio-temporal replication patterns in Rif1 null cells. To this end, we have taken advantage of the Rif1 conditional allele (Rif1F) (Buonomo et al, 2009) to induce acute Rif1 deletion in logarithmically growing early passage Rif1F/F pMEFs (Supplementary Figure S3A). Upon CRE infection of Rif1F/F pMEFs, cells were pulse-labelled with EdU and the six replication patterns typically observed in mouse cells were identified. A low but reproducible percentage of Rif1−/− cells displayed an aberrant EdU pattern that we classified as mixed S2–S4 (Figure 3A). A similar observation was reported in Suv39h1/2 double-null MEFs (Wu et al, 2006), where it was shown that the appearance of this aberrant pattern coincided with advanced replication timing of the major satellites. We next performed genome-wide profiling (Hiratani et al, 2008) of replication timing in Rif1+/+ and Rif1−/− pMEFs. Several cell cycles following CRE-mediated deletion of Rif1, the replication-timing programme was dramatically re-organized (Figure 3B and C), to an extent substantially greater than that seen during ESCs differentiation (Figure 3C) (Hiratani et al, 2008), the most dramatic re-organization observed to date. The correlation between Rif1+/+ and Rif1−/− pMEFs was reduced to an average of 0.63, yet the correlation of replicates between two independent lines was 0.90 (Supplementary Figure S3B and C), demonstrating that the disrupted pattern upon Rif1 loss was reproducible. Rif1 deletion caused both late-to-early (LtoE) and early-to-late (EtoL) switches of replication timing (Figure 3D and E), with 40% more domains switching from EtoL than LtoE timing (Figure 3C). The distribution of replication-timing values in Rif1 null cells clustered near the middle of S-phase (Figure 3F), but due to the nature of our genome-wide analysis, we cannot distinguish whether these regions are replicating in mid-S, replicating randomly throughout S-phase, or whether the two homologues are replicating at specific but asynchronous times. Figure 3.Rif1 deficiency deregulates the replication-timing programme. (A) EdU staining (green) in Rif1−/− pMEFs reveals a mixed S2–S4 pattern as compared with wild-type cells (Figure 2, S2). A pan-nuclear EdU signal excluding nucleoli typical of S2 co-exists with the characteristic S4 pattern, identified by the typical EdU rings around the chromocenters. DAPI (blue). Scale bar: 10 μm. (B) Hierarchical clustering and correlation matrix heat map comparing replication-timing data (each an average of two bio-replicates) from pMEF Rif1+/+ and Rif1−/− to different cell types (previously published ESC, neural progenitor cells-NPC, Myoblasts, induced pluripotent Stem cells-iPSC) along with published wild-type MEF and two replicates of pMEF Rif1+/+ and Rif1−/−. (C) Table summarizing the percentages of changes comparing previously published wild-type MEFs with Rif1+/+ pMEFs and the latter with littermate Rif1−/− pMEFs. *As a reference, the % changes during ESCs differentiation into NPCs from Hiratani et al (2008) is shown. The percentage of changes is calculated as the number of probes on the array that change by a factor of more than 1 versus the total number of probes. (D) Exemplary regions (shaded) in chromosome 17 (left) and 3 (right) whose replication-timing switches from LtoE in Rif1−/− (red) compared with Rif1+/+ (black) pMEFs, shown by Loess smoothed replication-timing profiles. Profiles for the same region from male and female published wild-type MEFs are shown in green. (E) Exemplary regions (shaded) in chromosome 3 (left) and 5 (right) whose replication-timing switches from EtoL, depicted as in (D). (F) Distribution of replication-timing values in Rif1−/− versus Rif1+/+ cells. Two independent lines are shown for each genotype. (G) Comparison of early (blue) and late (red) domain size distribution between Rif1+/+, Rif1−/− and ESCs (*ESCs from Hiratani et al, 2008). Download figure Download PowerPoint Interestingly, one striking effect of Rif1 deficiency on replication-domain structure was their fragmentation in size (Figure 3G). ESCs have the smallest replication domains observed among all cell types that have been analysed thus far, which consolidate into larger domains during differentiation (Hiratani et al, 2008; Ryba et al, 2010). Rif1 deletion in pMEFs induced fragmentation of replication-domain sizes into a total number even higher than in ESCs (1789 domains in ESCs versus 1425 in Rif1+/+ and 1956 in Rif1−/−pMEFs). These data show that Rif1 deficiency induces dramatic changes in the temporal replication programme, to an extent not matched by the disruption of 3D spatial replication patterns. However, during the analysis of the spatial organization of DNA replication by EdU staining, we observed that there were about 50% more Rif1 null cells displaying an early S-phase pattern (Figure 4A). This result could be interpreted as an actual accumulation of cells in the early stages of S-phase, reflecting a temporal delay in the early-to-mid S-phase transition. Alternatively, in many cells the pattern that we had identified as early S could result from a loss of spatial organization as well as a fragmentation of the replication domains. The early S-phase pattern is indeed characterized by a diffuse localization and by the presence of numerous small replication foci. To distinguish between these two possibilities, we quantified the distribution of Rif1−/− cells among different S-phase stages by flow cytometry, relying on DNA content, rather than visual inspection (Figure 4B) to assess whether the cells were delayed in early S-phase. We subdivided S-phase into three equal fractions according to DNA content and determined the percentage of total 5′-bromo-2′-deoxyuridine (BrdU)-positive S-phase cells found in each fraction for wild-type versus Rif1−/− pMEFs. This analysis revealed comparable progression through S-phase for cells of both genotypes. We can therefore conclude that the visual accumulation of cells with a diffuse early-S-like EdU pattern in the Rif1−/− cells results from spatially and temporally disorganized replication, potentially due to fragmentation of replication domains. Figure 4.Consequences of Rif1 deletion on spatial replication-foci distribution. (A) In the absence of Rif1, cells with early replication spatial patterns accumulate. Upon 30′ EdU pulse, cells were fixed and stained for EdU. S-phase substages were evaluated by visual inspection of the cycling population. Pie charts show the relative proportion (percentage of total S) of early, mid and late S-phase. Cells were scored blinded for two independent Rif1+/+ and four Rif1F/F treated with CRE or EV pMEF clones. In all, 200 EdU+ cells were counted for each clone. Averages are shown. P-values early P<0.0001, mid=P<0.0001, late P<0.01. (B) Six independent Rif1+/+ and Rif1F/F pMEF clones treated with CRE or EV were pulse-labelled with BrdU and the percentage of positive cells was evaluated by FACS analysis. S-phase was subdivided into three equal fractions of increasing propidium iodide content to define early, mid and late S-phase. The ratio of the percentage of total BrdU-positive cells found in each S-phase fraction between EV and CRE treated was plotted. Download figure Download PowerPoint Surprisingly, we found that Rif1 deletion had no major effect on the cell's transcriptome (Supplementary Figure S3D), suggesting that the effect of Rif1 deletion on replication timing is neither mediated by nor causing gross changes in the transcriptional landscape. However, our transcriptome analysis has been performed on the population of cycling Rif1−/− cells. Therefore, we cannot formally exclude that a small number of cell-cycle specific genes could be deregulated as a consequence of Rif1 deletion. The effect of Rif1 deficiency on the mammalian replication-timing profile that we report here is the strongest observed to date, indicating that Rif1 plays a key role in the control of the spatio-temporal organization of the replication origin-firing programme. Rif1 deficiency disrupts repackaging of newly replicated chromatin It has been speculated that the spatio-temporal organization of DNA replication could facilitate the repackaging of different types of chromatin after fork passage. Thus, one possible consequence of replication-timing deregulation could be disorganized chromatin assembly (Quivy et al, 2008; Maison et al, 2010). In agreement with this prediction, one of the most striking phenotypes associated with Rif1 deficiency is the presence of what we have defined ‘fluffy’ newly replicated chromocenters. In Rif1 null cells, the EdU-labelled DNA around the chromocenters appeared highly disorganized (Figure 5A) in comparison to wild-type controls, suggesting a defective pHC repackaging/assembly. However, none of the known post-translational histone modifications associated with heterochromatin analysed showed abnormal localization on newly replicated pHC by immunofluorescence, or overall levels by western blotting (Supplementary Figure S4A and B). We also analysed the general organization of newly replicated DNA by examining its accessibility to Micrococcal nuclease (MNase). For the same extent of total DNA digestion (visualized by ethidium bromide staining), a stronger BrdU signal in Rif1−/− pMEFs is detected (Figure 5B; Supplementary Figure S4C), suggesting increased accessibility of newly replicated DNA in Rif1−/− pMEFs. In summary, Rif1 deletion causes both an alteration of the replication-timing programme and the disorganization of chromatin re-assembly after the passage of the replication fork. Figure 5.Rif1 deficiency affects re-organization of newly replicated chromatin. (A) Rif1 deletion impairs post-replicative 3D re-organization of pHC. EdU staining (green) of cells in mid S-phase shows disorganization of newly replicated pHC in Rif1−/− cells. Scale bar: 10 μm. (B) BrdU pulse-labelled chromatin from two independent Rif1+/+ and three Rif1F/F; Rosa26CreERT2/+ (Rif1−/−) pMEFs treated with 4-hydroxytamoxifen was digested with different dilutions of MNase. DNA was extracted and used for Southern–Western with an anti-BrdU antibody (upper panel). The lower panel shows ethidium bromide staining of the agarose gel with the total DNA from MNase digested chromatin used for the Southern–Western. All the pMEFs lines showed a BrdU+ population around 4%, as quantified by FACS (not shown). The right panel shows the quantification of the BrdU immunoblot (WB) normalized for the major satellite Southern signal (SB) (Supplementary Figure S4C). U=enzymatic units. Download figure Download PowerPoint Replication-timing deregulation does not affect inter-origin distances or fork progression The profound effect of Rif1 deletion on replication timing offered the opportunity to investigate how alteration of this programme is related to DNA replication dynamics, namely the frequency of origin firing and fork progression speed. Upon CRE infection, we labelled Rif1-deficient and control pMEFs with consecutive pulses of IdU followed by CldU to visualize individual replication tracks by DNA Dynamic Molecular Combing (Michalet et al, 1997). Inter-origin distances were not significantly altered (Figure 6A) while replication fork speeds showed a minor increase in Rif1 null cells (Rif1+/+ 1.98 kb/min and Rif1−/− 2.16 kb/min), mostly resulting from a reduction in the abundance of slower replicating forks (Figure 6B and C). Moreover, we could not detect any difference in the number of collapsed or uni-directional forks, nor, by flow cytometry, in the amount of BrdU incorporated per cell (data not shown). In DT40 cells, it has been shown that Rif1 knockout causes an increase in the number of uni-directional replication forks (Xu et al, 2010). However differences both in cell type and system used could account for this discrepancy. Our data indicate that the correct establishment of the replication-timing programme is independent from and has no major impact on fork progression and frequency of origin firing. Figure 6.Rif1 deficiency does not affect the frequency of origin-firing or replication fork speed. (A) Boxplot diagram of inter-origin distance measured on individual DNA fibres in Rif1−/− (n=38) and Rif1+/+ cells (n=53). (B) Boxplot diagram of replication fork speed measured on individual DNA fibres in Rif1−/− (n=309) and Rif1+/+(n=392). P-value=3,75 10−5. (A, B) Bottom and top of the box indicate the upper and lower quartiles, respectively, the line in the box the median and the whiskers the 1.5 IQR of the lower and upper quartile. (C) Fork speed distribution in Rif1−/− and Rif1+/+ cells. Download figure Download PowerPoint Replication-timing deregulation is a primary consequence of Rif1 deficiency In order to determine whether replication-timing deregulation was upstream or downstream of the chromatin-repackaging defect, we examined replication timing in the first S-phase following Rif1 deletion. Rif1F/F pMEFs were arrested in G0, deletion was induced (Supplementary Figure S4D)