Présentée Par Baraah

A L Ahmad

Nachar Le

Devant Le

Valeria Mme

CRCN Naim

MCU Jean-Charles Cadoret

Pr Anne-Marie Martinez-Cavalli

DR Mme Chantal Desdouets

M Vincent

Thèse Pour

Obtenir Le Grade De

Docteur De L'université De Montpellier

Marta Bialic

Baraah Al

Ahmad Nachar

Vincent Coulon

Jean-Marc Lemaitre
email: jean-marc.lemaitre@inserm.fr

Etienne Schwob
email: etienne.schwob@igmm.cnrs.fr

Sous la direction de Etienne SCHWOB

Keywords: DNA replication, origin licensing, DNA combing, S-phase duration, MiDAS, chromosome instability, Palbociclib DNA replication, origin licensing, DNA combing, S-phase duration, MiDAS, chromosome instability, Palbociclib ESC, MEF, DNA combing, global instant fork density, DNA replication, S-phase duration, fork pausing, dormant origins, chromatin state, genome segmenting

Why solid tumours show high chromosome instability is still poorly understood. Seminal work in the host lab has shown in the yeast model system that precocious CDK activation and reduced origin licensing in G1 cause S-phase extension and chromosome rearrangements in mitosis. Since most cancer cells have genetic or epigenetic alterations in one or more G1/S cell cycle regulators that can impede origin licensing, we analysed chromosome replication dynamics in fourteen human epithelial cancer cell lines using newly developed techniques, and compared it to normal human fibroblasts, mammary and retinal pigment epithelial cells. Our results show that all cancer cells spend longer time in S phase (10-29h) than untransformed cells (7-9h). Interestingly, most cancer cell lines displayed a lower global instant density of replication forks (GIFD), partly compensated for some cell lines by increased fork velocity (FV). We define replication potency (RP = GIFD x FV) as a new descriptor of cells' capacity to synthesize DNA that integrates this compensation mechanism, and found that it was lower for cancer cell lines.

The consequences of this longer S phase on the cell cycle and mitosis were assessed by 4D microscopy. We detected mitotic DNA synthesis (MiDAS) and chromosome segregation failures in cancer cells not treated with replication drugs, indicating constitutive chromosome instability (CIN). Importantly, the low GIFD and long S phase of pRb+ cancer cells was reversed by slightly extending G1 using low dose of the CDK4/6 inhibitor Palbociclib. Our data strongly suggest that S-phase extension due to lowered origin licensing in G1 is a common feature and perhaps the main trigger for the genomic instability in cancer cells.

Résumé

L'instabilité chromosomique est une propriété cardinale des tumeurs solides dont on connait encore mal l'origine. Les travaux pionniers effectués au laboratoire ont montré, dans le système modèle levure, que l'activation précoce des kinases CDK et la formation d'un nombre insuffisant d'origines de réplication ('licensing') en G1 conduisent à une extension de la phase S et à des remaniements chromosomiques en mitose. La plupart des cellules cancéreuses ayant des altérations génétiques ou épigénétiques d'un ou plusieurs régulateurs G1/S du cycle cellulaire qui peuvent entraver la formation des origines, j'ai analysé à l'aide de nouvelles techniques la dynamique de réplication de l'ADN dans 14 lignées de cellules cancéreuses épithéliales humaines, comparativement à celle de fibroblastes humains normaux. Mes résultats montrent que toutes les lignées cancéreuses testées ont une phase S plus longue (10-29h) que les cellules non transformées (7-9h). Les cellules cancéreuses ont aussi une densité instantanée globale de fourches de réplication (GIFD) plus faible, partiellement compensée pour certaines lignées par une vitesse de fourche (FV) accrue. Je définis la force de réplication (RP = GIFD x FV) comme un nouveau descripteur de la capacité des cellules à synthétiser l'ADN, qui intègre ce mécanisme de compensation et qui est systématiquement plus faible pour les lignées cancéreuses.

Les conséquences de cette phase S allongée sur le cycle cellulaire et la mitose ont été évaluées par microscopie 4D. J'ai détecté une synthèse d'ADN en mitose (MiDAS) et des défauts de ségrégation des chromosomes dans les cellules cancéreuses non exposées à des perturbateurs de la réplication, indiquant la présence une instabilité chromosomique (CIN) constitutive. Un traitement avec des doses faibles de palbociclib, un inhibiteur des kinases de G1 CDK4/6, restaure une GIFD et durée de phase S normales dans des cellules cancéreuses pRb+. Nos données indiquent que la majorité des cellules tumorales peinent à dupliquer leur génome à cause d'une réduction de nombres d'origines actives, et que ceci pourrait être le déclencheur principal de leur instabilité génomique. Elles ouvrent aussi de nouvelles pistes pour le dépistage précoce ou le traitement des cancers.

Mots-clés : Réplication de l'ADN, 'licensing' des origines, peignage moléculaire de l'ADN, durée de phase S, MiDAS, instabilité chromosomique, palbociclib. A warm deep thank you goes to my lab members for being that amazing and for the great environment we have in the lab. Thank you Etienne, my director, for trusting me and for giving me the opportunity to grow up as an independent and strong scientist in addition to the valuable discussions that improved my critical thinking. Thank you also for your warm welcome and for your support at the human level. Thank you Vincent, my co-supervisor, for your scientific help and mainly for the psychological support especially when I get emotional over experiments.

Thank you for being a good listener and for absorbing my stress both at the scientific and personal levels. My combing goddess, Marjorie, thank you for all the moments we had together and for your help in my project. I know I am bossy sometimes, but you were kind enough to handle it. Thank you for all the calls, emails and letters you helped me with. You made my life much easier with French administration. You also made work so fun in addition to being supportive. Thank you Nicolas for giving me your time when you feel I am lost or angry and for the advices and the valuable discussions we had. Elisabeth, thank you for forcing me to practice my French and for the good times we used to have at lunch. Thank you Philippe for helping me optimizing and doing some experiments and Vjeko for lab meetings discussions. Thank you Juande for being a labmate friend who eased my stress and listened to all my shit. Thank you for the "bitching times" we used to have with Vincent. Michela, you also made working in the lab so fun and you handled my last year stress with Juande. A special thanks also goes to Laura, Theresa and Léna too. Isabelle, I know we did not have a lot of time to know each other but thank you for being kind and for practicing my French ;) Thank you for all the team for making it possible for me to reach this level you were literally my family in France. Thank you all IGMM members for making working at the institute so special! I am also so lucky and fortunate to have great and supportive friends though friend word is too little to describe what you have done to me over these four years. Without you guys life would have been much harder. Thank you Charbel and Jona for your support. Thank you for all the lovely kebbe we used to have because of you Jacinthe. Thank you Adham for organizing the special pots and for making life in the institute much more fun. Thank you Fabienne for being you and for your help. Thank you Fatima for your support as well. Hasan (ya 7elo), you would listen to my shit and know what is going on even without me telling you. Special thanks go to Hussein, my best gym partner ever! who pushed me to relief my stress through exercising. Great thank you goes to you Amani for handling all my stress, all my anxiety and my questions both at work and personal levels. You took the friendship level so high! Thank you Azam, David, Igor, Lorène, Fatimat, Mathias, Hulya and Monica for making pots and outings so specials. Thank you Marta for your help in culturing mouse ESC and for our discussions.

Special thanks go to you Rana for all your support and love! You too Razan J Thank you ladies for the good moments. Sarwat I would never forget the taste of your mlo5ye! Maha my ex bench mate, thank you for making Paris visits so special. I cannot move on without thanking Francoise and Sarah for their help in the administrative paper work and Virginie, Myriam and Stephanie for their help in the MRI facility.

Thierry, thank you for all the statistical analysis and for the good discussions we had at lunch! Finally, I would like to thank my family for all their support. Warm thanks go to my sister, Mariam who continuously supports and defends me. And you mom, thank you for your strength because you passed it to me. Thank you Dad! I dedicate this work to you.

Kinases

The cell cycle is composed of consecutive controlled steps by which the mother cell duplicates its genome and segregates identical copies of chromosomes into two genetically identical daughter cells. The two main steps of the cell cycle are the S phase (synthesis phase) where genome duplication occurs and the M phase (Mitosis) where chromosomes segregate.

Two additional phases take place before and after S phase called G1 and G2 respectively. The control of the cell cycle through several phases allows the cell to ensure complete DNA replication before undergoing cellular division. The transition from one phase to another is regulated by cyclins and cyclin-dependent kinases (CDKs) (Figure 1). In each cyclin / cdk dimer, the cyclin abundance fluctuates and it constitutes the regulatory subunit, while the cdk moiety possesses the kinase activity. In addition to these phases, most metazoan adult cells remain in a resting state without entering the cell cycle; they are referred to as quiescent or G0 cells. In 1974, Arthur Pardee suggested that untransformed cells have a restriction point past which cells irreversibly enter S phase (Pardee, 1974). Normal cells shift from proliferative to quiescent state under nutritional deprivation or absence of mitogens as a survival mechanism; however malignant cells lack this restriction point. In the presence of mitogens, retrovirusassociated sequence (Ras) pathway is activated leading to the expression and stabilization of cyclin D. The association of cyclin D with these CDK4/6 triggers its activation which plays a role in G1 phase. Regulation of G1 phase of the cell cycle occurs through several mechanisms.

One of them is through the retinoblastoma protein (RB) and its two paralogs p107 and p130.

However, if the restriction point is not passed in untransformed cells, cells enter G0 phase.

1.1.1.1.G0 Phase

The G0 phase is characterized by increased expression of the DREAM (DP, Rb-like, E2F

and Muv) complex which represses cell cycle entry. E2F4-DP1 and Muv B components of DREAM complex bind to promoters containing E2F and CHR elements, responsible for early and late cell cycle gene promoters respectively, inhibiting their expression (Litovchick et al., 2007). A recent study has shown that cyclin D-CDK4 is necessary for phosphorylation of P130, a component of the DREAM complex, triggering its dissociation and complex inactivation. This relieves the inhibition by DREAM on promoters involved in cellular proliferation (Schade et al., 2019).

1.1.1.2. G1 Phase Upon cells commitment to S phase entry, cells irreversibly enter the cell cycle. G1 phase is the phase of the cell cycle by which DNA is prepared for replication. The presence of RB protein in an hypophosphorylated state in G1 allows it to bind to E2F1-4 transcription factors, inhibiting their transcriptional activity. Cyclin D-CDK4/6 phosphorylates RB through its Cterminal region leading to the release from E2F factors. E2Fs then activate genes involved in Sphase entry including cyclin E. Cyclin E then binds and activates CDK2, inducing further phosphorylation of RB and full inactivation. As a result, E2Fs are further released (Figure 1) (Harbour et al., 1999).

Another mechanism by which G1 phase is regulated is through Cip/Kip proteins interactions with cyclin-Cdks. p21/p27 members of the Cip/Kip family of cyclin-dependent kinase inhibitors bind and inhibit cyclin E-Cdk1/2 in proliferating mouse fibroblasts. Moreover, these proteins play the role of a bridge enhancing cyclin D association to its Cdks and its nuclear transportation (Cheng et al., 1999). The binding of Cip/Kip proteins to cyclin D-Cdk titrates Cip/Kip leading to the relief of cyclin E-Cdk2 suppression inducing cell cycle progression. This explains the oncogenic activation caused by p27 sequestering cyclin D-Cdk (Bouchard et al., 1999). However, this inhibitory effect is dependent on the phosphorylation state of Cip/Kip proteins which modulates their interactions with cyclins-CDK complexes. P27 Kip1 was shown to be associated with cyclin-D-CDK4 in both proliferating and non-proliferating cells. However, the outcome of this association is different in both cases. P27 Kip1 phosphorylation at tyrosine Y88 and Y89 on the N-terminus is seen in proliferating cells where p27 does not act as an inhibitor.

Whereas loss of these phosphorylations enhance p27 inhibitory effect seen in non-proliferating cells (James et al., 2008). Beside its direct interaction with cyclins, p21 was shown to act as a transcription repressor enhancing the repression activity of RB-E2F complex (Delavaine and La Thangue, 1999).

S Phase

The increase in cyclin E levels triggers G1/S transition, and the cells enter S phase to replicate the genome. DNA replication is a dynamic process that involves complex enzymes that induce several genomic events including chromatin changes, DNA unwinding and epigenetic modifications. In addition, cyclin A was shown to be localized in the nucleus during S phase till G2 (Girard et al., 1991) and cyclin A precocious expression in G1 triggers early G1/S transition (Resnitzky et al., 1995). In addition, cyclin A-CDK2 interacts with components of DNA replication complex, mainly minichromosome maintenance-7 (MCM-7) promoting pre-initiation complex formation (pre-IC) (Chibazakura et al., 2011). The association of cyclin A with CDK2

indicates DNA synthesis.

G2/M Phases

The association of CDK1 with cyclin A marks G2 phase. The presence of checkpoint kinase 1 (CHK1) protein in basal levels during the S phase inhibits mitotic entry. However, the modulation of CHK1 levels by cyclin A-CDK1 at the end of S phase allows cells to undergo mitosis (Oakes et al., 2014). The reduction in CHK1 levels releases the inhibition on cyclin B by CHK1, leading to an increase in cyclin B-Cdk1 levels upon mitotic entry. Greatwall kinase 1 (Gwl) associates with cyclin B-cdk1 activity, allowing mitotic spindle assembly and then cellular division (Hara et al., 2012).

E2F-dependent Roles of RB

Understanding how alteration in the RB pathway induces carcinogenesis leads to crucial insights about cancer properties as it is one of the most commonly altered pathways in cancer cells. As discussed in section 1.1.1.2. RB plays a major role in the regulation of the transition between G1 and S phases. Thus, RB is involved in cell cycle regulation and cellular proliferation. Loss of RB in this case induces oncogene-induced DNA damage caused by premature expression of cyclin E. The details of how cyclin E in this case acts as an oncogene will be discussed in section 3.1.4.

Other roles of RB have also been highlighted and reviewed suggesting the importance of this multifunctional protein in preventing genome instability (Dick and Rubin, 2013). Loss of RB induces overexpression of mitotic arrest deficient 2 (MAD2) gene leading to overactivation of spindle-assembly checkpoint or failure in detachment of microtubules from sister chromatids in anaphase. This would result in chromosome mis-segregation. Furthermore, overexpression of E2F due to RB loss induces premature S-phase entry, alteration in the nucleotide pools, replication stress and double strand breaks. The mechanism of nucleotide pool alteration will be developed in later section. It has been shown that RB is recruited to DNA double-strands breaks in ataxia-telangiectasia mutated (ATM)-dependent manner. RB also stabilizes Topoisomerase IIbinding protein1 (TopBP1) and recruits BRG1 proteins involved in double-strands breaks repair (Vélez-Cruz et al., 2016). All of these data suggest several roles of RB in maintaining genome stability, thus it acts as a tumor suppressor gene to prevent genome instability, a hallmark of cancer.

E2F-independent Roles of RB

In addition to E2F-dependent roles of RB, RB also acts on preserving genome stability in E2F-independent manner. RB depletion alters condensin II and cohesin loading to chromosomes leading to chromosomes mis-segregation and genome instability (Manning et al., 2010). RB was shown to be required for H4-K20 trimethylation at pericentric heterochromatin. Epigenetic changes induced by RB depletion altered cohesin affinity to pericentric regions. This induced altered chromatin structure, chromosomes fusion and lagging chromosomes (Manning et al., 2014). These data suggest the role of RB as tumor suppressor gene in preventing genome instability as well. However, alteration the RB pathway is not restricted only to RB depletion. It also includes epigenetic modifications that alter RB expression. Promoter hypermethylation is known to downregulate the expression of genes. CCCTC-binding factor (CTCF) binding to RB promoter prevents DNA methylation allowing the expression of RB gene. In the absence of CTCF, the RB promoter is silenced. This suggests a possible contribution of CTCF in maintaining an optimal chromatin state that allows RB expression (De La Rosa-Velázquez et al., 2007).

Effect of Palbociclib, a CDK4/6 Inhibitor, on the G1/S Transition

The oncogenic consequence of cyclin D deregulation has been reviewed and highlighted in several types of cancer (Musgrove et al., 2011). In addition, previous data from the lab have emphasized on the importance of G1/S regulation in preventing incomplete replication and chromosomal aberrations (Lengronne and Schwob, 2002a) and (Coulon et al in preparation).

This emphasizes on the importance of targeting G1/S regulation in cancer therapy. PD0332991

or palbociclib has been described as pyridopyrimidine compound, a potent selective inhibitor of CDK4 and CDK6 (Fry et al., 2004) and one of the FDA-approved treatments for several solid tumors and leukemias (Schwartz et al., 2011). The key challenge in using palbociclib in treating tumors is selecting those that do not harbor genomic or functional loss of RB, otherwise they are expected to be resistant to CDK4/6 inhibitors (Figure 2) (Condorelli et al., 2018). Treatment of human fibroblasts with palbociclib was shown to inhibit cell cycle progression through the inhibition of CDK4 kinase. As a result, RB and p130 phosphorylation was reduced, leading to a reduction in the expression of early cell cycle genes including CDC6 and MCM5 (Schade et al., 2019).

Molecular Mechanisms and Regulation of DNA Replication

Control of Origins of Replication during the Cell Cycle

In order to ensure equal genetic inheritance to progeny cells, initiation of replication at origins is highly regulated throughout the cell cycle. In yeast, DNA replication was shown to be initiated at conserved genomic sites called autonomously replicating sequences (ARS) (Stinchcomb et al., 1979). In 1994, Diffley and his colleagues have shown that origins of replication are present in two different states during the cell cycle: pre-replicative and post-replicative. The pre-replicative state appears at the end of mitosis till the beginning of S phase and post-replicative during S, G2 and mitosis. DNA footprint studies at ARS showed that some proteins are recruited to origins of replication in the pre-replicative state to allow further protein recruitment during S phase (Diffley et al., 1994b). The loading and activation of origins are controlled through two exclusive steps in order to ensure that each origin is replicated only once per cell cycle. The first step occurs in G1 phase where replication proteins are loaded to origins, and it is referred to as origin licensing step. The second step occurs in S phase where only licensed origins could be activated to initiate DNA replication, and it is referred to as origin activation step.

Licensing for Replication in G1

The mechanism controlling DNA replication licensing is conserved in yeast and mammalian cells but with some significant differences. Several proteins are recruited to ARS to form pre-replication complexs (pre-RCs) including origin recognition complex (Orc) (Bell and Stillman, 1992), cell division cycle 6 (Cdc6) (Cocker et al., 1996), cdc10-dependent cell division cycle transcript 1 (Cdt1) (Hofmann and Beach, 1994) and mini-chromosome maintenance MCM (Chong et al., 1995).

Previous studies in Saccharomyces cerevisiae showed that a multiprotein complex specifically binds ARS and initiates DNA replication in ATP-dependent manner. Such complex was named Orc (Bell and Stillman, 1992). The ORC complex is a ring-shaped heterohexamer consisting of several subunits including ORC1, ORC2, ORC3, ORC4, ORC5 and ORC6 that have been identified in yeast, Xenopus laevis and mammalian cells. It has been shown to be associated to chromatin throughout the cell cycle. However, in some species such as Xenopus laevis ORC complex relocalizes to the cytoplasm during mitosis for a brief period of time. The association of ORC to the chromatin in G1 provides a landing pad for preRC (Figure 3, a) (Lygerou and Nurse, 1999;Romanowski et al., 1996).

Furthermore, Cdc6 expression in Saccharomyces cerevisiae was shown to be variable throughout the cell cycle, and it reaches its peak in late G1 then it declines during DNA replication and mitosis. It was proven to have an essential role in DNA replication. Cdc6 mutant studies showed that mutant cells fail to replicate their genome, and they still undergo reductional division with unreplicated chromosomes. This emphasizes the role of Cdc6 loading in G1 that is required to activate replication later during S phase (Piatti et al., 1995). Cdc6 and Cdc18 are the orthologs of the yeast Cdc6 in mammalian cells and S. pombe respectively (Williams et al., 1997). Cdt1 was also shown to be an essential gene which expression peaks at the G1/S boundary. Deletion of Cdt1 from fission yeast led to cell cycle arrest and DNA replication inhibition, emphasizing its role in initiating DNA replication. The mammalian ortholog of CDT1 has also been identified, and was shown to be regulated during the cell cycle with an accumulation in G1 and absence upon S phase entry (Nishitani et al., 2001).

CDC6 recruitment to ORC and its regulation throughout the cell cycle is different in yeast and mammalian cells. Cdc6 was shown to be degraded upon G1/S transition in yeast.

However, mammalian CDC6 was shown to be constant during the cell cycle with a decrease in early G1 due to its degradation in mitosis. The subcellular localization of CDC6 changes during the cell cycle: it was shown to be nuclear in G1 then CDC6 translocates to the cytosol after the G1/S transition through phosphorylation by cyclin A-CDK2. However, a defined amount of CDC6 remains bound to the chromatin during S and G2. Loading of CDC6 during G1 was shown to be essential for human mini-chromosome maintenance (MCM) complex loading.

MCM is another protein involved in pre-RC formation. This complex consists of a heterohexamer of related proteins MCM2 to MCM7 (reviewed in Tye, 1999). The essential role of MCM2-7 in DNA elongation during S phase has been demonstrated in Saccharomyces cerevisiae where MCM2-MCM-7 helicase is involved in initiation and fork progression (Labib et al., 2000). However, MCM loading occurs only when cyclin B is degraded suggesting that full origin licensing requires the absence of cyclins during G1 (Méndez and Stillman, 2000).

Both CDC6 and CDT1 were shown to be essential for MCM loading during G1 in Xenopus eggs. CDC6, CDT1 and MCM are loaded sequentially in this order to form the pre-RC. In addition, CDT1 immunodepletion from Xenopus eggs did not alter ORC-CDC6 loading.

However, it reduces the competence of MCM loading, highlighting the essential role of CDT1 on MCM loading during licensing. CDT1 was also shown to be destabilized from the chromatin once the MCM complex is loaded and pre-RCs are formed (Maiorano et al., 2000).

Overexpression of Cdt1 in vivo was later shown to induce origin re-replication and lethality in both embryonic and adult intestinal mouse cells. This was accompanied with an increase in apoptosis highlighting on the importance of Cdt1 regulation in maintaining proper DNA replication (Muñoz et al., 2017).

Following Orc binding, Cdc6 associates with ORC and recruits Mcm2-7-Cdt1. How the second Mcm2-7 is loaded on the DNA is proposed to happen through two models. The first one suggests that two Orc-Cdc6 complexes recruit Mcm2-7 independently and they translocate on the origin (Coster and Diffley, 2017). The second suggests that a single Orc complex loads the first Mcm2-7 followed by the second (Ticau et al., 2015). Both Mcm2-7 complexes encircle DNA in a head-to-head orientation and bidirectionally leave the origin upon activation in opposite directions. Champasa and colleagues confirmed the latter model through singlemolecule studies providing evidence that the loading of the second Mcm2-7 is facilitated through interaction with the first -Mcm2-7. The formation of a double hexamer is essential for DNA unwinding and replication bubble formation (Champasa et al., 2019).

Activation of Licensed Origins for Replication

After origins have been licensed in G1, a subset of them is chosen to be activated in S phase by G1/S kinases: CDK and Dbf4 dependent cdc7 kinase (DDK). DDK and CDK then phosphorylate and allow the loading of several proteins to pre-RCs to form pre-ICs. These proteins include MCM10, REQL4, CDC45, GINS and treslin (Xu et al., 2009). Both DDK and CDK phosphorylate the MCM double hexamers inducing their dissociation and DNA unwinding (Figure 3, b) (Heller et al., 2011). Once Cdc6 and Cdt1 are released, they result in the closing of Mcm2-7 ring around DNA where two MCM2-7 rings are needed at each origin (Ticau et al., 2017). In addition, this activation enables recruitment and formation of the CMG complex consisting of cell division control protein 45 (CDC45), MCM and GINS (synthetic lethality with Dpb11-5 (Sld5, Psf1, Psf2 and Psf3). Dpb11 is TOPBP1 in higher eukaryotes. The association of these three components enhances the ATPase activity of MCM2-7 and thus its helicase activity (Ilves et al., 2010). However, in order for the CMG to be loaded to origins, a set of loading factors has to be activated through phosphorylation by CDK, including Sld2, Sld3 and Dpb11 in yeast orthologs of TOPBP1-interacting replication stimulating protein, treslin, and TOPBP1 in mammals respectively. Phosphorylation of treslin by CDK allows its interaction with TOPBP1. The binding of treslin to TOPBP1 allows the functional integration of other replication initiating proteins (Boos et al., 2011). GINS is also recruited by RecQ helicase allowing GINS interaction with MCM2-7 in S phase (Xu et al., 2009). Moreover, CDC45, a component of the CMG complex was shown to be loaded to replication origins through treslin. Depletion of treslin in the nucleus inhibits CDC45 loading, thus altering S-phase progression in human cells

After the activation of replication origins (Figure 3, c), polymerases (pol) are recruited to the origins selected to initiate DNA synthesis. The primase subunit of pol α initiates RNA-DNA short primers synthesis on both leading and lagging strands. The number of RNA-DNA hybrids on the leading strand is much less than RNA-DNA hybrids on the lagging. This comes from the fact that replication at the leading strand is continuous whereas it is discontinuous on the lagging.

Thus polymerase α synthesizes a primer for each Okazaki fragment and have little contribution in the elongation process (Ottiger and Hübscher, 1984). DNA products in pol ƍ and pol ɛ depleted Xenopus extracts were analyzed. The results showed that pol ɛ-depleted gene products are longer than pol ƍ-depleted products suggesting the contribution of pol ɛ in the replication of leading strand while pol ƍ is involved in the extension of Okazaki fragments. However, pol ƍ depletion induced an increase in chromatin-bound replication factor C (RFC), proliferating cell nuclear antigen (PCNA), replication protein A (RPA) and flap endonuclease 1 (FEN1) (Fukui et al., 2004). RFC binds to the 3'-OH of the primer in an ATP-dependent manner, then PCNA binds to RFC to form an unstable complex. After hydrolysis of ATP, RFC loads the PCNA clamp on chromatin then RFC gets released. Thus, this complex is loaded through nicks on double-stranded DNA to interact with pol ƍ and pol ɛ and replicate single-stranded DNA (Podust et al., 1995). In addition, another study has shown that RPA controls pol α loading on primed sites by targeting RFC-PCNA complexes to the primed ends. During primer extension, RPA binds to pol α, allowing its firm attachment to the site. After that pol α dissociates. This dissociation at the primed sites allows RFC to load PCNA to the site of DNA synthesis. Pol ƍ then associates with PCNA, allowing stable interaction of pol ƍ to the DNA and further extension of primers (Yuzhakov et al., 1999).

Inactivation of Licensing Factors during S Phase

As mentioned previously, the pre-RC complex is formed in G1 and activated in S phase by DDK and CDK. Upon S-phase entry, licensing factors including CDC6 and CDT1 and other pre-RC complex components were shown to be inactivated by several mechanisms in order to ensure that each origin of replication will be activated only once per cell cycle. These mechanisms include CDT1 inhibition by geminin, phosphorylation of pre-RC components by

CDKs and ubiquitin-proteasome-targeted degradation.

Wohlschlegel and his colleagues have shown that recombinant human geminin disrupts pre-RC assembly in Xenopus nuclei by the inhibition of Xenopus MCM4 loading in a CDT1dependent manner. Furthermore, HeLa cells were shown to express CDT1 during G1 and S phases but not in G2, while geminin was shown to be expressed in late S and G2. This suggests a role for geminin in inhibiting CDT1 and consequently re-replication. The role of geminin in inhibiting CDT1 was further demonstrated by Tada and colleagues. Addition of recombinant geminin to Xenopus nuclei led to inhibition of replication licensing which was rescued by the addition of recombinant CDT1 (Tada et al., 2001;Wohlschlegel et al., 2000). CDC6 was also shown to be regulated in a cell-cycle-dependent manner where cyclin A-CDK2 induces its phosphorylation and consequent translocation from the nucleus to the cytoplasm during S phase (Petersen et al., 1999). Similarly, CDT1 was also shown to be inactivated by cyclin A/CDK1and cyclin A/CDK2-dependent phosphorylation without affecting geminin binding, suggesting that CDT1 is regulated by cyclin A/CDK1-, CDK2-dependent phosphorylation independently of geminin-targeted regulation (Sugimoto et al., 2004).

Selection of Origins of Replication for Activation

Regulation of DNA Replication in Space

The very first evidence concerning origin distribution in mammalian cells was described by Huberman and Riggs in 1968. Their results show that chromosomal DNA is made up of tandemly-joined replication sections. DNA synthesis occurs un-equidistantly and at different times on these different sections (Huberman and Riggs, 1968). Potential replication origins are arranged in a replicon or replication unit where only a single origin is activated to replicate the region. Each replication unit has different potential replication origins with different affinities.

The activation of origins within the replication unit is stochastic and flexible. It differs from one cell to another even within the same cell type depending on chromatin folding (Fragkos et al., 2015a). Since a replication unit is defined as a region of DNA replicated by a single origin, not all of the potential origins present within the replication unit can be used. The rest of the potential pre-RCs are removed passively by upcoming nearby forks of more efficient origins (Lebofsky et al., 2006a). Origin efficiency is defined as the frequency of a certain origin getting activated in a population of cells.

Replication origins are further arranged in clusters and origins within the same cluster are activated synchronously through positive interference mechanism (Marheineke and Hyrien, 2004). Origins fired at the same time within a cluster can be visualized using single-molecule analysis by DNA combing. Organization of replication units in clusters forms replication domains (RD). Cohesin-depleted HeLa cells showed decrease in the density of active origins, associated with a decrease in the intensity of replication foci. This suggests that cohesins might involve chromatin loop formation where several replicons are tethered together by cohesins to form a single domain (Figure 4) (Guillou et al., 2010). However, it has been long difficult to study these structures at the molecular level due to limitations of methodologies used. DNA synthesis occurs in subnuclear structures called replication foci. The pattern of these foci changes dynamically during the S phase (see section2.2.2). However, this pattern is conserved throughout several rounds of replication within the same cell type. This suggested that replication foci are a fundamental structure of mammalian chromosomes (Ma et al., 1998a;Nakamura et al., 1986a).

Regulation of DNA Replication in Time

Besides spatial organization, replication origins are further controlled in time depending on several factors including chromosome organization in the nuclear space, transcriptional activity and concentration of replication factors.

In the Ma and colleagues study mentioned above, synchronized 3T3 mouse fibroblasts showed changes in replication foci patterns throughout S phase. 3T3 fibroblasts were synchronized and labeled with 5-chloro-2'-deoxyuridine (CldU) for 2 minutes detected in green in early S phase followed by 4 hours, 6 hours and 10 hours chase. Cells were then pulsed with 5-Iodo-2 -deoxyuridine (IdU). After 4 hours, 6 hours and 10 hours chase cells were in mid-S phase, late-S phase and G2 as shown in (Figure 5). The green foci correspond to foci replicating in early S phase. While the red foci correspond to foci replicated later in mid and late-S phase, and they disappear after 10 hours of chase since the cells entered G2. The early-replicating foci are small and more numerous compared to late replicating foci which are larger and less numerous (Ma et al., 1998a). Genome-wide analysis of replication timing has been devised by Azuara (Azuara, 2006).

It is based on sorting S phase cells according to bromodeoxyuridine (BrdU) incorporation profile into early and late S phase. This is followed by, polymerase chain reaction, microarrays or nextgeneration sequencing analysis to map the replicated segments. Then the log2 of the ratio of early/late signal of each probe is determined. If the log2 ratio is higher than 0 then the region is early replicating whereas a log2 ratio lower than 0 suggests that the region is late replicating (Ryba et al., 2011). Most of the mapped chromosomes segments are either early or late RD, and they are referred to as constant timing replication regions (CTRs). These RDs replicating at different times are separated by timing transition regions (TTRs) in which DNA replication progressively advances (Figure 6, a).

Early replicating domains are origin-rich regions that have the features of euchromatin.

They are gene-rich, transcriptionally active and located mainly in the interior of the nucleus (Dellino et al., 2013). On the other hand, late-replicating domains are gene-poor regions, not transcriptionally active and resemble the features of heterochromatin. These regions are usually associated with nuclear lamina and referred to as lamina-associated domains (LAD) (Figure 6 (Pope et al., 2014a;Xiang et al., 2018).

Origin mapping using short-nascent strand in HeLa and mouse embryonic stem cells (ESCs) revealed a correlation between origin density and enrichment of genes. The results

showed that 44 regions covering 30 Mb have regions of 500 Kb devoid of strong initiation events. These regions were shown to replicate in mid to late-S phase indicating that these regions are either TTR or late-replicating regions (Cadoret and Prioleau, 2010). The replication timing program is conserved among related eukaryotic species. The main biological function of it is to ensure full replication of the genome through temporal activation of replication forks in order not to exhaust limiting factors. Thus, the initiation of replication in eukaryotes is extended throughout the S phase. Besides ensuring the availability of replication factors, the reason behind having evolutionary conserved temporal order might be attributed to regulating gene dosage of histone genes that are highly needed during S phase (Müller and Nieduszynski, 2017). Furthermore, the replication timing program allows concentrating genomic polymorphism in particular sites of the genome. Given that mutations frequencies vary throughout S phase, it has been shown that the frequency of point mutations increases in latereplicating regions due to a decrease in the efficiency of DNA mismatch repair. As most of essential genes are replicated early, this temporal regulation prevents the accumulation of mutations in essential genes preserving their function (Supek and Lehner, 2015).

In addition, this temporal regulation of replication was shown to be correlated with cell fate transition. The time by which RDs are replicating and their position in the nucleus are developmentally regulated. Human embryonic stem cells show weak correlation between replication timing and nuclear compartmentalization; however, this correlation becomes more evident as cells differentiate. Replication domains get closer leading to compartment consolidation (Dileep et al., 2019).

Transcription Favors Replication

It has been shown a long time ago that transcription and replication origin activation are correlated. Goldman and his colleagues have shown that transcriptionally active genes, cell type and tissue dependent, replicate in early S phase whereas inactive genes replicate later (Goldman et al., 1984).. Origins of replication tend to be enriched in CpG islands (CGI) devoid of methylation near transcription binding sites (Delgado et al., 1998). This suggests that decondensed chromatin allows the access of replication machinery; however, this depends on the transcriptional activity of these regions. Using short RNA-primed nascent DNA sequencing, MYC, HBB and HPRT1 genes were shown to be enriched in replication-initiation events near genes promoters. The technique is based on the fact that newly replicating RNA-primed DNA strands are synthesized symmetrically and bi-directionally from the replication origins.

Treatment of DNA with lamda exonuclease allows the purification of RNA-primed DNA. This is followed by purification of nascent strands based on their size. Short-nascent strands indicate newly synthesized DNA which are sequenced later. This allows the mapping of origins genomewide. Then the frequency of replication initiation was determined in several genes with different expression levels in MCF-7 cell line. The results showed that replication-initiation events are less frequent in genes with very low or very high transcriptional activity. In addition, the absence of replication-initiation events from the TSS was directly related to transcription activity. This suggests that the chromatin conformation of transcribed genes helps pre-RC loading on the chromatin. However, the transcriptional activity itself may interfere with replication events (Martin et al., 2011).

The problem with short RNA-primed nascent DNA sequencing is it can be biased due the fact that the replicative polymerases may pause when they encounter G-quadruplexes. Thus, the purified short nascent strands might be contaminated with these short strands. However, it has been shown that using the experimental conditions in Prioleau's lab, the possibility of contaminating short RNA-primed nascent DNA sequencing with short sequences encountered Gquadruplexes has been excluded (Valton et al., 2014). Mentioning that G-quadruplexes can impede fork progression in this case, here we come to the conflict in research whether Gquadruplexes act as mediator or an impeder. The mechanism by which G-quadruplexes can be considered as a problem will be further discussed in section 3.1.3.2.

The expression of common fragile sites (CFS) are defined as breaks, gaps or decondensations seen on preferential locations in the genome when cells are exposed to replication stress. They are late replicating regions that harbor very long genes. The transcription of these long genes would start in G2/M phase of the previous cell cycle and lasts till the end of S phase till the next cell cycle. As a result, transcription-replication collisions occur leading to fork collapse and strand breaks. However, inducing the transcription of some CFS in chicken DT40

and human HCT116 cells led to a redistribution of the initiation events in that region with enrichment at the promoter accompanied with advancing the replication timing. This advancement of replication timing in S phase rescued the fragility of the CFS studied. All these suggest that transcription affects the distribution of initiation events and might affect the replication timing (Blin et al., 2019). Furthermore, deletion of early replication control elements (ERCEs), involved in nuclear compartmentalization, led to alteration in transcription and replication timing for some genes, however in other genes it altered transcriptional activity but not replication timing (Sima et al., 2019) suggesting that transcription does not directly alter replication timing.

Chromatin Modifications Affecting Origins Activity

Role of Histone Modifications in Origins Localization and Initiation

In mammalian cells, the exact sequence features involved in determining the initiation frequency of origins is to be elucidated. They do not exhibit any consensus sequences, but they share common features including CGI, G-quadruplexes and transcription start sites in addition to others (Besnard et al., 2012a;Foulk et al., 2015). However, whether G-quadruplexes act as a necessity or a problem is still controversial and will be further discussed in section 3. 1.3.2. This observation suggests a possible role of the replication machinery in modulating chromatin structure to make it more permissive for transcription and replication. However, binding of the pre-replication complex to replication origins is essential for replication initiation, but it cannot be the only reason explaining consistent replication patterns seen in higher eukaryotic cells. Histone modifiers affect origin activity locally or globally. For example, the methylation of histone H4 on lysine 20 by SET8 globally enhances the loading of the prereplication complex through the interaction with ORC1 facilitating licensing (Tardat et al., 2010). Furthermore, chromatin modifiers can also act locally to control origin activity.

Euchromatin histone modifications such as some histone H3 methylation (H3K4me, H3K4me2

and H3K4me3) and acetylation (H3K9ac, H3K18ac and H3K29ac) are prominent in earlyreplicating regions whereas other forms of modifications are associated with late-replicating regions such as H3 and H4 hypoacetylation, H3K9me, H3K9me3 and H3K27me3 (Picard et al., 2014). The association of chromatin modifiers with certain replication origins is correlated with cellular differentiation status. Constitutive origins are usually associated with euchromatin markers mentioned above and coupled with unmethylated CGIs, whereas cell-type-specific replication origins are associated with heterochromatin markers (Smith et al., 2016).

It has been suggested that transcription alone is not sufficient to alter replication timing and it is probably the chromosomal organization that impact the regulation of replication timing.

A mid-late replicating region devoid of origins in chromosome 1 was selected in chicken DT-40 cell line. In one of the alleles on this site, transcription was activated through the insertion of strong chicken β-actin promoter and had faint effect on advancing replication timing. In addition, this site was shown to be enriched with H3 acetylation. This indicated that transcription and histone modifications alone are not sufficient in inducing a change in replication timing.

However, when β A -globin origin was inserted, a strong shift in replication timing was induced.

Furthermore, introducing the insulator element HS4 next to β A replicator induced further shift in replication timing. This shift was dependent on the USF binding motifs of the HS4 insulator (Hassan-Zadeh et al., 2012).

Finally, the relationship between replication timing and transcriptional activity is similar to the chicken and egg conundrum. The above-mentioned studies in sections 2.2.3. and 2.3.1. involve correlation, however the mechanistic links are still unknown. In the Blin study, enhancing transcriptional activity of long genes harbored by CFS induced a shift in replication timing. However, it is worth mentioning that the shift in replication timing is seen in long genes that might extend over 1 replication domain (Blin et al., 2019). On the other hand, the Sima study has shown that alteration in transcriptional activity does not always change replication timing (Sima et al., 2019). Moreover, the mentioned studies in this section showed that histone modifications allow the access of the replication machinery and enhance replication. All these confirm that the relation between replication timing and transcription is not really direct.

Trans Acting Elements Affecting Origins Activity

Several proteins that are not members of the pre-replication complex have been identified

through interactions with DNA at replication-initiation sites and have an impact on replication timing. Replication-initiation determinant (RepID) is a member of DDB1-and CUL4-associated factors (DCAF) family of proteins that might act as an adaptor for specific sequences facilitating early replication in human beta-globin locus in erythroid cells (Zhang et al., 2016).

Another protein that has been shown to regulate origin firing is Rap1-interacting factor (RIF1). Mammalian RIF1 has been shown to be involved in regulating the timing of replication foci and domains. Loss of RIF1 enhanced replication initiation events in early S phase associated with changes in the pattern of early, mid and late replication foci. Mid-replicating foci were lost upon RIF1 depletion with more prevalence of early replication foci. In addition, RIF1 loss induced changes in the replication timing of some replication domains but not all of them.

Immunostaining of RIF1 revealed overlap between RIF1 signal and Lamin B1 suggesting its role in generating chromatin loops. It associates with the nuclear lamina and binds to constitutive heterochromatin. This association possibly regulates origin affinity for limiting replication factors (Yamazaki et al., 2012a).

Cohesins have been shown to be enriched at replication origins as well. They are thought to bring neighboring origins in proximity within the replication factories allowing them to fire simultaneously. This deposition concentrates initiator proteins and kinases allowing origins activation. Down-regulation of cohesins resulted in fewer origins activated during S phase (Guillou et al., 2010).

Besides its role in translesion synthesis, pol Ѳ was detected in cells exiting mitosis and recruited to the chromatin in early G1 in absence of exogenous replication stress.

Downregulation of pol Ѳ resulted in an increase in MCM loading to the chromatin. This increase in MCM loading was not due to licensing of more pre-RCs. Instead Pol Ѳ was shown to have a similar but independent role as RIF1 in regulation of the replication timing of replication domains. Depletion of pol Ѳ in rectal cancer cell line RKO lead to increased loading of MCM during G1 without affecting the density of active origins during S phase. However, genome-wide analysis of replication timing in pol Ѳ-depleted RKO cells revealed a change in replication in some domains indicating that pol Ѳ have a role in TDP. Overexpression of pol Ѳ delayed replication timing in subset of domains confirming the role of pol Ѳ in regulating replication timing (Fernandez-Vidal et al., 2014).

pSIRT1 is a NAD-dependent protein deacetylase sirtuin1. When phosphorylated on Thr530 (T530-pSIRT1), it associates with replication origins that are activated during middle and late S phase. This association was prominent in gap phases of the cell cycle and colocalized with origins of replication but not on elongating forks. Depletion of SIRT1 in mammalian cells causes them to activate additional origins and induces DNA breaks emphasizing the role of SIRT1 in preventing dormant origins activation. This prevents origins overactivation and genome instability (Utani et al., 2017).

Regulation of DNA Replication in Embryonic Stem Cells

The study of stem cells has started in 1961 where Till and McCulloch had accidently observed the presence of proliferating cells in the spleen of irradiated mice that have been injected with bone marrow cells. This opened the gate later for studying and isolating different types of stem cells based on their importance in a wide range of clinical uses (Till and McCULLOCH, 1961). These bone marrow stem cells are tissue-specific stem cells. They are able to generate several types of cells related only to the bone marrow. Some other tissues such as the gut and organs have their own tissue-specific stem cells responsible for replacing turnedover cells. These tissue-specific stem cells are referred as adult stem cells. ESCs are another type of stem cells obtained only during embryonic development. They are pluripotent stem cells that are able to give rise to specialized cells of the whole body. Pluripotent stem cells embryonic cells that give rise to the ectoderm, endoderm and mesoderm. These germ layers give rise later to the several types of cellular tissues. In our study, we will focus on the properties of mouse ESCs.

Characteristics of Embryonic Stem Cells

Three days after fertilization, the blastocyst is formed before implantation. it consists of the outer trophectoderm (TE) layer and the inner cell mass (ICM). The TE layer gives rise to the extra-embryonic tissue whereas the ICM is the source of the pluripotent embryonic stem cells (Figure 7.). Distinct pluripotent states exist in vitro depending on the stage of isolation of the ESCs.

Mouse ESCs isolated from pre-implantation epiblast embryos are considered to be naïve whereas those isolated from post-implantation are considered to be primed. Primed mouse ESCs or epiblast stem cells are referred as EpiSCs.

Pluripotency

Mouse ESCs were the first pluripotent stem cells to be successfully cultured (Martin, 1981). They were isolated from the epiblast and maintained in vitro in the presence of leukemiainhibitory factor (LIF), a cytokine secreted by trophoblasts and important for implantation and embryogenesis, permitting their homogenous growth by suppressing their spontaneous differentiation (Smith et al., 1988). In 2000, Reubinoff and colleagues isolated human ESCs from human epiblast (Reubinoff et al., 2000). Chimera assay has been used to assess the developmental potency of human stem cells in mouse host embryos, based on the ability of stem cells to generate an organism (Mascetti and Pedersen, 2016). At the molecular level, ESCs can be distinguished based on the expression of certain transcription factors such as Oct4, Sox2 and Nanog which maintain their pluripotent state (Kim et al., 2008). While metabolites play a role in inducing epigenetic changes, metabolite profile switching during different stages of pluripotency identifies cellular fate during developmental stages. Thus, pluripotent stem cells have their unique metabolic signatures. For example, EpiSCs are characterized by having high levels of H3K27me3 on genes involved in development, leading to their repression (Sperber et al., 2015). telomeres showed an increase in copy variant number compared to the control. This suggests that maintaining telomerase activity is essential for maintaining self-renewal in human ESCs (Zeng et al., 2014).

Tumorigenesis

The pluripotency of stem cells has been confirmed by characterizing their ability to form teratoma in immunodeficient mice in vivo. However, besides their clinical potential in therapy, transplanting human patients with them is facing a lot of hurdles because of their tumorigenic capacity. It has been shown that stem cells and tumor cells share common molecular signatures.

Bioinformatic analysis has revealed the oncogenic properties of stem cells when compared to primary cells (Ghosh et al., 2011). heterogeneity in molecular and cellular properties depending on their spatial attribution (Wu et al., 2015). ESCs have the ability to proliferate and differentiate into adult cell lineage which is the principle of cell-replacement therapy using ESCs.

Cell Cycle Control of Embryonic Stem Cells Compared to Somatic Cells

The cell cycle profile of ESCs has been previously characterized. While proliferating somatic cells require around 24 hours to cycle, ESCs need only 16 hours. How the shortening in the cell cycle affects the length of each phase has been questioned as well. G1 phase was also shown to be shortened which might be the reason behind this fast proliferation (Becker et al., 2006). In 1998, Aladjem and colleagues have attributed this shortening of G1 to an absence of cell cycle checkpoint control suggesting that cell cycle regulation and checkpoint controls are acquired during differentiation, and DNA-damage pathways can be tissue-specific (Aladjem et al., 1998). The question whether this shortening of G1 and cell cycle length have a role in maintaining pluripotency has been addressed in 2013. LIF signaling is important in regulating ESC cycle and contributes in G1/S regulation. LIF withdrawal from the media of in vitro cultured ESCs led to alteration in G1 kinetics, inducing G1 prolongation accompanied with differentiation. Similar results were seen by knocking down cyclin E, leading to delayed S-phase entry and facilitating differentiation supporting the idea that short G1 maintains pluripotency (Coronado et al., 2013a). It is though worth to mention that ESCs are susceptible to differentiation when they encounter any type of stress.

Replication Dynamics of Embryonic Stem Cells Compared to Somatic Cells

Since G1 is the point where maximum loading of MCM occurs and stem cells spend a short time in G1, how these cells manage to replicate their genome with enough origins has been addressed. The replication stress in S phase is influenced by the amount of loaded MCM. Despite having short G1, ESCs load MCM on chromatin at a faster rate compared to differentiated cells.

This fast loading of MCM complexes in ESCs is attributed to increased levels of chromatinbound CDC6 and CDT1 in human ESCs during G1 (Ballabeni et al., 2011a;Fujii-Yamamoto et al., 2005a;Matson et al., 2017) and contributes to efficient licensing. Besides the presence of excessive licensing factors and fast loading on chromatin, the chromatin of ESCs is more homogenous. The ratio of heterochromatin to euchromatin is higher in differentiated cells compared to ESCs. This state of euchromatin in ESCs also facilitates the accessibility of chromatin to licensing factors which also may contribute to efficient licensing.

Drivers of Replication Defects and Genome Instability

DNA Replication Defects Induce DNA Damage

While DNA replication is progressing, the opened DNA gets exposed to damage impeding the replication forks from going on by inducing either their stalling or collapse. This is defined as replication stress which alters efficient DNA replication inducing genome instability, a hallmark of cancer. Fork stalling occurs when replication is stopped without altering the integrity of the replisome on DNA: it continues when the problem is resolved. In contrast, fork collapse involves disruption of replisome integrity on DNA. Though there is no single definition that describes replication stress. It does not necessarily include replication defects originating from alteration of origin number and re-replication (Zeman and Cimprich, 2014). However, defects in origin licensing might induce fork stalling or collapse.

Exogenous DNA Damage and the Repair Pathways Commonly Activated

Several exogenous factors have been reviewed to interfere with DNA integrity and induce DNA damage by causing base modifications, cross-linking and other DNA lesions leading to single strand breaks (Tsegay et al., 2019). Depending on the type of damage, DNA is either repaired by base excision repair or nucleotide excision repair. Base excision repair involves DNA glycosylases which remove the damaged base by flipping it out. The gap is then filled using polymerase β and ligated. Deamination of cytosines leads to the incorporation of uracil, and it is one of the DNA damages repaired by base excision repair (Courcelle et al., 1999). Nucleotide excision repair involves repairing damaged bases with their surroundings because it usually recognizes damages that distort DNA helix. DNA damaged caused by UV radiation or cigarette smoking. Failure to repair these regions might result in mismatches and unresolving of stalled forks. In addition to single strand breaks, double strand breaks (DSB) can also be generated. These breaks can be repaired by several pathways including homologous recombination (HR) and non-homologous-end joining (NHEJ). As the name implies, HR utilize a homologous template (the sister chromatid, when available) to repair the gap, resulting in less error-prone repair repair. NHEJ on the contrary, does not use homologous template resulting in nucleotide deletion and / or insertion, thus it is considered to be an error-prone pathway. These pathways in addition to translesion synthesis are considered DNA damage tolerance pathways that are used to avoid DNA breakages. Because not all of the DNA lesions created can be repaired by template switching, they require the involvement of less accurate and efficient DNA polymerases to repair the DNA stretches. These are called translesional DNA polymerases that lack the proof-reading activity. These polymerases are divided into several subfamilies however, the term usually refers to Y-family polymerases (REV1, pol eta, pol iota, and pol kabba) and Bfamily (pol zeta). The mechanism involves switching of replicative polymerase on stalled forks with translesional polymerases followed by nucleotides extension (Livneh et al., 2010;Ma et al., 2020).

Alteration of Nucleotides Pools

Another factor that contributes to replication stress since early stages of carcinogenesis is alteration of the nucleotide pool. It has been previously shown that infection of primary keratinocytes with E6 and E7, human papilloma virus onco-proteins, induces a decrease in fork progression rate and an increased fork asymmetry between forks originating from the same origin, in addition to activation of dormant origins. This was accompanied by a reduction in nucleotide pool biogenesis. Exogenous supply of nucleosides to E6/E7-induced keratinocytes rescued the phenotype, indicating that the replication stress was induced because of shortage in the nucleotides pool. The study proposed that insufficient nucleotides were generated because of precocious S-phase entry which might affect the transcription of genes involved in nucleotide biogenesis (Bester et al., 2011). This concept was further studied by Beck and colleagues. The inhibition of WEE1 mitotic kinase, known to be a CDK inhibitor, induced deregulated CDK levels. This inhibition of WEE1 led to activation of more origins. As a result, more nucleotides were consumed and became limiting which in turn led to reduction in the apparent fork speed and induction of DSB due to fork collapse (Beck et al., 2012). Similar results can be obtained upon treatment of cells with hydroxyurea. Hydroxyurea is a well-established ribonucleotide reductase inhibitor that has been used in scientific research more than 60 years ago. The inhibition of ribonucleotide reductase alters nucleotides pool biogenesis which depending on the does induces forks arrest and DNA damage (Singh and Xu, 2016).

DNA Secondary Structures

Replication-Transcription Interference and R-Loop Formation

Origins of replication were shown to partly overlap with transcriptional promoters (Martin et al., 2011). Given the fact that chromatin is more open during transcription this might allow easier loading of components of replication origins. In order for replication and transcription to occur, both mechanisms use the same DNA templates leading to the evolvement of mechanisms to limit replication-transcription conflicts in prokaryotes and eukaryotes. Studies on mapping human genome suggested that replication and transcription do not occur codirectionally (Necsulea et al., 2009). Sasaki and colleagues had shown that transcription negatively regulates the activation of origins within the dihydrofolate reductase locus in Chinese hamster cells. Active transcription affects the selection of active origins within the region where the presence of transcriptional machinery allows the access of replication factors. In addition, active origins were mainly localized to the intergenic sequences. Inhibition of the dihydrofolate reductase promoter resulted in intragenic initiation of replication (Sasaki et al., 2006).

The replication and transcription are coordinated processes. While active transcription occurs mainly in G1, some long genes would still be transcribed during S phase. This requires coordination between replication timing and transcription. The genomic regions harboring long genes, such as CFS, are more prone to replication-transcription collisions. These regions are hotspots of genomic rearrangements and instability. Since the transcription of long genes is throughout the cell cycle, this might lead to the formation of stable RNA:DNA hybrids or so called R-loops. FHIT, WWOX and IMMP2L are very long genes harbored by FRA3B 3p14.2, FRA16D 16q23 and FRA7K 7q31 respectively. CFS break formation was compared in different mammalian cell lines with different gene expression level suggesting that transcription induces fragility at CFS. CFS breaks were partly rescued by transfecting cells with RNase H1 which reduced RNA:DNA hybrids or so called R-loops (Helmrich et al., 2011). The mechanisms of replication occurring in silent versus transcriptionally active genes are illustrated in (Figure 8).

G-Quadruplexes

Besides R-loop formation, the DNA strand by itself can form structures that alter genome instability. Such structures are referred as secondary structures and their form is determined by the DNA sequence, including G-quadruplexes formed by repetitive G-rich sequences. The stability of these structures might be even stronger than the double-stranded B-DNA itself. The ability of forming G-quadruplexes comes from guanines rolling within single-stranded DNA forming a uni-molecular stable structure (Gellert et al., 1962). In human B-lymphocytes, around 700,000 potential G-quadruplexes-forming sites whose height and loop size vary depending on the sequence have been identified (Chambers et al., 2015). Similar to the R-loops, Gquadruplexes are thought to act as barriers that impede replication forks from progression. DNA double strand breakpoints inducing translocations and deletions were shown to be enriched with DNA secondary structures, including G-quadruplexes, suggesting a possible role of Gquadruplexes in inducing genome breakpoints and instability (Bacolla et al., 2016).

Given the fact that the lagging strand remains single-stranded more than the leading strand, then it might be more prone to G-quadruplexes formation. However, the competition of RPA to bind exposed single-stranded DNA prevents the formation of G-quadruplexes (Safa et al., 2016). The leading strand is also prone to have G-quadruplexes in the absence of helicases necessary to unwind the DNA (Sarkies et al., 2012). In case of polymerase arrest or slowing down, the helicase continues to unwind the DNA while the replicative machinery remains stalled or slowed down, which increases the distance between helicase and polymerase. This leads to more exposure of single-stranded DNA, which may allow G-quadruplexes formation. The presence of G-quadruplexes motif in the leading strand heading to the TSS exacerbate the genome instability induced by hydroxyurea of the BU-1 locus in DT-40 chicken cells. This emphasizes on the contribution of G-quadruplexes in inducing genome instability (Papadopoulou et al., 2015).

In order to overcome G-quadruplexes formation, in addition to RPA binding, several helicases are utilized including bloom (BLM), werner (WRN) and fanconi J (FANCJ). BLM helicase was shown to resolve G-quadruplexes through interaction with 3'-single-stranded overhang (Sun et al., 1998). G-quadruplexes were shown to be strongly enriched close to the promoters in down-regulated genes in BLM-depleted human fibroblasts, suggesting the role of G-quadruplexes in altering gene expression as well (Nguyen et al., 2014). Another helicase which was shown to be involved in resolving them is WRN helicase found to be mutated in werner syndrome. It efficiently unwinds the G-quadruplexes (Kamath-Loeb et al., 2001). FANCJ is another helicase mutated in fanconi anemia. Similar to the above helicases, it helps resolving G-quadruplexes to maintain chromatin epigenetic status (Sarkies et al., 2012).

Oncogene-Induced DNA Damage

The term oncogene-induced replication stress is referred to cells acquiring sustained proliferation through the activation of oncogenes. The alteration in both types of genes alters the regulation of cellular proliferation. Cyclin E is frequently amplified in many human cancers leading to deregulation in the cell cycle and gene expression. In addition, it also induces replication stress mainly through affecting origin licensing and firing. The mechanism by which origin licensing and firing are affected and induce replication stress will be addressed in section 3.1.5.

The P53 tumor suppressor protein is involved in inducing apoptosis following DNA damage (Schumacher et al., 2001). Once DNA double strand breaks are formed, they are detected by the checkpoint kinase ATM protein which in turn phosphorylates P53 leading to its release from the P53 inhibitory protein MDM2. Once released, P53 in turn activates genes involved in apoptosis cell cycle arrest and DNA repair (Shiloh, 2003).

Deregulation of Replication Licensing

Recent studies have started shedding light on the implication of altered origin licensing on replication and how it is orchestrated with oncogene-induced replication stress. As mentioned in the first chapter, replication licensing is a highly-regulated event where any alteration, including relicensing, over-licensing and under-licensing, drives genome instability. The importance of origin licensing is a two-edged sword. If licensing occurs even during the S phase, the same DNA template will be replicated more than once driving DNA re-replication. However, if no enough origins were licensed during G1, the cells will enter S phase with few numbers of origins. As a result, the cells will not be able to rescue DNA replication by firing additional origins if replication stress was encountered.

Origin Relicensing

The reason behind the regulation of DNA replication in two exclusive steps is to ensure that origins are activated only once per cell cycle. Thus, origin licensing is restricted to G1

phase. If licensing occurs outside G1, the same DNA region might be replicated several times during the same cell cycle which leads to re-replication, replication stress and gene amplification (Alexander and Orr-Weaver, 2016). Origin relicensing can occur by overexpressing licensing proteins including CDT1 and CDC6, inducing re-replication and checkpoint activation (Vaziri et al., 2003). Cdt1 overexpression induces re-replication and tissue dysplasia in vivo (Muñoz et al., 2017). The reason why re-replication induces DSB comes from increased fork collisions.

Deregulation of origin licensing was shown to induce origin refiring, accumulation of RPAcoated single-stranded DNA, and increased fork reversal, suggesting that the DNA breaks generated are the result of fork collisions due to re-replication (Neelsen et al., 2013). In vivo in mouse intestinal cells, the deregulation of Cdc6 and Cdt1 was shown to increase the expression of p21 using immunohistological analysis , suggesting a DNA-damage response (DDR) mediated by the p53 pathway accompanied by increased apoptosis (Muñoz et al., 2017). However, since many cancers have an altered p53 pathway, then this barrier is lost. In this case origin relicensing acts as a fuel to induce genome instability.

Ectopic Licensing and Over-licensing of Origins

A connection between oncogene-induced replication stress, ectopic licensing and genome instability has been addressed by Macheret and colleagues. Replication origins were mapped in U2OS cells overexpressing either cyclin E or MYC. Both cyclin E-and MYC-overexpressing cells fire a new subset of origins that are absent in control cells. These new origins are located in the intragenic regions which would create replication-transcription collisions and fork collapse.

Cyclin E overexpression was also associated with genomic rearrangements Intragenic pre-RCs are usually removed by transcription before the cells enter S phase. However, premature S-phase entry prevents the removal of the pre-RCs. As a result, these origins fire while transcription is taking place which induces fork collapse (Macheret and Halazonetis, 2018).

Shoaib and colleagues also showed that maintaining a threshold of chromatin compaction during G1 regulates origin licensing and preserves genome integrity. SET8 mediates H4K20 methylation after mitosis. Depletion of SET8 from synchronized U2OS led to chromatin relaxation and increased binding of ORC and MCM to DNA. This was also associated with an increase in the single-stranded DNA in S phase. This suggests that SET8 maintains fine-tuned chromatin compaction in G1 which prevents over-licensing thus DNA damage in S phase (Shoaib et al., 2018). Taken all together, these data emphasize on the importance of G1 duration and chromatin compaction state on maintaining origin licensing and genome stability.

Origin Under-licensing

Eukaryotic cells license more origins than needed in an unperturbed S phase. Most of the licensed, unfired origins usually remain dormant and fire only when S phase is challenged. They serve as a backup to rescue replication when forks are slowed down or arrested. As the cells cannot license more origins when they are in S phase, it is essential to license an excessive number of origins during G1 (Alver et al., 2017). Thus, the reduction in the number of licensed origins will result in DNA regions with a reduced number of backup origins. If replication stress occurs in these regions, cells will struggle to finish replication. This causes these regions to be under-replicated and induces genome instability. This was supported by the work of Ge and colleagues. Using DNA fiber analysis, they showed that treating U2OS with replication inhibitors such as aphidicolin and hydroxyurea led to a decrease in the space between active origins. This implicated that additional dormant origins had been activated to rescue replication stress. However, knocking down MCM5 led to a decrease in the total loading of MCM. Thus, fewer origins were available to replicate the genome. si-MCM5 treatment of U2OS did not affect origins seen by DNA fiber analysis. However, when MCM5 knockdown cells are challenged with hydroxyurea, they fail to activate dormant origins. This suggested that knocking-down MCM5 led to a decrease in the available dormant origins. In addition, this was associated with increased levels of DNA damage (Ge et al., 2007).

Whether oncogene-induced replication stress can induce origin under-licensing has been addressed by Ekholm-Reed and colleagues. Overexpression of cyclin E in KB cells induced early S-phase entry. This was accompanied with reduced BrdU incorporation and PCNA foci numbers in S phase. Furthermore, MCM4 and MCM7 loading was highly reduced in cells released from mitosis indicating that impairment of replication was due to alteration of licensing (Ekholm-Reed et al., 2004a). Live-imaging of MCM dynamics during the cell cycle showed that MCM loading starts at the end of mitosis and peaks at the end of the G1 phase (Symeonidou et al., 2013).

Therefore, premature S-phase entry escapes the last wave of MCM loading, leading to S-phase entry with a lesser number of origins. Furthermore, seminal yeast work from our lab has shown that G1 deregulation prevents full replication origin licensing achieved by the loading of pre-RC on a large number of chromosomal sites in M/G1. As a consequence of G1 deregulation, cells proceed through S phase with fewer active origins, delaying the completion of chromosome replication until after the initiation of mitosis. This leads to chromosome breaks, rearrangements and mis-segregation (Lengronne and Schwob, 2002a;Tanaka and Diffley, 2002a). Using singlemolecule analysis of DNA replication, our lab showed that mouse fibroblasts lacking all three pRb paralogs that control G1/S progression have a lower Global Instant replication Fork Density (GIFD) as well as an extended S phase (Coulon et al., in preparation). GIFD is defined as number of fork divided by the total length of DNA measured (Bialic et al., 2015a). GIFD is expressed as forks/Mb corrected for the amount of DNA analyzed in cells of S phase determined by flow cytometry on the same sample.

The debate remains about the existence of a licensing checkpoint that delays S-phase entry if licensing was not complete. Some studies have shown that depletion of licensing factors induces apoptosis or G1 arrest in the untransformed human diploid fibroblasts and liver cells in a p53-dependent manner. It was associated with CDK2 phosphorylation and RB hypophosphorylation which prevented S-phase entry. Since cancer cell lines have defects in p53 pathway, they enter S phase even in the absence of licensing factors. This was associated with an increase in the phosphorylated CHK2, a marker of the DNA damage response (Feng et al., 2003;Nevis et al., 2009). On the other hand, partial depletion of MCM2 in an untransformed cell line allowed its progression into S phase. This was associated with increased frequency of anaphase bridges suggesting that replication was not efficient (Passerini et al., 2016). Similar results have been seen in mouse embryonic fibroblasts (Kawabata et al., 2011). Thus, G1 arrest or apoptosis seen in the above studies might be due to total depletion of licensing factors while their downregulation does not induce same phenotype. Accordingly, knocking down ORC in cancerous cell lines sensitizes them to hydroxyurea-induced replication stress. This also alters cellular viability which was not the case in the untransformed controls (Zimmerman et al., 2013).

An alternative way in targeting licensing for cancer therapy might be through the induction of re-replication. This would be through downregulation of geminin which should relieve the inhibition of CDT1. Geminin downregulation was shown to induce DNA rereplication selectively in some cancerous cell lines. The induction of DNA re-replication in these cells resulted in apoptosis. Phosphorylated p53 was also elevated in cancer cell lines depleted of geminin but not in the untransformed controls. The reason why re-replication was not induced in the untransformed cell lines is attributed to the fact that cyclin A-CDK2 phosphorylates CDT1

targeting it for degradation. Depletion of cyclin A along with geminin depletion from the untransformed cell lines resulted in re-replication (Zhu and Depamphilis, 2009). This suggests that targeting licensing could be a new approach in targeting malignant tumors.

Impact of Under-replication on Genome Integrity

Microsatellite and Chromosomal Instability

Genome instability is further classified into microsatellite instability and chromosomal instability. Microsatellite instability is defined by a defect of DNA mismatch repair which induces mutations in DNA coding sequences leading to alteration in the gene expression.

Replication timing was shown to be highly correlated with different types of mutation. Latereplicating regions have more chances of having a variety of point mutations compared to earlyreplicating regions. As discussed in earlier in section 2.2.2., this might be due to changes in DNA polymerases proofreading efficiency in late-replicating regions (Koren et al., 2012).

At the chromosomal level, chromosomal instability is defined by high rate of gain or loss of whole or parts of chromosomes. It also involves alterations in gene copy number and gene rearrangements which contribute to the genomic heterogeneity seen in cancer. Alterations in gene copy number include loss of heterozygosity (LOH) and gene amplifications. LOH at the RB locus is frequently seen in several types of human cancers, and it is associated with a decrease in RB expression. LOH of RB in patients with epithelial squamous cell carcinoma was frequently associated with alteration of P53. This would lead to an imbalance between proliferative activity and apoptotic control (Xing et al., 1999). Gene amplification occurs when regions from two different chromosomes remain fused together until they reach anaphase. As a result of pulling by spindle fibers, these regions may tear apart. One cell would receive two copies of the gene while the other will miss it. The constitutive breakage-fusion-bridge cycles may lead to amplifications of genes (Coquelle et al., 1997;Umbreit et al., 2020). An example of a gene that have been

shown to be amplified in cancers is the human epidermal growth factor receptor-2 (HER-2) gene seen in breast cancer and associated with poor prognosis (Jiang et al., 2012). Another type of genomic rearrangement is gene translocation which triggers hematologic tumors. It occurs due to end joining of DSB from different chromosome segments inducing their fusion and alteration of the expression of downstream gene or the production of a fusion protein. Philadelphia chromosome t(9;22) is commonly seen in acute lymphocytic leukemia and chronic myeloid leukemia. This translocation results mainly in the fusion of breakpoint cluster region (BCR) gene and abelson (ABL) gene. As a result ABL gene coding for tyrosine kinase is constitutively expressed leading to uncontrolled cellular proliferation (Achkar et al., 2010). While chromosomal translocations are usually the drivers of oncogenesis in hematological cancer, it is not considered as a main trigger in solid tumors. However, most of the other rearrangements contribute to drive genome instability seen in solid tumors. As mentioned above, microsatellite instability is more frequent in late replicating regions. Genomic rearrangements may also occur at preferential locations on chromosomes, mainly CFS. The importance of studying these regions and their implication in cancer will be elaborated in the next sections.

Characteristics of Late-Replicating Regions and Fragile Sites

The importance of studying CFS came from Yunis and Soreng study in 1984. Blood cultures of individuals with mental retardation and acute leukemia were prepared. 70-82% of the total chromosomal gaps coincide with 51 different identified CFS. This suggested that the expression of CFS contributes to genome instability seen in several types of cancer (Yunis and Soreng, 1984). Then a question arose whether the fragility induced at CFS is attributed to alteration in the replication process. Le Beau and colleagues revealed that the fragility induced at In order to further understand the fragility that occurs at CFS, several models have been proposed. The proposed causes include fork stalling (Glover et al., 2005), R-loop formation (Helmrich et al., 2011) and paucity of initiation events (Letessier et al., 2011). The first two models have been addressed in the previous sections. CFS harbor genes so long that transcription would start G2/M of the previous cell cycle, continue in S phase of the next cell cycle. The presence of such long genes increases the chances of RNA-DNA hybrids formation and replication-transcription interference. This leads to the induction of DSB. However, besides the replication-transcription conflicts, there is another reason explaining fragility at CFS. Letessier and colleagues studied the initiation events in FRA3B using combined FISH on combed DNA.

This allows single-molecule analysis of DNA within a specific region. FRA3B was known to be fragile in human lymphocytes but not fibroblasts. The study was set up in both cell lines in the presence and absence of replication stress. The results revealed the presence of initiation events within the core of FRA3B in fibroblasts but not in lymphocytes in unperturbed S phase. Thus, in order to replicate FRA3B core in lymphocytes, origins of replication flanking the core region would replicate the core itself. When cells are challenged with aphidicolin, the fibroblasts compensate to the replication stress by firing on dormant origins. However, this was not possible for the lymphocytes as the FRA3B core lacks licensed replication origins. As a result, lymphocytes would exit S phase with FRA3B being under-replicated and fragility occurs (Figure 11) (Debatisse et al., 2012;Letessier et al., 2011). This emphasizes the importance of origin licensing in preventing genome instability. If lymphocytes had enough origins licensed in the FRA3B core, instability would not occur at this site.

Mechanisms of Resolving Replication Stress

Checkpoint Activation during Replication in S Phase

During replication stress when the forks get stalled or collapsed, a series of cellular responses are activated to rescue the forks. Treating U2OS with hydroxyurea led to a reduced fork rate associated with a decrease in the number of replication foci, indicating that not all of the different clusters are activated. However, the intensity of each focus was higher with hydroxyurea-treated cells, indicating that more origins were being fired within a replication focus. This indicated that the dormant origins had been fired locally to compensate for the replication stress (Figure 12). Activation of local dormant origins and inhibition of firing of new foci were shown to be ATR/CHK1-dependent. ATR/CHK1 is known to be activated in response to single-stranded DNA sensed by RPA and acts to stabilize the forks (Ge and Blow, 2010). Several other roles of of ATR/CHK1 have been reviewed. In an unperturbed cell cycle, CHK1 has a role in the regulation of S phase, mitotic entry and spindle checkpoint. Knocking down or inhibiting CHK1 leads to accumulation of cells in S phase, increased initiation of DNA synthesis and DNA damage. In response to DNA damage, CHK1 phosphorylates CDC25 phosphatase. CDC25 phosphatase phosphorylation targets it for degradation. As a result, CDK1 and CDK2 will remain phosphorylated thus not active. This leads to cell cycle arrest. CHK1 also activates with WEE1 kinase which in turn phosphorylates and inhibits CDK1. As a result, mitotic entry is inhibited. CHK1 activation also inhibits polo-like kinase 1 (PLK1) which is a mitotic kinase. Moreover, CHK1 has a role in recruitment of translesional polymerases to repair DNA (Patil et al., 2013).

However, single stranded DNA may break and produce DSB. Even the exposure of interphase cells to radiation might result in DSB. In response to DSB, ATM/CHK2 is activated.

CHK2 is phosphorylated mainly by ATM. As a consequence, CHK2 gets activated and phosphorylates many proteins involved in a variety of cellular processes including DNA repair through HR and NHEJ in addition to other proteins involved in cell cycle regulation, checkpoints activation and apoptosis (Zannini et al., 2014).

As the name implies HR requires the presence of undamaged homologous template mainly sister chromatids. Thus, this process of repair occurs mainly in S and G2 phases. The process by which HR occurs starts with 5' to 3' resection to form 3'-single-stranded DNA using several nucleases including MRE11. RPA then binds the resected DNA to protect it from degradation. RPA is then displaced by RAD51 with the help of breast cancer gene 1 (BRCA1)

and BRCA2. Then the 3'-strand invades the homologous strand forming a D-loop then DNA polymerases extend it using the homologous strand as a template. This would create 4-stranded DNA structures called holliday junctions. These junctions will be resolved later by endonucleases or by helicase/topoisomerase. This mechanism of repair is known to be less errorprone compared to NHEJ.

On the other hand, the NHEJ pathway is a more mutagenic mechanism to repair the DSB.

Classical NHEJ requires the binding of Ku70/80 which is a heterodimer of X-ray repair cross complementing 6 (XRCC6) and XRCC5. The binding of Ku70/80 prevents DNA resection followed by DNA ligation. However, there are two alternative NHEJ pathways which are more error-prone compared to the classical one that are the micro-homology mediated end joining and repair by polymerase Ѳ-mediated end joining. These are usually activated when the classical NHEJ is impaired. Micro-homology mediated-end joining involves resection and exposure of short-strand. This is followed by cleavage of exposed 3'-end, filling the single-stranded gap and ligation. As a result, this type of repair leads to deletions and loss of genomic information.

Polymerase Ѳ-mediated end joining involves bridging of broken ends based on homology.

However, this may result in either deletions or insertions (Rodgers and McVey, 2016). The three different types of NHEJ are summarized in (Figure 13).

Resolving Under-replicated DNA during G2 Phase

In addition to inducing the checkpoint response in S phase in response to stalled or collapsed forks, ATR was shown to have a role in mitotic entry. S-phase retinal pigment epithelial cells (RPE1-hTERT) cells treated with ATR inhibitors were shown to have accelerated mitotic entry. ATR inhibition also led to premature accumulation of cyclin B through the expression of forkhead box protein M1 (FOXM1) in S-phase cells. In unperturbed S-phase, ATR is activated at a low level during early S phase and it declines at the end. Cyclin A-CDK1 then phosphorylates and activates FOXM1 allowing transition to G2 and mitosis (Saldivar et al., 2018).

Another protein involved in G2/M transition is RNA-binding protein TIAR. Knocking down TIAR from HeLa cell line resulted in premature mitotic entry associated with an increase in chromosomal breaks. It has been shown that it localizes in G2/M transition granules in G2/prophase cells. These granules are sites of active transcription around stalled replication complexes in G2 and prophase. Furthermore, TIAR sequesters CDK1 in these granules upon G2/M checkpoint activation in order to prevent mitotic entry (Lafarga et al., 2019).

Resolving Under-replicated DNA during Mitosis

However, the checkpoints mentioned in the previous sections might not be sufficient to prevent the mitotic entry with under-replicated regions. As a result, the cells evolved other mechanisms in order to finish replication in mitosis and prevent genome instability. As low levels of under-replicated regions might not be detected by ATM/ATR pathway, cells would still enter mitosis with under-replicated regions. One mechanism by which under-replicated regions are resolved in mitosis is through the fanconi D2 (FANCD2) pathway. FANCD2 was shown to be enhanced in aphidicolin-treated cells and colocalized with g-H2AX and chromosome breaks.

However, the presence of FANCD2 spots did not inhibit cellular division. Thus, FANCD2 spots were also seen on lagging chromosomes and anaphase bridges. FANCD2 spots seen on mitotic chromosomes were due to unresolved DNA replication structures from the previous S phase.

Moreover, BLM helicase which is known to resolve ultrafine anaphase bridges was shown to be enhanced in aphidicolin treated FANC-deficient cells indicating that FANCD2 has a role in recruitment of BLM to ultrafine bridges to prevent abnormal chromosome segregation (Naim and Rosselli, 2009). ERCC1 and MUS81 are endonucleases that have been shown to colocalize with FANCD2 on mitotic chromosomes. ERCC1 and MUS81 also colocalize on DNA gap from prophase till metaphase. ERCC1 and MUS81 depletion resulted in an increase in chromosomal abnormalities and mitotic catastrophes in HeLa cells both treated and untreated with aphidicolin, whereas it resulted in mitotic failure in human fibroblasts. These data suggested that these endonucleases have a role in maintaining proper chromosomes segregation. Furthermore, ERCC1, MUS81 and FANCD2 foci were colocalizing with incorporated 5-ethynyl-2'deoxyuridine (EdU) in G2 and M cells treated with aphidicolin, suggesting that they are involved in resolving late-replicating regions. In addition, ERCC1 and/or MUS81 depletion resulted in a decrease in the expression of CFS without altering replication dynamics during S phase. This suggested that these proteins are involved in resolving under-replicated regions (Naim et al., 2013).

It has also been shown that MUS81 is required to promote DNA synthesis in mitosis.

Thus, this process of DNA synthesis is referred to as mitotic DNA synthesis (MiDAS). MiDAS occurs mainly at CFS where cells enter mitosis with under-replicated DNA. Cells trigger this process in order to preserve genome integrity when earlier checkpoints have failed. This atypical DNA synthesis was shown to be POLD3 dependent. MUS81 depletion reduced POLD3 recruitment to chromatin suggesting that MiDAS is mediated by MUS81 through POLD3 (Minocherhomji et al., 2015). Later, it has been shown that the recruitment of both POLD3 and MUS81 requires the presence of RAD52 which promotes the repair of DSB through HR. RAD52

depletion led to phenotypes similar to POLD3 and MUS81 depletion where depleted cells showed increased frequencies of both bulk and ultrafine anaphase bridges. This emphasized the importance of RAD52 in initiating MiDAS (Bhowmick et al., 2016a). As a summary, under replication stress conditions the cells would enter mitosis with under-replicated DNA mainly at CFS. To prevent genomic instability, unscheduled DNA synthesis occurs in a POLD3-dependent mechanism to process these regions mainly in prophase and prometaphase to prevent anaphase anomalies such as lagging chromosomes and ultrafine anaphase bridges.

Once the cells enter mitosis, the spindle assembly checkpoint (SAC) is activated to ensure proper orientation of chromosomes on mitotic spindles. The mitotic checkpoint complex inhibits the anaphase-promoting complex (APC). The inhibition of APC prevents mitotic exit

Resolving Under-replicated DNA in the Next Cell Cycle

Despite the fact that MiDAS is capable of resolving under-replicated DNA, some underreplicated regions might escape the process and lead to ultrafine anaphase bridges formation.

Ultrafine anaphase bridges are formed because of the entangling of under-replicated chromosome regions. Upon chromosomes segregation during anaphase, the pressure exerted on these regions leads to the formation of DNA lesions inherited in both daughter cells. p53 binding protein1 (53BP1) nuclear bodies were shown to sequester DNA lesions. U2OS cells arrested in G1 phase showed persistent 53BP1 nuclear bodies. Upon S phase entry these nuclear bodies colocalized with PCNA then dissolved at the end of S phase. This confirmed that the repair of sequestered DNA lesions from the previous cell cycle requires active replication. The dissolution of 53BP1 nuclear bodies at the end of the S phase was due to the activation of local origins which is assisted by the presence of RIF1. RAD52 was shown to be associated with late stages of 53BP1 dissolution in late S phase to repair DNA lesions. Furthermore, the frequency of ultrafine anaphase bridges was measured in cells depleted of 53BP1 over two consecutive cell cycles and was compared to the control. In control cells, the frequency of ultrafine anaphase bridges was decreased in the second cell cycle compared to the first one. This was not the case in 53BP1depleted cells, supporting the role of 53BP1 in preventing ultrafine anaphase bridges (Figure 14) (Spies et al., 2019).

Hereditary Diseases Associated with Replication Components

Meier-Gorlin syndrome is a recessive autosomal disorder associated with mutations in ORC1, ORC4, ORC6, CDT1 and CDC6 components of pre-replication complex. Meier had reported the first case in 1959 and the second case was reported by Gorlin in 1975. Thus, it was referred to as Meier-Gorlin syndrome. Phenotypically the syndrome is characterized by primordial dwarfism in addition to pulmonary emphysema, skeletal abnormalities, microcephaly and others. The phenotypic features of the diseases are linked to the gene mutated. For example ORC2 mutation causes more severe growth retardation compared to others (Munnik et al., 2012).

Another genetic syndrome associated with replication process is the Bloom syndrome.

The disease was first described by Bloom in 1954 who named it as congenital telangiectatic erythema resembling lupus erythematosus in dwarfism. It is a recessive autosomal disease characterized phenotypically by facial skin lesions. Cells from these patients grown in culture displayed increased genomic rearrangements and genome instability. Bloom is a RecQ helicase protein which is involved in resolving entangled lagging chromosomes in anaphase. This highlights the importance of this protein in preventing genome instability (German et al., 2007).

Mutation in another RecQ helicase, WRN, is associated with Werner syndrome. Werner syndrome is a recessive autosomal disease characterized by premature aging. It has been discovered by Werner in 1904. Patients with werner syndrome a particularly unstable genome.

Loss of WRN helicase was shown to induce chromosomal aberrations and telomere fusion in cells with oncogenic stress compared to wild type WRN controls (Crabbe et al., 2007).

Another hereditary disease associated with replication components is fanconi anemia. It was first described by Fanconi in 1927. The disease is characterized by bone marrow failure and abnormalities in development. Twenty-two fanconi anemia genes have been identified. All of them are inherited as recessive autosomal except for FANCB, which is X-linked. Patients with fanconi anemia are more susceptible for developing cancer and display hypersensitivity to chemotherapy. The chromosomal instability seen in these patients is due to a failure to repair DNA defects. As a result, they display a high level of chromosomal breaks and aberrations (Rageul and Kim, 2020).

These suggest that patients with genetic syndromes associated with mutations in the components of replication are more prone to have genome instability and cancer except for patients with Meier-Gorlin syndrome. Although the life span of these patients is usually normal, it has not been shown whether these patients are more prone to cancer or not. In addition, how alteration in DNA licensing induces developmental defects is not clear. One hypothesis could be that alteration in DNA licensing in these patients affect cellular division leading to a delay in development. Given the fact that the frequency of this disease is really low there might be not enough cases to study the prevalence of cancer in these patients.

Under-replicated Regions from an Evolutionary perspective

Late replicating regions, mainly CFS, are highly unstable and they represent preferential sites of sister chromatid exchange, translocations, deletions and retroviral integration. The pattern of human papilloma virus integration in human genome of 26 cervical tumors has been studied. Viral integration was seen in 14 different chromosomes on the same loci. Most of these loci colocalized with CFS (Thorland et al., 2003). Furthermore, it has been mentioned previously that the fidelity of the polymerases and checkpoints diminish with late replication. The conservation of CFS between species, also raises the question of the importance of these sites in maintaining polymorphism and genome flexibility. Whether mammalian evolution could be partly driven by CFS has been addressed by Ruiz-Herrera and colleagues. The study revealed that the distribution of 2,304 evolutionary breakpoints was concentrated at specific chromosomal locations coinciding with CFS. This emphasizes the importance of CFS expression in evolutionary genome reorganization (Ruiz-Herrera et al., 2006).

All of these suggest that CFS are a two-edged sword. They are necessary for genome flexibility which is important for speciation and evolution. However, at the same time they put the cells at risk because of genome instability which would induce carcinogenesis.

Transfection of U2OS

150 mM NaCl containing 2µg of DNA plasmid were added to 150 mM NaCl containing 4 µl JET-PEI (Polyplus). The solution was incubated at room temperature for 20 minutes. Fresh medium containing the transfection solution was added to the cells and incubated for 4-6 hours.

JET-PEI containing-medium was then replaced by fresh medium.

Measuring intermitotic time using Time-lapse microscopy

Untransformed and cancer cell lines were grown on microscopy dishes and were imaged every 10 min for 72h on a widefield inverted microscope, either by phase contrast or by fluorescence using the vital DNA stain SiR-Hoechst (SC007, Tebu) at 100 nM. SiR-Hoechst stain was added 4 hours before acquisition. Cells were maintained in 37°C humid incubator in the presence of 5% CO2 and 21% O2.

DNA Preparation for Molecular Combing

DNA Labeling with Thymidine Analogues

Asynchronous cells growing in exponential phase were pulsed with 25µM IdU (Sigma) for 15 minutes. Medium containing IdU was removed and replaced by warm medium containing 200µM CldU (Sigma) for 15 minutes. Except for the U2OS, IdU and CldU pulses were performed using 30 minutes interval. Medium was also removed and replaced by warm medium containing 200µM thymidine (Sigma) for 1-2 hours. Cells were trypsinized, resuspended with cold PBS 1X and counted. 125,000 cells were used to prepare agarose plugs for molecular combing. The rest of cells were centrifuged and fixed with 96% of ethanol for determining the fraction of cells in S phase.

DNA Preparation in Agarose Plugs

Low-melting agarose plugs were prepared by counting 25,000 cells per plug and added to an equal volume of 1% low-melting point agarose. 100 µL of mix was casted in the mold. Plugs were left to solidify at 4°C for 30 minutes and then extruded in proteinase K buffer containing 0.2 mg/mL proteinase K (Sigma) and incubated at 37°C overnight. Buffer was removed gently and plugs were washed 5 times with TNE50 buffer (1 hour each) at RT. Plugs were stored in TE buffer at 4°C. Coverslips were washed 3 times with PBS-0.05% triton X-100, 3 minutes each. DNA was immunodetected as in previous steps using primary antibody (mouse anti-ssDNA, Dshb autoimmune ss-DNA, 1/100) and secondary antibody goat anti-mouse IgG2a Alexa647 (A21241, Life Technologies, 1/50 dilution).

DNA coverslips were mounted with 10 µl of ProlongGold anti-fade reagent (Molecular Probes) and were left to polymerize at RT overnight (Bialic et al., 2015a).

Molecular Combing Microscopy

The acquisition of DNA combing images was done using an epifluorescence Zeiss AxioImager Z1 microscope equipped with scMOS Zyla 4.2MP camera (2048x2048 pixels) using 40x objective.

For measuring the GIFD, images were acquired without bias for IdU/CldU, based solely on DNA fibers (Cy5 channel). About 50 images, depending on DNA density on coverslips, were taken per sample to get more than 90 Mb of DNA to be analyzed. For fork velocities, IdU tracks were measured.

Image Analysis of DNA Combing

DNA combing images were analyzed using IDeFix (Intelligent DNA Fiber Identification and Analysis software) developed in the lab. The software allows the measurement of fiber length in Kb.

Flow Cytometry

Flow Cytometry for S-Phase Fraction in the Cell Population

Since the combing experiments were performed on asynchronous cells, GIFD measurements require the knowledge of the percentage of cells incorporating the nucleosides analogues. About 10 6 cells were fixed in 96% cold ethanol and stored at 4°C. For IdU detection, cells were rehydrated by an equal volume of PBS 1X, washed again and DNA depurinated with 2N HCl, 0.5% Triton X-100 for 30 minutes. Cells were washed with PBS, spun at 500g, neutralized with sodium borate pH8.5, then incubated with anti-BrdU antibody (347580, Becton

Dickinson, 1/30) diluted in PBS 1X, 1% BSA, 0.5% Tween 20. Cells were washed and incubated with Alexa 488 Fab goat anti-mouse (A11017, Thermo Fisher, 1/1000) diluted in PBS 1X, 1% BSA, 0.5% Tween 20. Cells were washed and incubated overnight at 4°C with 7aminoactinomycin D (7-AAD) solution (Sigma) containing 200 µg/mL RNase (Sigma). Cell aggregates were removed by filtering using a 70 µm mesh and acquisition was carried out using BD FACS CantoII instrument.

Estimation of the DNA Amount of Cancer Cell lines Using Flow Cytometry

HCT116, U2OS, PC-3, HCC-1937, MK-45, 22RV1, HCC-38 and BXPC-3 were cultured and collected along with the untransformed cell line RPE1-hTERT. Cells were fixed with 70% ice-cold ethanol. Cells were then counted and around 1,200,000 million cells were split into four tubes. This was followed by staining with 0.5 (1X), 1.5 (3X), 4.5 (9X) and 13.5 (27X) µg/ml 4',6-diamidino-2-phénylindole (DAPI) (Sigma). Cell were then washed and filtered using 70 µm mesh and acquisition was carried out using BD FACS CantoII instrument. The FACS parameters were kept the same for all the RPE1-hTERT and the cancer cell lines analyzed. DNA amount was then estimated based on the shift in G1 peak of the cancer cell lines compared to the G1 peak of the diploid RPE1-hTERT per each DAPI concentration. The ratio of G1 peak shift of cancer cell lines to the G1 peak of RPE1-hTERT was determined.

Measurements of S-Phase Duration by Flow Cytometry

1.6.3.1. Principle
The measurement of S-phase duration by double labeling using a protocol previously developed in the lab (Bialic M. et al., in preparation).

Asynchronously Cells were filtered before the acquisition using 70 µm mesh filter. The acquisition was carried on using BD FACS CantoII. Flow cytometry analysis was done using FlowJo software.

The percentage of cells incorporating both EdU and BrdU at each time point was analyzed and plotted as linear regression on a graph as a function of thymidine chase times. The S-phase duration is defined as the intersection of the plot with the x-axis.

Percentage of Mitotic Cells Undergoing MiDAS in Cancerous and Untransformed

Cell Lines

Cell Culture and Immunodetection

Cells were cultured in glass bottom 24 well plate (662892, Greiner bio-one).

Exponentially growing cells were pulsed with 10 µM EdU for 15 minutes. Cells were fixed with an equal volume of 7.4% formaldehyde for 15 minutes at RT (medium was not removed to prevent the loss of mitotic cells). Cells were washed twice with PBS 1X then permeabilized with PBS 1X, 0.2% Triton-X100 for 10 minutes at RT followed by 2 washes of PBS 1X. Cells were blocked with PBS 1X, 1% BSA, 0.5% Tween 20 for 30 minutes at RT. For EdU detection, a volume of 25 µL Click-it reaction solution containing 2 mM CuSO4, 10 mM ascorbic acid, 0.02 mM of di-sulfo-cyanine5 azide was added to each well and covered with a cover glass for 45 minutes in humid chamber at 37 ºC. Cells were washed 3 times with PBS 1X then incubated in rabbit anti-phosphoH3 antibody (06-570, Millipore, 1/1000) diluted in PBS 1X, 1%BSA, 0.1% Triton X-100 for 1 hour at 37 ºC. Cells were washed 3 times with PBS 1X, 0.05% Triton and incubated in Alexa488 goat anti-rabbit (A11034, Molecular probes, 1/2000) for 1 hour at 37ºC.

Cells were washed 3 times with PBS 1X, 0.05% Triton X-100, and DNA was stained with 500 µl of 0.5 µg/mL DAPI then stored at 4 ºC. For the negative control, EdU was not added to the medium before fixation. The following steps were as described above.

Microscopy Screening Using Opera Phenix

PhosphoH3 positive mitotic cells were pre-scanned using a 10x objective followed by confocal scanning (13 Z stacks 0.1 µm each) using a 63x objective. Stacks were merged as maximum intensity projection to check for the presence and number of EdU foci on mitotic cells.

Negative control well lacking EdU was also scanned. The EdU positive threshold was set above the intensity of EdU signal in the negative control. Mitotic cells incorporating EdU were scored automatically. The percentage of EdU incorporating mitotic cells from the total analyzed mitotic cells was determined. In order to test for the sensitivity of the assay, all of the phosphoH3 positive mitotic cells selected as EdU positive were checked for positivity and 50 random phosphoH3 positive mitotic cells were checked for the absence of EdU using Harmony program.

Analysis of Mitotic Cells for Mitotic Defects

From these thousand cells analyzed per each cell lines, two hundred mitotic cells were randomly selected. Mitotic cells were analyzed for mitotic defects according to the following categories: lagging chromosomes, chromosome bridges, failure of alignment, uncondensed DNA and others.

G1 Rescue with Palbociclib Treatment

HCT116 Rb positive cells were treated with PD 0332991 (S1579, Sellechem) for 24 hours with different concentrations: 0.16, 0.32, 0.625 and 1 µM followed by release in fresh medium and nucleosides pulses according to the FACS protocol mentioned above. Using 0.16 µM concentration, G1 fraction was increased by almost 20% without completely blocking the cells. RPE1-hTERT (Rb positive), HCT116 (Rb positive) and HeLa (Rb negative) were treated with 0.16 µM PD for 24 hours. Cells were then released and pulsed with IdU then CldU for DNA molecular combing according to the protocol mentioned above.

1.9. MCM Loading Assay by Flow Cytometry

Principle

The principle of this technique is based on the fact that during G1 MCM is loaded to the chromatin and become bound to it and then the MCM gets deloaded in S phase as forks from opposite directions merge. This increase in MCM binding to chromatin during G1 and deloading in S phase can be visualized using CSK buffer containing triton as a detergent. This permits the removal of non-bound MCM and only MCMs that are bound to the chromatin will visualized using this assay after immunodetection with MCM-specific antibodies. This permits to visualize the difference in the amount and the rate of MCM loading on the chromatin.

Chromatin Extraction and Fixation

Cancer cell lines were untreated, treated with DMSO or 0.16 µM PD for 24 hours. Cells were collected, and the pellet was either frozen at -80 ºC or processed directly. The pallet was then treated with ice cold cytoskeletal (CSK) buffer containing 10 mM PIPES pH=6.8 (P1851, Sigma), 100 mM NaCl, 300 mM sucrose, 1 mM MgCl2, 0.3% triton X-100, 0.5 mM dithiothreitol (DTT), 1 mM ATP, protease inhibitors (11873580001, Sigma) and ddH2O to complete the volume. CSK buffer should be freshly prepared. DTT, ATP and protease inhibitors were added at the last step before use. A volume of 100-500 µl of buffer was used to resuspend the pellet gently. Pellet was incubated on ice for 10 minutes then centrifuges at 2000 g for 7

minutes. The pellet was washed again with CSK after incubation on ice for 10 minutes. The pellet was then resuspended with 1 ml PBS-1X followed by fixation with ice cold methanol of 70% total concentration.

Immunodetection of the

Molecular DNA Combing

Detection of Fork Pausing Using Molecular Combing

Asynchronously growing MEFs were treated with 500 nM pyridostatin (CT-PYRI, Chemietik). Cells were then pulsed with 25 µM IdU (Sigma) for 15 minutes in medium containing 500 nM pyridostatin. Medium was then replaced by warm medium containing 200 µM CldU (Sigma) for 60 minutes and 500 nM pyridostatin. Medium was then replaced by 200 µM thymidine containing medium for 2 hours. Plugs preparation, DNA combing and immunodetection was done as described in section 1.2.

Asynchronously growing MEFs and mouse ESC were pulsed with 25 µM IdU (Sigma)

for 7 or 15 minutes. Cells were pulsed with 200 µM CldU (Sigma) for 60 minutes followed by 2 hours of 200 µM thymidine (Sigma) chase. Plugs preparation, DNA combing and immunodetection was done as described in section 1.2.

Testing for Variation in Origin Distribution in MEFs and Mouse ESC

Asynchronously growing MEFs were treated with 20 nM trichostatin A (TSA) (T8552, Sigma) for 1, 4, 5 and 10 hours. Cells were released in fresh medium containing 25 µM IdU (Sigma) for 15 minutes followed by 200 µM CldU (Sigma) containing medium then 200 µM thymidine (Sigma) chase. Plugs preparation, DNA combing and immunodetection was done as described in section 1.2.

Asynchronously growing MEFs were pulsed with 25 µM IdU (Sigma) for 15 minutes.

Cells were pulsed with 200 µM CldU (Sigma) for 15 minutes followed by 2 hours of 200 µM thymidine (Sigma) chase. Cells were collected and fixed with 70% cold ethanol. Cells were then rehydrated and stained with 0.5 µg/ml DAPI (Sigma). Cells were then sorted based on their DNA content into early S1, mid S2 and late S3 cells. Cell sorting was carried on using BD FACS ARIA IIU. Cells were then collected and prepared in plugs. DNA preparation and Immunodetection was done as described in section 1.2.

Project 1: Causes and Consequences of Altered DNA Replication

Dynamics in Cancer Cells

In this part of my project I studied DNA replication dynamics in a variety of cancer cell lines and compared them to untransformed controls. The results I have obtained in this part represent the core of my thesis and my main contribution during the four years of my Ph.D. The results will be presented in details in my first scientific article.

Aims and Objectives

Most if not all solid tumors show genetic or epigenetic alteration of one or more G1/S cell cycle regulators and display genome instability. However, it is not yet clear how these two events are linked. Based on previous data from the lab on yeast and mouse models, we showed that upon G1/S deregulation cells enter S phase with fewer number of origins and spend more time to replicate the genome.

1) The first aim of my thesis was to see if these phenotypes (low GIFD, long S) are frequent or prominent in cancer cells.

2) Determine the consequences of decreased replication origin usage and S-phase extension on doubling time in cancer cell lines, using time-lapse microscopy.

3) Rescue the decrease in the replication potency in one of the tested cancer cell lines by prolonging G1 using Palbociclib, a cdk4/6 inhibitor, and check whether this would restore normal S phase length. Despite their molecular and cytogenetic differences, several types of cancers share common properties referred to as hallmarks of cancer (Hanahan and Weinberg, 2011). Among these hallmarks, replication stress and genome instability have been extensively studied during the last decade. Faithful genome duplication requires setting up a sufficiently large number of replication origins followed by a well-orchestrated activation during S phase of a fraction of these licensed origins so to complete chromosome replication before mitosis begins [START_REF] Zhao | High-resolution Repli-Seq defines the temporal choreography of initiation, elongation and termination of replication in mammalian cells[END_REF]. Genome instability is defined as rapidly acquiring genomic changes such as copy number variations, translocations or complex chromosome rearrangements that can provide selective advantages to cells exposed to various stresses [START_REF] Tubbs | Endogenous DNA Damage as a Source of Genomic Instability in Cancer[END_REF]. A large body of work has shown that cells are normally well equipped to defeat endogenous and exogenous replication stress, which is recognized as the major fuel that drives genome instability. However, a small number of cells escape these controls, are transformed, become genetically unstable and able to create the genetic diversity that nurtures selection and tumor progression.

Why and how a few cells in a large population of unchallenged cells start behaving differently and become cancerous has been difficult to address via ensemble cellular and biochemical methods.

Chromosome replication is an inherently stochastic and plastic process, which inevitably will generate at a low rate some variant, from which evolution proceeds. Once per cell cycle DNA replication involves two mutually exclusive steps operating at different phases of the cell cycle (Diffley, 2011). The first step entails licensing of replication origins in G1 by sequential loading of the origin recognition complex (Orc1-6), the Cdc6 and Cdt1 licensing factors, and two inactive Mcm2-7 replicative helicases.

This assembly is referred to as the pre-replication complex (pre-RC). The second step consists in activating these pre-RCs and occurs throughout S phase, when the S phase cyclin-dependent kinases (S-CDK) and Dbf4-dependent kinases (DDK) are fully active. This leads to loading and activation of the Cdc45-MCM-GINS (CMG) complex, DNA unwinding, recruitment of DNA polymerases and bidirectional DNA synthesis from origins. Crucially, the presence of high CDK activity during the S and G2

phases inhibits the formation of further pre-RCs in order to prevent DNA re-replication (Arias and Walter, 2007;Nishitani and Lygerou, 2002;Truong and Wu, 2011). Thus, the loading of Mcm2-7 is restricted to G1, where cells can load an excess of Mcm2-7 complexes that act as dormant origins, which are needed under situations of replication stress to resume DNA synthesis between stalled converging forks (Alver et al., 2017;Ge et al., 2007;Ibarra et al., 2008). The importance of efficient origin licensing has also been studied in embryonic stem cells (ESC). The fast loading of Mcm2-7 complexes in ESC is attributed to increased levels of chromatin bound Cdc6 and Cdt1. This contributes to efficient origin licensing despite the short length of G1 phase. Alteration in the G1 kinetics was shown to be associated with cellular differentiation (Ballabeni et al., 2011;Coronado et al., 2013;Fujii-Yamamoto et al., 2005;Matson et al., 2017). The exact nature of replication origins is still debated as their specification involves chromatin modifications subjected to dynamic changes and cell-type specific adaptations [START_REF] Ganier | Metazoan DNA replication origins[END_REF][START_REF] Hyrien | Peaks cloaked in the mist: The landscape of mammalian replication origins[END_REF]. In human cells up to 100,000 sites can be used as replication origins, from which less than 20,000 are used in every cell [START_REF] Kumagai | Binding of the Treslin-MTBP Complex to Specific Regions of the Human Genome Promotes the Initiation of DNA Replication[END_REF][START_REF] Prioleau | DNA replication origins-where do we begin?[END_REF].

Very few studies have emphasized the importance of origin under-licensing in inducing genomic aberrations [START_REF] Limas | Preparation for DNA replication: the key to a successful S phase[END_REF]. Seminal work in yeast has shown that loss of a Cdk inhibitor or overexpression of G1 cyclins, both causing precocious Cdk activation and G1 shortening, impede full origin licensing, leading cells to proceed in S phase with fewer active origins, which resulted in chromosomes breaks, rearrangements and mis-segregation (Lengronne and Schwob, 2002;Tanaka and Diffley, 2002). Similarly, overexpression of cyclin E in mammalian cells led to decreased Mcm2-7 loading onto chromatin, a lower number and intensity of replication factories, as well as an increase in -H2X foci that denote replication stress (Ekholm-Reed et al., 2004). These results identified origin licensing defects as a source of genome instability. However, whether altered origin licensing and changes in replication dynamics are a common endogenous feature of cancer cells has not been thoroughly addressed, particularly in unperturbed conditions.

In order to get insights into how cancer cells replicate their genome, we determined several parameters of DNA replication on long single DNA molecules labelled in vivo with thymidine analogs and stretched by DNA combing (Bensimon et al., 1994, Bialic et al., 2015), in fourteen different cancer cell lines compared to three untransformed cell lines. We also determined the total duration of S phase in these cells, using a newly developed method (Bialic et al, submitted), along with mitotic DNA synthesis (MiDAS) (Bhowmick et al., 2016;Minocherhomji et al., 2015) and chromosome segregation defects. We find, strikingly, that all tested cancer cell lines display a decreased capacity to synthesize DNA, have a longer S phase and an increased frequency of MiDAS and lagging chromosomes during anaphase.

Imposing a longer G1 phase in Rb-positive HCT116 cancer cells with low doses of the Cdk4/6 inhibitor Palbociclib (Fry et al., 2004) restored normal S-phase dynamics, indicating that replication origin underlicensing is a likely cause for the extended S phase and genome instability of cancer cells.

MATERIALS AND METHODS

Culture of cancer cell lines

Eleven

Culture of untransformed cell lines

Primary human fibroblasts (BJs) were cultured in DMEM/Glutamax-I supplemented with 10% fetal bovine serum (Eurobio) and 100 U/mL penicillin-streptomycin, 1 mM pyruvate and 2 mM L-glutamine.

Immortalized RPE1-hTERT cells were cultured in DMEM-F12 without glutamine, supplemented with 10% fetal bovine serum (Eurobio) and 100U/mL penicillin-streptomycin and 2mM L-glutamine.

Immortalized HMEC-hTERT cells were cultured in serum-free MEGM medium (Lonza) supplemented with bullet kit containing 5 ng/mL epidermal growth factor, 0.5 µg/mL hydrocortisone, 5 µg/mL insulin, 70 µg/mL bovine pituitary extract, 5 µg/mL transferrin, 10 -5 M o-phosphoethanolamine and 100 U/mL penicillin-streptomycin.

DNA preparation for molecular combing

Preparation of genomic DNA for combing and immunodetection was performed as described (Bialic et al., 2015). Asynchronous cells growing exponentially were pulsed sequentially with IdU

Determination of G1, S and G2+M phase fractions

Asynchronously-growing cells labelled with IdU-CldU were harvested and fixed with cold ethanol, rehydrated with PBS, and DNA was denatured with 2N HCl, 0.5% Triton X-100 for 30 min. Cells were washed, spun at 500g, neutralized with sodium borate pH8.5, then incubated with anti-BrdU antibody (347580, Becton Dickinson). Cells were washed and incubated with Alexa 488 Fab goat anti-mouse (A11017, Thermo Fisher, 1/1000). DNA was stained with DAPI solution (Sigma, 0.5 µg/ml) containing 200 µg/mL RNase. Cell aggregates were removed by filtering using a 70 µm mesh and acquisition was carried out using Becton Dickinson FACS CantoII. The fraction of cells in G1, S and G2+M was calculated based on IdU-DAPI bivariate analysis for each experiment.

Measurement of replication parameters by DNA combing

Global instant fork density (GIFD, in forks/Mb) was calculated by dividing the number of bicolor forks by the total length of all DNA molecules analyzed, and normalized for the fraction of cells in S phase for each experiment (see Fig 2B). This value was then corrected for the fraction of cells in G1 and G2+M to take into account the doubling of DNA in the latter, according to the formula: GIFD = fork density x (0.66x G1% + S% +1.33xG2+M)/S%. Fork velocity (FV, in kb/min) was obtained by dividing the length of the IdU track (in kb) by the pulse time for each bicolor track. Because a low fork density is sometimes compensated by increased fork velocity (Conti et al., 2007), we propose here a new descriptor of the DNA synthesis capacity of cells, named replication potency (RP), which combines GIFD x FV (see results).

Estimation of DNA content of cancer cells using flow cytometry

HCT116, U2OS, PC-3, HCC-1937, MK-45, 22RV1, HCC-38 and BXPC-3 cells were cultured and collected along with the untransformed cell line RPE1-hTERT. Cells were fixed with 70% ice-cold ethanol, counted and 1.2 x 10 6 cells were split into four tubes. Cells were stained with 0.5 (1X), 1.5 (3X) and 4.5 µg/ml (9X) DAPI (Sigma), washed and filtered on a 70 µm mesh and acquired on a Becton

Dickinson FACS CantoII. The FACS settings were kept identical for all the cell lines analyzed. The DNA content of cancer cell lines was estimated based on the shift of the G1 peak compared to the G1 peak of the diploid RPE1-hTERT for each DAPI concentration.

Measurements of S-phase duration

Asynchronously-growing cells were pulsed sequentially with EdU (10 µM, 30 min) then BrdU (100 µM, 30 min) plus nocodazole (100 ng/mL) for the first time point. For the other time points, the EdU and BrdU pulses were separated by thymidine chases (200 µM) of increasing times. Cells were harvested and fixed with cold ethanol, rehydrated with PBS and treated with 2N HCl, 0.5% Triton X-100 for 30 min, then neutralized with sodium borate. For EdU detection, cells were blocked and incubated in a Click-it reaction containing 2 mM CuSO4 (Sigma), 10 mM ascorbic acid (Sigma) and 0.02 mM di-sulfocyanine 5 azide (Interchim, FP-1JV6320). Cells were washed then incubated in mouse anti-BrdU solution (MoBu-1, 11-286-C100, Exbio, 1/1000) followed by Alexa 488 Fab goat anti-mouse (A11017, ThermoFisher, 1/1000). Cells were washed and incubated in DAPI (0.5 µg/mL) containing 200 µg/mL RNase A, then filtered. The percentage of cells incorporating both EdU and BrdU at each time point was analyzed and plotted as linear regression on a graph as a function of thymidine chase times. The S-phase duration is defined as the intersection of the plot with the x-axis.

Measuring intermitotic time using Time-lapse microscopy

Untransformed and cancer cell lines were grown on microscopy dishes and were imaged every 10 min for 72h on a widefield inverted microscope, either by phase contrast or by fluorescence using the vital DNA stain SiR-Hoechst (SC007, Tebu) at 100 nM. SiR-Hoechst stain was added 4 hours before acquisition. Cells were maintained in 37°C humid incubator in the presence of 5% CO2 and 21% O2.

Assessing cell cycle entry using flow cytometry

Cells were grown exponentially then pulsed with BrdU (20 µM, 0, 6, 12, 18, 24 and 36 h). Cells were then collected, fixed with 70% cold ethanol and rehydrated. Then cells were immunodetected for BrdU according protocol mentioned above for determimning the different fractions of the cell cycle.

Percentage of mitotic cells undergoing MiDAS

Cells were grown exponentially in glass bottom 24 well plates (662892, Greiner bio-one), pulsed with EdU (10 µM, 15 min), fixed with an equal volume of 7.4% formaldehyde, washed twice and permeabilized with PBS, 0.2% Triton-X100 for 10 minutes at RT. Cells were blocked with PBS 1X, 1% BSA, 0.5% Tween 20. Click-it reaction was used for EdU detection. Cells were washed and incubated in rabbit anti-phosphoH3 antibody (06-570, Millipore, 1/1000 dilution) followed by Alexa488 goat antirabbit (Molecular probes, A11034, 1/2000 dilution). DNA was stained with DAPI (0.5 µg/mL). For the negative control, EdU was omitted to the medium. PhosphoH3-positive mitotic cells were spotted on an Opera Phenix HCS microscope on pre-scan mode using a 10x objective, followed by confocal imaging (13 z stacks, 0.1µm) of mitotic cells using a 63x objective. EdU foci present in mitotic cells were identified on maximum intensity projection images. The EdU-positive threshold was set above the intensity of EdU signal in the negative control, and the percentage mitotic cells undergoing DNA synthesis was scored automatically from the total mitotic cells analyzed. All of the phosphoH3-positive mitotic cells selected as EdU positive were rechecked visually for nuclear localization and colocalization with DNA. Fifty random phosphoH3 positive mitotic cells were also scored for the absence of EdU signal. About 1,200 to 1,500 mitotic cells were analyzed for each cell line to detect MiDAS and chromosome segregation defects.

G1 lengthening with low-dose Palbociclib

RPE1-hTERT, HCT116 and HeLa cells were exposed to DMSO (vehicle) or 0.16µM PD 0332991 (S1579, Sellechem) for 24 hours, followed by release in fresh medium with nucleosides pulselabelling as described above. G1 extension was monitored by the percentage of cells in G1 by flow cytometry, and the absence of cell cycle arrest by exposing cells for 24h to BrdU.

Transfection of U2OS

150 mM NaCl containing 2µg of DNA plasmid were added to 150 mM NaCl containing 4 µl JET-PEI (Polyplus). The solution was incubated at room temperature for 20 minutes. Fresh medium containing the transfection solution was added to the cells and incubated for 4-6 hours.

JET-PEI containing-medium was then replaced by fresh medium.

MCM chromatin loading

Cell lines were treated with DMSO or 0.16 µM PD for 24 hours, collected and either processed directly or frozen at -80ºC. The cell pellet was then treated with 100-500 µl of freshly prepared ice cold cytoskeletal (CSK) buffer containing 10 mM PIPES pH6.8 (P1851, Sigma), 100 mM NaCl, 300 mM sucrose, 1 mM MgCl2, 0.3% Triton X-100, 0.5 mM dithiothreitol (DTT), 1 mM ATP, protease inhibitors (11873580001, Sigma) and ddH2O to complete the volume. DTT, ATP and protease inhibitors were added at the last step before use. The cell pellet was incubated on ice for 10 minutes, then centrifuged at 2000 g for 7 min. The pellet was washed again with CSK after 10 min on ice, then resuspended with 1 ml PBS-1X followed by fixation with ice 70% cold methanol.

RESULTS

Cancer cells have a longer cell division time than untransformed cells

Cancer cells are often portrayed as having a higher proliferation rate than untransformed cells.

This was analyzed most of the time at the cell population level and whether individual cancer cells divide more rapidly than untransformed cells is not known. Here we monitored inter-mitotic times of individual cells by time-lapse microscopy and cell tracking in eight epithelial cancer cell lines derived from six different tissues (U2OS, HCT116, Colo320, HeLa, 786-O, PC-3, 22RV1, MKN-45) in comparison to immortalized but untransformed cell lines RPE1-hTERT and BJ-hTERT (Supp Table I). Cells were grown on microscopy dishes and imaged every 10 min for 72h on a widefield inverted microscope, either by phase contrast or by fluorescence using the vital DNA stain SiR-Hoechst at 100 nM (Lukinavičius et al., 2015) (Supp Movies 1, 2). Since some of the cell lines showed longer intermitotic times in the presence of SiR-Hoechst than without, indicating photosensitivity or toxicity at least in our illumination conditions, we present here only the data obtained without SiR-Hoechst and without fluorescence illumination. In order to limit bias for cells with short intermitotic times, most cells were followed over at least two consecutive cell cycles (three mitoses). Figure 1A shows that cancer cells usually display higher heterogeneity (between 12 and 47 h) in intermitotic times compared to untransformed cells (13-28 h).

While HCT116, Colo320 and 786-O had median intermitotic times within the range of untransformed cells (16.5-18 h), the five other cancer cell lines had significantly longer cell doubling times (median of 18.3-22.1 h) compared to untransformed cells. Although there is considerable heterogeneity from cell to cell (or from one cycle to the next) in every cell line, we conclude that cancer cell lines taken as a group have longer intermitotic times than untransformed cells. This suggests that transformed cells, at least a significant fraction of them, struggle to complete their cell cycle.

One way to reconcile the notion of high proliferation rate for cancer cells with their longer intermitotic times is to propose that cancer cells stop dividing less often than untransformed cells that need to integrate mitogenic stimuli before they can commit to a new division cycle. To test this hypothesis, we grew untransformed (BJ-hTERT, HMEC-hTERT) and transformed (U2OS, HCT116, HeLa) cells with 20 µM BrdU for 6 to 36 hours and monitored by flow cytometry the fraction of cells that did not undergo S phase (BrdU-negatives). A significant fraction (18%) of untransformed BJ-hTERT cells were still BrdUnegative cells after 30h, while 100% of HCT116 were BrdU-positive after 24 hours (Fig. 1B). Additional data is being introduced here; Fig1C will be a graph showing for how long each cell line still contains BrdU-neg, i.e. non-cycling cells. While all 3 cancer cell lines become fully BrdU-pos in less than 20h, BJ and HMEC cultures contain cells that are significantly slower (G0 or long G1). We conclude that untransformed cells have shorter division cycles but frequently pause between cycles whereas cancer cells cycle continuously, albeit at a slower pace. The increased proportion of cells in S phase (Fig. 1C) suggests that cancer cells might have a longer S phase, but this could also simply be due to a shortening of G1 and/or G2+M phases.

S phase is extended in cancer cells

To test directly whether cancer cells take longer to replicate their genome, we turned to a method newly developed in the lab (Bialic et al., submitted) that can measure S-phase duration in exponentially growing cell populations, without having to rely on synchronization procedures that perturb cell cycle kinetics. This method is based on dual pulses of EdU and BrdU separated by a thymidine chase of increasing times, the rationale being that the fraction of EdU-BrdU double-positive cells become null when the thymidine chase time equals the duration of S phase (Fig. 2A). These two thymidine analogs are detected using orthogonal methods, click reaction and immuno-staining, respectively, allowing populations of single and double-positive cells to be quantified accurately by flow cytometry (Fig. 2B).

Plotting the fraction of double-positive cells against dT chase times produces a linear regression whose intersection with the x-axis provides S-phase duration (SPD, Fig. 2C). This method is robust and produced consistent SPDs of around 8h for three untransformed cell lines (BJ, hMEC-hTERT, RPE1-hTERT). We expanded this analysis to fourteen cancer cell lines (Supp Table I), which strikingly all displayed longer S-phase durations (from 10.5 to 29.8h) compared to the untransformed cells (Fig. 2D, Supp. Fig. 3). Most cancer cell lines have a S phase 30% to 90% longer than untransformed cells, yet within the range of increased intermitotic times calculated above. Two cancer cell lines (HCC-1937, ASPC-1) displayed a much longer S phase, 23 and 30h respectively, and their intermitotic times could not be measured using our setup, suggesting that they are very long. The SPD of cancer cell lines were generally more variable between duplicates than untransformed cells, suggesting that the former might be more sensitive to external conditions or evolve more rapidly in culture. We conclude that all tested cancer lines have longer S phases and seem to struggle to duplicate their genome in time before mitosis.

The density of replication forks is lower in cancer cells

In order to identify the cause for the S-phase extension in cancer cells, we used the currently most sensitive method to assess replication dynamics in single cells, that is single-molecule analysis of DNA replication by DNA combing (Bensimon et al., 1994;Bialic et al., 2015;Lengronne et al., 2001). The fourteen exponentially growing cancer cell lines, along with three untransformed cell lines, were sequentially pulsed with IdU and CldU (15 min each), then long individual DNA molecules were stretched by DNA combing on silanized coverslips and replication forks detected with anti-IdU (red) and anti-CldU (green) antibodies along DNA fibers stained in blue (Fig. 3A). Divergent red-green signals can identify replication origins (Fig. 3B), and the presence of two or more such signals on the same fiber allows to measure inter-origin distances (IODs) that have been used as a proxy for origin density on chromosomes. Because IODs are a local measure that is limited by the size of the DNA fibers (Técher et al., 2013), we prefer to use the global instant density of replication forks (GIFD, in forks/Mb), which is the number of forks divided by the total length of DNA analyzed and normalized for S-phase cells, as a proxy for origin usage genome-wide (Bialic et al., 2015). Between 467 and 1,200 Mb of single DNA molecules were analyzed for each of the seventeen cell lines, in three biological experiments (Supp Table II). The GIFD was between 1.2 and 1.9 forks/Mb for the control cells, but significantly lower (between 0.6 and 1.2 forks/Mb) for the transformed cell lines, indicating that cancer cells likely replicate their chromosomes from fewer active origins (Fig. 3C).

We also measured the speed at which individual replication forks progress in the same cell lines. The results indicate that individual fork velocities (FV) are highly variable over a 10-fold range within the same cell line, and that each cell line has its own range of FV. The mean FV was between 1.0 kb/min (hMEC-hTERT) and 1.55 kb/min (BJ) for untransformed cells, and within 0.6 and 1.5 kb/min for the cancer cell lines without any specific trend (Supp Fig. 1 supp table IV). It has been shown that dNTP pool availability is a key factor for regulating FV (Bester et al., 2011;Malinsky et al., 2001); the various FV therefore probably represent the metabolic and biosynthetic state of each cell line.

Replication potency, an integer of fork density and velocity, is decreased in cancer cells

Several studies provided evidence for replication plasticity where origin density and fork velocities are inversely correlated (Conti et al., 2007;Courbet et al., 2008). It was therefore possible that cancer cell lines with low fork density would compensate by increasing fork speed. To test this, we define here a new descriptor for the strength at which cells can synthesize DNA. We name it replication potency (RP), which is calculated by multiplying GIFD with FV, and which represents the amount of DNA (in kb) a cell in S phase can synthesize per minute for every Mb of its genome. Interestingly, this parameter is calculated for every Mb of DNA and is therefore independent of genome size, which can be highly variable in the various cancer cell lines (Supp Fig. 2, Supp Table V). Calculating the RP for the fourteen cancer cell lines was revealing, because all were significantly lower than in the untransformed cell lines (Fig. 3D). Some cancer cell lines including PC-3, 22RV1 and MKN-45 had average fork velocities similar to that of BJ and RPE1-hTERT cells, thus excluding the fact that the reduction in replication potency is solely due to alteration in the nucleotide pool (Bester et al., 2011). This is the first demonstration, to our knowledge, that a common property of cancer cells is their weaker capacity to synthesize DNA, likely due to under-licensing of their replication origins.

Palbociclib extends G1 and rescues the low GIFD of HCT116 cancer cells

Many of the most prominent oncogenes and loss of tumor-suppressor genes exert their function by accelerating the G1/S transition of the cell cycle. As shown previously, this can potentially decrease the efficiency at which replication origins are licensed and, consequently, the rate of DNA synthesis (Ekholm-Reed et al., 2004;Lengronne and Schwob, 2002;Tanaka and Diffley, 2002). To test the notion that cancer cell lines have a lower fork density due to a shortened G1, and implicitly incomplete origin licensing, we extended the G1 phase of cancer cells with low doses of the Cdk4/6 inhibitor Palbociclib, which prevents pRb phosphorylation and G1 transit, to see if it rescues their slow S phase. HCT116 cells (Rb positive), HeLa cells (Rb negative as a control) and untransformed RPE1-hTERT cells were treated with 0.16, 0.32 and 1 µM Palbociclib for 24 or 48 hours, and the increase of their G1 fraction analyzed by flow cytometry. As expected, Palbociclib had no effect on Rb-negative HeLa cells (Fig. 4A and Supp Table III). In contrast, Palbociclib at 0.16 µM increased the fraction of G1 cells by ~20% in HCT116 and RPE1 cells without blocking the cell cycle (compare 24 and 48 h), whereas higher doses did so in agreement with a previous report (Lee et al., 2016). To confirm that 0.16 µM Palbociclib prolongs G1 rather than blocking it, we added BrdU to Palbociclib-treated cells and showed that 100% became BrdU positive after 24h (Fig. 4B), demonstrating that they are indeed dividing, yet with an extended G1 phase.

We then measured GIFD by DNA combing in RPE1, HCT116 and HeLa cells treated with DMSO (vehicle) or 0.16µM Palbociclib. This led to a 30-40% increase in fork density in RPE1 and HCT116 cells that displayed an extended G1, but not in HeLa cells that did not due to their Rb-negative status (Fig. 4C).

This demonstrates that increasing the duration of G1 can increase the number of active forks as seen in yeast (Lengronne and Schwob, 2002), likely by improving the licensing of origins in G1. We tried to detect an enhancement of Mcm2-7 loading onto chromatin using a flow cytometry assay (Matson et al., 2017), but failed to do so probably because this biochemical assay is not sensitive enough to reveal slight changes in MCM loading in vivo, as in the case of Cdt1 overexpression which did not lead to more detected MCM on chromatin despite obvious re-replication (Supp Fig. 6). Strikingly, the increased fork density in HCT116 cells treated with Palbociclib also led to a decrease in S-phase duration (Fig. 4D, Supp Fig. 5). This indicates that low doses of Palbociclib can restore a normal S phase to Rb + cancer cells, possibly explaining its therapeutic success in treating various cancers.

Mitotic DNA synthesis and chromosome segregation defects in cancer cells with extended S phase

Previous work has shown that slowing down of replication forks using low doses of aphidicolin, an inhibitor of DNA polymerases, leads to late replication intermediates that are resolved through DNA synthesis in mitosis (MiDAS), detected as EdU foci on condensed chromosomes (Minocherhomji et al., 2015). Similar mitotic EdU foci have been seen on common fragile sites (CFS) that are known to replicate late in the cell cycle and break in mitosis (Bhowmick et al., 2016;Naim et al., 2013). Since we showed above that cancer cells inherently struggle to replicate their genome, even without the addition of aphidicolin or other replication stressors, we sought for signs of MiDAS in exponentially growing U2OS, HCT116 and HeLa cancer cells, compared to RPE1-hTERT untransformed cells. Besides not using aphidicolin or nocodazole, we also shortened the EdU pulse from 30 to 15 min to lower the risk of carrying into mitosis EdU previously incorporated in late S phase. Such MiDAS events likely being rare in the already scarce population of mitotic cells in the cell population, we used a high-content screening (HCS) microscope to automatically detect mitotic cells, using phospho-H3 staining, among hundreds of thousands of cycling cells. These mitotic cells were then screened for EdU foci in confocal images, and confirmed visually. We found that 5.5% of mitotic U2OS cells displayed MiDAS, a 27-fold increase compared to the untransformed RPE1-hTERT cell line, whereas this increase was 5.5-fold and 3.5-fold for HCT116 and HeLa, respectively (Fig. 5A,D). Thus, cancer cell lines have an intrinsic tendency to resolve late replication intermediates or complete DNA synthesis in mitosis. The confocal DAPI images also allowed us to score for chromosome segregation defects, which were increased between 2.7 and 5.4-fold in cancer cells compared to RPE1 cells (Fig. 5B). Defects consisted mainly in a failure for chromosomes to align on the metaphase plate, the presence of lagging chromosomes and anaphase bridges (Fig. 5C,D). Altogether our results provide strong evidence that cancer cells share the new hallmark of enfeebled DNA synthesis capacity, which translates into an extended S phase, mitotic DNA synthesis and segregation defects that fuel chromosome instability.

Discussion

Using single-molecule analysis of DNA replication, which in contrast to 'ensemble' techniques takes into account the stochasticity and plasticity of origin firing, as well as a novel method to measure Sphase duration from asynchronous cell populations, we provide here firm evidence that all the fourteen tested cancer cell lines suffer from a weaker DNA synthesis capacity than untransformed cells. Their longer S phase is mainly due to a decrease in replication fork density, which can be reversed by prolonging the G1 phase using low doses of the FDA-approved cancer drug Palbociclib. We also show that all three tested cancer cell lines have constitutively higher frequencies of mitotic DNA synthesis (MiDAS) and chromosome segregation defects. These novel findings have strong implications for understanding why cancer cells have high genome instability, but also provide new perspectives for the detection and therapy of cancer.

Analyzing chromosome replication dynamics from a single DNA molecule perspective allowed us to propose two new descriptors for how cells duplicate their genome. The global instant fork density (GIFD) is a genome-wide snapshot of how many replication origins have been fired at any moment during S phase in a given cell population. It includes firing events located in distant replication domains and is not affected by DNA fragmentation, in contrast to inter-origin distances measured on single molecules. GIFD is computed per Mb of DNA and is therefore independent of ploidy changes in various cancer cell lines. The second descriptor, replication potency (RP), integrates the average speed of replication forks to the fork density, and therefore reflects how many kb of DNA a cell in S phase can synthesize per minute, for every Mb of its genome. These measurements of GIFD and RP capture the essence of genome replication because they can correctly predict the duration of S phase measured using an orthogonal method, and are consistent with S-phase lengths found in the literature. RPE1 cells, for example, have a GIFD of 1.2 forks/Mb and a FV of 1.6 kb/min. For a genome of 6.6 Gb, this computes as 7,920 forks and 12,672 kb synthesized every min during S phase, and an extrapolated 521 min or 8.7 h to replicate the entire genome that is very close to the measured SPD, although this is not the case for all cell lines (Supp Table V, , Supp Fig. 3). Cancer cell lines having the lowest replication potency are expected to spend the longest time in S phase. Figure 6A shows this correlation between RP and SPD for the fourteen cancer cell lines studied, and validates RP as a correct predictor of S-phase duration. The three untransformed cell lines cluster clearly away from cancer cell lines in this representation (Fig. 6A), which could potentially be used for tumor cell stratification.

The lower replication potency and increased S-phase duration that we uncovered in cancer cells have two additive components, at least: a decreased density of active replication forks (Fig. 3C) and a variable but generally lower fork speed (Sup Fig. 1B). The latter has been described and attributed to lower dNTP pools in primary keratinocytes infected with E6 and E7 oncoproteins (Bester et al., 2011), as well as in cells overexpressing cyclin E or after knock-down of WEE1 (Beck et al., 2012;Jones et al., 2013). Pennycook et al. (2020) have proposed that the E2F-dependent transcription regulates fork speed and S-phase length, without affecting the number of active forks (Pennycook et al., 2020). Our results, in contrast, show that cancer cells utilize fewer replication origins, globally, compared to untransformed cells. The former studies used inter-origin distances as a proxy for origin density, but this a local measure that is highly biased by fiber fragmentation (Jones et al., 2013;Técher et al., 2013). The global instant fork density (GIFD) used here does not suffer the same flaw, integrates origins that are at far distances one of another (different replication domains) and showed a general trend of reduction in cancer cell lines. Combining this fork density to the measure of fork velocity generated a new descriptor of replication potency that is clearly lower in all fourteen tested cancer cell lines.

Macheret and Halazonetis showed that MYC and cyclin E-induced U2OS fire a small subset of new origins that are absent in the relative controls and do not serve as dormant origins. EdU sequencing showed that these origins are located in intragenic regions and can create transcription-replication conflicts. The results confirmed the shortening of G1 in oncogene-induced cells, but the overall density of origins was not measured (Macheret and Halazonetis, 2018). Cyclin E overexpression in mammalian cells was shown to reduce MCM loading (Ekholm-Reed et., 2004), which is consistent with our results. A shortage or imbalance in nucleotide pools has also been invoked as a cause of oncogene-induced replication stress (Bester et al., 2011), but this is debated (Manning et al., 2014). Furthermore, our results on Palbociclib-treated HCT116 cells confirms the importance of G1 duration and origin licensing in maintaining a strong replication potency in S phase. The fact that Palbociclib neither extended G1 nor rescued the long S phase of HeLa cells (Rb-negative) confirms that this drug targets the Rb pathway, as previously shown (Schade et al., 2019).

The question regarding the existence of a G1/S checkpoint delaying S-phase entry upon origin under licensing is debatable. Some studies showed that depletion of licensing factors induce G1 arrest in p53-dependent manner (Shreeram et al., 2002). However, the fact that cancer cells often have defects in the p53 pathway allows them to proceed with S phase even in the absence of licensing factors (Feng et al., 2003;Nevis et al., 2009). On the other hand, downregulation of Mcm2 in untransformed cells allowed them to progress into S phase but led to an increase in the frequency of anaphase bridges suggesting that replication was not efficient (Passerini et al., 2016). Thus, origin under-licensing is a valid cause for explaining the extended S phase and chromosome instability of cancer cells.

Previous reports have analyzed cancer specimens by flow cytometry and used the S-phase fraction as a proliferative index and a predictive parameter of relapse (Clark et al., 1989;Pinto et al., 2001), in addition to others, but without taking into consideration the variability of cell cycle profile among different cancer cells. The growth rate among several cancer cell lines was shown to be significantly variable through the measuring of population doubling time (Bazzocco et al., 2015;Khosravi-Maharlooei et al., 2015). These early attempts to use S phase as a prognostic tool were un successful and emphasize the necessity of assessing S-phase dynamics using novel and more direct techniques, as the ones described here. Our results demonstrate a novel property shared among most cancer cell lines, which is that they spend a longer time in S phase and struggle to replicate their genome.

One consequence of cells spending a longer time in S phase due to reduced origin licensing, as shown here, or after slowing down fork elongation rates with aphidicolin, is mitotic DNA synthesis seen as EdU foci on condensed chromosomes (Bhowmick et al., 2016;Minocherhomji et al., 2015). Here, we extended these observations by assessing the penetrance of MiDAS using high-content screening in asynchronously-growing U2OS, HCT116, HeLa and RPE1-hTERT cells in the absence of aphidicolin treatment. By monitoring 1200-1500 mitotic cells we detected naturally occurring MiDAS events and chromosome segregation defects at 5-25x increased frequency in cancer cells compared to untransformed RPE1-hTERT cells (Fig. 5). Our results underscore the importance of assessing rare events in unperturbed cell populations to grasp the underlying genome instability of normal cells that allows evolution to take place, as well as its inflation in cancer cells that fuels tumor progression and the frequent resistance to cancer therapies. The stochastic nature of origin firing is permissive for rare events of incomplete DNA replication that generate genome instability, and its increased frequency in pre-cancerous cells with deregulated G1 control and origin under-licensing would be sufficient to drive tumorigenesis (Fig. 6 B). It is conceivable, in this regard, to revert genome instability to acceptable levels and to lower cancer burden by restoring a normal S phase to pre-cancerous cells.

Palbociclib is a specific inhibitor of CDK4/6 kinases and of the G1/S transition by preventing Rb phosphorylation. It has been approved by the FDA as a first line treatment of women with advanced estrogen receptor-positive and human epidermal growth factor-2 (HER2)-negative breast cancer combined with hormonal therapy (Finn et al., 2015), and is currently being tested for several other cancer indications. How Palbociclib cures or prevents cancer is not fully understood. At high doses, it blocks the cell cycle in G1 and is thought to affect preferentially cancer cells with higher proliferation rates.

However, it also generates de novo resistance through Rb loss or cyclin E overexpression (Guarducci et al., 2018;Herrera-Abreu et al., 2016). Our results demonstrate that Palbociclib administrated at low doses that do not block the cell cycle, but solely extend G1 duration by 20%, can increase origin licensing and 2.51x10 -6 ) between untransformed and transformed group. .193x10 -6 ) between untransformed and transformed group. D) Replication potency for the same cell lines, which takes into account both the number of forks every Mb (GIFD) and the mean fork speed (FV, see Suppl Fig. 1B). Values are mean (n=3); no statistical significance by Kruskal-Wallis (P=6.218x10 -2 ) between BJ, HMEC-hTERT and RPE1-hTERT; statistical significance by Wilcoxon test (P= 6.57x10 -10 ) between untransformed and transformed group.

Measuring Inter-mitotic Time of Untransformed and Cancer Cell Lines Using SiR-Hoechst Vital Stain

In order to follow intermitotic times of untransformed and cancer cell lines, cell lines were grown on microscopy dishes and were imaged every 10 min for 72h on a widefield inverted microscope, either by phase contrast or by fluorescence using the vital DNA stain SiR-Hoechst at 100 nM. However, using the given microscope and the microscopy conditions we used, we have noticed that even at lower

Assessing the Replication Potency as a Factor of Measuring Replication Strength Despite Differences in the Genome Size

Cancer cell lines are known for having cytogenetic and genomic abnormalities (supp. table I.). In order to assess whether our measurement of the replication potency is not biased by the variation of the DNA content between cancer and untransformed cell lines, we have designed a new approach to address this question. We speculated that if cells with more DNA content, they will have more DAPI signal compared to diploid cells when stained with a saturated concentration of DAPI. Thus, an equal number of cells from PC-3, HCC-1937, BXPC-3, HCC-38, 22RV1, MKN-45 and RPE1-hTERT were stained with increasing concentrations of DAPI then washed. Cells were then analyzed by FACS and the difference in the DNA amount was calculated as a ratio of G1 peak signal in the cancer cell lines versus G1 peak signal of the untransformed diploid RPE1-hTERT (Figure 16. and supp. Table II). Knowing the genome size per cell line and the replication potency would allow us to determine the DNA synthesis rate per min as a factor of genome size and replication potency. As a result, SPD can be extrapolated based on combing data by dividing the genome size per the time of DNA synthesis per min. However, if both the DNA synthesis rate and the extrapolated SPD are dependent on genome size, the genome size factor is annihilated leaving behind the replication potency as the only factor to calculate the extrapolated SPD (Suppl. Table III). This further enhances the strength of replication potency as a factor to evaluate DNA replication independent of the variation in genome size among different cell lines. However, in order to confirm that the decrease in the replication potency is mainly attributed to origin underlicensing, origin licensing was addressed by trying to increase origin licensing efficiency through G1 prolongation. RPE1-hTERT

MKN

Project 2: Replication Dynamics in Pluripotent Stem Cells

ESCs are characterized by having a peculiar cell cycle where they spend less time in G1 and G2 phases compared to somatic cells. Having short G1 was shown to be important for maintaining pluripotency. Altering G1 kinetics induces G1 prolongation which is accompanied with cellular differentiation (Coronado et al., 2013a). Despite having short G1 efficient DNA licensing in ESC should be attributed to fast MCM loading and increased expression of CDT1 and CDC6 (Matson et al., 2017). Furthermore, cell fate transition and differentiation domains of

ESCs was shown to be less compartmentalized and characterized by switching of replication timing compared to constitutive domains. This suggested that regulation of replication timing might be involved in cellular differentiation (Dileep et al., 2015a). However, the link between cell cycle remodelling, DNA replication regulation and development needs further investigations. The main aim of this project is to get more insights about the regulation of replication dynamics in ESCs relative to their differentiated state.

2.1.The main results of this project

1) Single-molecule analysis by DNA combing was used to test whether replication dynamics contribute to stem cells pluripotency. The results showed little change in the fork velocities and local inter-origin distances (IODs) between mouse ESCs and their differentiated counter parts MEFs.

2) Measuring the origins density globally in both cell types revealed that MEFs use 2-fold more origins compared to the mouse ESCs.

3) As a result of using more number of origins, MEFs are expected to spend less time in S phase compared to the mouse ESCs. S-phase duration measurements revealed that both cell types spend almost 8 hours to replicate the genome.

4) Replication parameters measured by DNA combing fit very well the S phase duration measured. However, MEFs use 2-fold more origins than required to replicate the genome with the given 8 hours of S phase.

My contribution to the project

Based on the above data, three main hypotheses were suggested to explain the increase in fork density in MEFs.

1) Replication is continuous in mouse ESCs while it is interrupted in MEFs due to pausing events. Fork pausing in MEFs would allow dormant origins to fire which would explain the increase in fork density in MEFs. Thus, pausing events have been tested in both

MEFs and mouse ESCs using modified DNA combing protocol.

2) mESC display more even distribution of origins on DNA while origins are more clustered in MEFs. Using our DNA combing protocol IOD was used as aparameter to study origins fired within the same cluster. Considering that origins are more clustered in MEFs, the IOD that was previously measured in MEFs might not include the distance between origins of different clusters. This might explain the absence of significant difference in IODs between MEFs and mouse ESC. In order to test this hypothesis, two approaches have been used. The first approach is through optimizing DNA combing protocol through increasing fibres length which might allow the measurement of IOD between clusters.

The second is through decreasing the compaction of chromatin in MEFs which might allow more even distribution of origins. Maintaining even origins distribution in MEFs is expected to reduce the number of origins used by MEFs to replicate the genome.

3) Another possibility behind the increase in GIFD in MEFs might be attributed to difference in the distribution of origins used in early, mid and late S phase. Having more origins in early S phase while fewer origins in late S phase would also allow the firing of dormant origins if they were licensed. If not this would require the cells to spend longer time to replicate the late replicating regions despite earlier increase in the number of origins used. In order to test this hypothesis, MEFs have been sorted into early, mid and late S phase cells based on their DNA content profile. GIFD was determined and compared in each of the mentioned fractions.

INTRODUCTION

Pluripotent stem cells (PSC) such as embryonic stem cells (ESC) or induced pluripotent stem cells (iPSC) are renowned for their ability to self-renew in culture and their capacity to differentiate into cells of three germ layers giving rise to all cells of the developing embryo. During senescence or terminal differentiation most cells exit from the cell cycle, which allows them to avoid tumorigenesis. ESCs and iPSCs in contrast can proliferate indefinitely in culture, yet be driven to differentiate into specific cell types, which explains their huge interest for developmental biology, disease modelling and regenerative medicine. One key question is how stem cells establish and maintain their pluripotent epigenome. This clearly involves an intertwined network of stem cell specific transcription factors [START_REF] Chambers | The transcriptional foundation of pluripotency[END_REF][START_REF] Morey | Pluripotency and Epigenetic Factors in Mouse Embryonic Stem Cell Fate Regulation[END_REF], peculiar cell cycle regulation [START_REF] Boward | Concise Review: Control of Cell Fate Through Cell Cycle and Pluripotency Networks[END_REF][START_REF] Fluckiger | Cell cycle features of primate embryonic stem cells[END_REF][START_REF] Stead | Pluripotent cell division cycles are driven by ectopic Cdk2, cyclin A/E and E2F activities[END_REF]White and Dalton, 2005), as well as hyper-dynamic chromatin architecture [START_REF] Dixon | Chromatin architecture reorganization during stem cell differentiation[END_REF][START_REF] Mattout | Heterochromatin Protein 1β (HP1β) has distinct functions and distinct nuclear distribution in pluripotent versus differentiated cells[END_REF]Meshorer and Misteli, 2006;Meshorer et al., 2006).

Interestingly ESCs and iPSCs proliferate more rapidly than somatic cells (~12 h vs ~24 h doubling time)

and have a unique cell cycle structure, with shortened G1 and G2 phases and S phase representing around 60% of the total cell cycle, a trend that is even more evident during early embryogenesis in fly, frog and fish [START_REF] Boward | Concise Review: Control of Cell Fate Through Cell Cycle and Pluripotency Networks[END_REF]. A short G1 seems particularly important for naïve pluripotency since G1 lengthening was shown not only to correlate with ESC progression to a primed state but also to induce their differentiation [START_REF] Calder | Lengthened G1 phase indicates differentiation status in human embryonic stem cells[END_REF]Coronado et al., 2013b), although dissonant reports exist [START_REF] Li | Gap 1 phase length and mouse embryonic stem cell self-renewal[END_REF]. The cell cycle of ESCs is peculiar by having a constitutive high cyclin-dependent kinase (CDK) activity, which is brought about by the altered expression, compared to somatic cells, of many regulators including i) absence of the Rb-dependent restriction point in late G1 (White et al., 2005), ii) high levels of cyclin E-Cdk2 and cyclin A-Cdk2 activity and their positive regulator Cdc25A (van der [START_REF] Van Der Laan | High Dub3 expression in mouse ESCs couples the G1/S checkpoint to pluripotency[END_REF][START_REF] Stead | Pluripotent cell division cycles are driven by ectopic Cdk2, cyclin A/E and E2F activities[END_REF] iii) high Emi1 levels and low APC-Cdh1 activity (Ballabeni et al., 2011b), iv) lack of p21 and p27 Cdk inhibitors (Fujii-Yamamoto et al., 2005b;[START_REF] Neganova | Expression and functional analysis of G1 to S regulatory components reveals an important role for CDK2 in cell cycle regulation in human embryonic stem cells[END_REF][START_REF] Zhu | JMJD5 regulates cell cycle and pluripotency in human embryonic stem cells[END_REF]. All these factors contribute to ESCs progressing rapidly through G1, which is reinforced in a positive feedback loop by high levels of the Oct4, Sox2 and Nanog pluripotency genes [START_REF] Boward | Concise Review: Control of Cell Fate Through Cell Cycle and Pluripotency Networks[END_REF]. A key unresolved question is how this remodelled cell cycle architecture can impact on the establishment or maintenance of a pluripotent chromatin and undifferentiated genome. Furthermore how DNA replication can take place in ESCs without marked CDK inactivation in G1 is particularly intriguing.

DNA replication in all eukaryotes entails first the selection, from late mitosis to late G1, of numerous chromosomal sites (~150,000 in mammals) that can later be used as replication origins (Besnard et al., 2012b;[START_REF] Cayrou | Genome-scale analysis of metazoan replication origins reveals their organization in specific but flexible sites defined by conserved features[END_REF][START_REF] Cayrou | The chromatin environment shapes DNA replication origin organization and defines origin classes[END_REF][START_REF] Miotto | Selectivity of ORC binding sites and the relation to replication timing, fragile sites, and deletions in cancers[END_REF][START_REF] Petryk | Replication landscape of the human genome[END_REF]. In this process called origin licensing, the Orc1-6, Cdc6, Cdt1 and Mcm2-7 pre-replication complex (pre-RC) proteins sequentially bind to these sites, making them competent for initiation of bi-directional DNA synthesis. This step is antagonized by Cdk activity and thus takes place during G1 when cyclin proteolysis and Cdk inhibitors keep this activity at a low level (Diffley et al., 1994a;[START_REF] Siddiqui | Regulating DNA replication in eukarya[END_REF]. Loss of Cdk inhibitors or overexpression of G1/S cyclins interferes with pre-RC assembly, extends S phase and causes genomic instability in yeast (Lengronne and Schwob, 2002b;Tanaka and Diffley, 2002b) and somatic cells (Ekholm-Reed et al., 2004b), but apparently not in ESC (Ballabeni et al., 2011b;Fujii-Yamamoto et al., 2005b). Efficient replication licensing in ESC, despite high constitutive Cdk activity, may be due to the elevated levels of Cdc6 and Cdt1 licensing factors. Later only a subset of these competent origins (~15,000) is activated during S phase, when raising CDK and Dbf4-dependent kinase (DDK) activities recruit further proteins into a Cdc45-MCM-GINS (CMG) complex capable of unwinding the DNA duplex and triggering the initiation of DNA replication [START_REF] Tanaka | Helicase activation and establishment of replication forks at chromosomal origins of replication[END_REF]. All origins do not fire simultaneously at the beginning of S, but sequentially throughout S phase in cohorts of replication foci of various size and nuclear location that are activated following a stereotyped spatio-temporal replication pattern delineating the distinct states of euchromatin, facultative and constitutive heterochromatin [START_REF] Chagin | 4D Visualization of replication foci in mammalian cells corresponding to individual replicons[END_REF][START_REF] Dimitrova | The spatial position and replication timing of chromosomal domains are both established in early G1 phase[END_REF]Ma et al., 1998b;Nakamura et al., 1986b). The position, timing and efficiency of origin firing is influenced by limiting initiation factors and titration effects [START_REF] Collart | Titration of four replication factors is essential for the Xenopus laevis midblastula transition[END_REF][START_REF] Mantiero | Limiting replication initiation factors execute the temporal programme of origin firing in budding yeast[END_REF][START_REF] Tanaka | Multiple Regulatory Mechanisms to Inhibit Untimely Initiation of DNA Replication Are Important for Stable Genome Maintenance[END_REF][START_REF] Yoshida | The histone deacetylases sir2 and rpd3 act on ribosomal DNA to control the replication program in budding yeast[END_REF] as well as by histone modifications regulating chromatin accessibility [START_REF] Aparicio | The Rpd3-Sin3 histone deacetylase regulates replication timing and enables intra-S origin control in Saccharomyces cerevisiae[END_REF][START_REF] Pasero | Single-molecule analysis reveals clustering and epigenetic regulation of replication origins at the yeast rDNA locus[END_REF], DNA damage checkpoint responses [START_REF] Guo | Interaction of Chk1 with Treslin negatively regulates the initiation of chromosomal DNA replication[END_REF][START_REF] Zegerman | Checkpoint-dependent inhibition of DNA replication initiation by Sld3 and Dbf4 phosphorylation[END_REF], local inhibition [START_REF] Cornacchia | Mouse Rif1 is a key regulator of the replication-timing programme in mammalian cells[END_REF][START_REF] Davé | Protein phosphatase 1 recruitment by Rif1 regulates DNA replication origin firing by counteracting DDK activity[END_REF][START_REF] Hiraga | Rif1 controls DNA replication by directing Protein Phosphatase 1 to reverse Cdc7-mediated phosphorylation of the MCM complex[END_REF]Yamazaki et al., 2012b) and nuclear localization [START_REF] Knott | Forkhead transcription factors establish origin timing and long-range clustering in S. cerevisiae[END_REF] (see (Fragkos et al., 2015b;[START_REF] Marks | Replication origins: determinants or consequences of nuclear organization?[END_REF] for recent reviews). Single DNA molecule analysis revealed that replication initiation sites are chosen stochastically among a vast excess of potential origins (Lebofsky et al., 2006b;[START_REF] Patel | DNA replication origins fire stochastically in fission yeast[END_REF]. At the population level however these stochastic initiation events coalesce into larger replication domains showing reproducible timing [START_REF] Bechhoefer | Replication timing and its emergence from stochastic processes[END_REF][START_REF] Rhind | DNA replication timing[END_REF]. Interestingly these replication domains correspond to the topologicallyassociated domains (TAD) that have been described by chromosome conformation capture (3C) methods, and represent stable structures of genome organization [START_REF] Pope | Topologically associating domains are stable units of replication-timing regulation[END_REF]. However up to 45% of these replication domains have been shown to switch their timing during the differentiation of ESCs towards various lineages, suggesting that changes in replication timing accompany or may dictate chromosome reorganization and gene expression patterns in different cell types (Dileep et al., 2015b;[START_REF] Dixon | Chromatin architecture reorganization during stem cell differentiation[END_REF][START_REF] Hansen | Sequencing newly replicated DNA reveals widespread plasticity in human replication timing[END_REF][START_REF] Hiratani | Global reorganization of replication domains during embryonic stem cell differentiation[END_REF][START_REF] Hiratani | Genome-wide dynamics of replication timing revealed by in vitro models of mouse embryogenesis[END_REF][START_REF] Rivera-Mulia | Dynamic changes in replication timing and gene expression during lineage specification of human pluripotent stem cells[END_REF][START_REF] Ryba | Evolutionarily conserved replication timing profiles predict long-range chromatin interactions and distinguish closely related cell types[END_REF].

However despite intensive studies, it is not yet clear what makes developmentally regulated replication domains switch their timing [START_REF] Rivera-Mulia | Replication timing and transcriptional control: beyond cause and effect-part III[END_REF].

In order to gain new insights into the links between cell cycle control, S phase and the pluripotent genome of ESCs, we compared the dynamics of DNA replication in ESCs and differentiated cells using the high-resolution and quantitative method of DNA combing [START_REF] Bensimon | Alignment and sensitive detection of DNA by a moving interface[END_REF][START_REF] Michalet | Dynamic molecular combing: stretching the whole human genome for high-resolution studies[END_REF], whereby genomic DNA from exponentially-growing cells pulse-labelled with thymidine analogues is stretched on glass surfaces for subsequent detection of DNA synthesis sites along individual DNA molecules (Bialic et al., 2015b;[START_REF] Huberman | Autoradiography of chromosomal DNA fibers from Chinese hamster cells[END_REF]Lengronne et al., 2001b). Crucially this single molecule approach is not confounded by the population averaging produced by ensemble methods that sum up all stochastic events in the cell population. We define here a new indicator of cellular replication activity, the Global Instant Fork Density (GIFD), which scores the average number of replication forks that are active at any time during S phase in each cell. We show that this GIFD is 2-fold lower in mouse and human pluripotent stem cells relative to their differentiated counterparts. Using a newly developed method to determine S-phase duration from asynchronous cultures, we find that S-phase length is unchanged (~8.5 h) in mESC and MEF. Our quantitative data indicate that differentiated cells use twice more replication origins than stem cells to duplicate their genome, mainly because their replicons are shorter due to intrinsic constraints on fork progression compensated by the activation of dormant origins. We propose that DNA replication is more processive in pluripotent stem cells whereas it is more temporally segmented in differentiated cells, and speculate that these different modes of DNA synthesis may underlie the establishment of their distinct chromatin states.

RESULTS

Cell cycle architecture in pluripotent stem cells and during their differentiation

Pluripotent stem cells are known to proliferate rapidly and have a truncated G1 phase. In order to more precisely determine their cell cycle structure and S-phase fraction, we performed bivariate BrdU/DNA FACS analyses on various pluripotent stem cells from mouse or human origin as well as on their differentiated cell counterparts. The stemness of our ESC and iPSC cultures was verified by Oct4, Nanog and Klf4 expression using RT-qPCR and by alkaline phosphatase staining (not shown). Sites of DNA synthesis in exponentially growing cells were labelled with consecutive pulses of iododeoxyuridine (IdU, 15 min) and chloro-deoxyuridine (CldU, 15 min) and processed for FACS using an anti-BrdU antibody that also recognizes IdU. Figure 1 shows that all PSCs tested had fewer cells in G1 and a much higher fraction of cells in S phase than their differentiated counterparts. The difference was most striking when mESCs (10% G1, 76% S) were compared to MEFs (54% G1, 27% S), but was still evident for hESCs and hiPSCs (~35% G1, ~50% S) compared to BJ fibroblasts (60% G1, 18% S). This difference between mESC and hESC may be related with the latter being in a less naïve state of pluripotency [START_REF] Tesar | New cell lines from mouse epiblast share defining features with human embryonic stem cells[END_REF].

We also differentiated mESC into neural precursor cells (NPC) and followed the restructuration of their cell cycle along their lineage commitment [START_REF] Bibel | Differentiation of mouse embryonic stem cells into a defined neuronal lineage[END_REF]. The fraction of S phase cells steadily decreased and the fraction of G1 cells steadily increased during the nine days of differentiation, while the fraction of G2/M cells remained low without varying much (Fig. 1C,D). Thus the accumulation of ESC in S phase does not induce a G2/M delay, suggesting that ESC either experience little or no DNA damage, that this damage is invisible to the G2/M checkpoint or that it is dealt with during S phase (Ahuja et al., 2016a). After nine days of differentiation (EB9) cells adopted the cell cycle architecture typical of differentiated cells, with a longer doubling time (not shown) and extended G1 phase. Interestingly this state was preceded at EB3 and EB6 by a 3-fold increase in the level of gH2AX measured by quantitative western blotting, suggesting that a moderate replication stress might come with the earliest steps of lineage commitment (Fig. 1E).

Fork velocities and local inter-origin distances vary little in ESC and MEF

Next we used DNA combing to determine several parameters pertaining to the dynamics of DNA replication on the cellular level. DNA combing can stretch and align DNA molecules in the mega base pair range (Mb) for single-molecule detection by fluorescence microscopy. This approach, in contrast to ensemble methods, is quantitative and monitors replication fork progression and origin densities on individual DNA molecules, taking into account the high cell-to-cell variability and stochastic nature of DNA replication.

Pluripotent and differentiated cells were labelled with two consecutive short pulses of IdU and CldU (15 min each) then chased 1-2 h with thymidine (dT) before fixation and harvesting cells. The two pulses allow determining fork direction whereas the dT chase moves the replication forks, which tend to break during DNA combing, away from the labelled tracks. The concentration of nucleosides and duration of labelling were chosen not to interfere with cell cycle progression. After detection of IdU, CldU and DNA with specific antibodies, we selected for fork velocity (FV) measurements only unambiguous bicolour red-green tracks (inside a DNA fibre) corresponding to elongating forks, thus excluding monocolour and tricolour signals that correspond to stalled or initiating/terminating forks, respectively (Fig. 2A). Because DNA combing stretches DNA uniformly (2kb/µm), we can derive FV by dividing CldU track length (in kb) by duration of the pulse. We found that individual fork rates vary significantly (3-5-fold) and that each cell type studied had a specific mean FV: 1.72 kb/min in mESC, 2.0 kb/min in MEF, 1.01 kb/min in hESC, 1.15 kb/min in hiPSC, and 1.61 kb in BJ human fibroblasts. These differences likely stem from the specific growth conditions for each cell type (medium, oxygen level) and/or their intrinsic metabolic differences (e.g. dNTP pools) but overall, replication forks tend to move more slowly in stem cells compared to their differentiated cell counterparts (Fig. 2B). This lower FV in PSC may be related to the high level of Rad51 protein and homology directed repair that was shown to slow replication forks in mESC (Ahuja et al., 2016a;[START_REF] Yoon | Rad51 regulates cell cycle progression by preserving G2/M transition in mouse embryonic stem cells[END_REF].

Besides generally moving more slowly, replication forks can also slow down or pause locally due to intrinsic or extrinsic impediments to fork progression, which can sometimes be detected by asymmetries in the distance covered by left-and right-moving sister forks. Furthermore fork slowing or stalling can lead to the activation of dormant origins located nearby, which can be detected by a decrease in local inter-origin distances (IOD). We found that sister fork asymmetry is slightly bigger (Fig. 2C) and IODs slightly shorter (Fig. 2D) in MEF compared to mES cells. Although these differences are not statistically significant by Mann-Whitney test at this depth of analysis, taken together they suggest that the kinetics of chromosome replication are slightly different in mESC and MEF.

Global instant fork density is lower in mouse and human pluripotent stem cells

Inter-origin distances measured by DNA combing only relate to the density of origins locally, on the same DNA fibre, within replication domains containing clustered origins that fire synchronously.

However only a fraction of the genome contains origins clustered at short (50-200 kb) distance [START_REF] Berezney | Heterogeneity of eukaryotic replicons, replicon clusters, and replication foci[END_REF], and IOD measurements are very biased by the limited size of the DNA fibres that can be visualized by DNA combing (Técher et al., 2013b;[START_REF] Tuduri | Defining replication origin efficiency using DNA fiber assays[END_REF]. To circumvent these caveats and get a more precise idea of how many origins fire globally instead of locally, we defined a new parameter called global instant fork density (GIFD) that is calculated by dividing the total number of moving replication forks (IdU-CldU bicolour signals) detected after pulse labelling by the total length of single DNA molecules measured in the same set of images (Fig. 2A). GIFD is expressed in forks/Mb, it is global and not restricted to clustered origins, instantaneous as defined by the transition between the red and green track, independent of DNA fibre fragmentation and covers all temporal domains of S phase.

However there are two constraints for GIFD to accurately represent the replication activity of the entire cell population: first, images have to be acquired without bias for those containing labelled fibres; second, the measured density ought to be corrected for the fraction of cells that are not in S phase at the time of the pulse. The former is done by random selection of fibres in the DNA (blue) channel, while the latter is achieved by dividing the measured GIFD by the fraction of cells that are in S phase (measured by BrdU/DAPI FACS on the same samples). We validated this correction method by FACS sorting of S-phase cells (Suppl Fig. S1A,B), but post-sorting DNA combing turned out to be inconvenient due to fluorescence-induced fragmentation of DNA fibres.

GIFD was measured for mESC and MEF on the same samples used above for FV and IOD analyses.

Interestingly, fork density was found to be two-fold lower in mESC (0.98 ±0.18 Fork/Mb) than in MEF (1.93 ±0.12 fork/Mb). This was obtained from several biological replicas and after analysing hundreds of individual DNA molecules totalling 400-900 Mb, which is a sizable fraction of the whole genome (Fig. 2F). The low GIFD of mESC is not a consequence of the normalization used to correct for the fraction of cells in S phase in the population, as a similar low GIFD was found after FACS sorting of a pure S-phase population (Suppl. Fig. S1A,B). It is also not a consequence of selecting only IdU + CldU containing forks, since the difference remained when mono-colour signals were also taken into account (Suppl. Fig. S1C). Strikingly a correspondingly low GIFD was calculated for human ESC and iPSC compared to BJ fibroblasts, indicating that a low GIFD is a conserved feature of pluripotent stem cells (Fig. 2F). We conclude that stem cells use fewer concomitantly active replication forks, and likely origins, at any time during S phase to replicate their genome. This is the largest difference in DNA replication kinetics ever observed between stem cells and differentiated cells, and thus establish GIFD as an interesting new functional marker of cellular replication.

S-phase duration is identical in mESC and MEF

All replication parameters that we determined on single DNA molecules, slower FV, higher IOD and 2-fold lower GIFD, point to a generally slower DNA replication in mESC compared to MEF. An expected consequence of this is that S phase should last longer in mESC than in MEF. Although previous reports suggested, using indirect methods, that S phase is not fundamentally different or longer in mESC relative to MEF [START_REF] Li | Gap 1 phase length and mouse embryonic stem cell self-renewal[END_REF][START_REF] Panning | Spatio-temporal organization of DNA replication in murine embryonic stem, primary, and immortalized cells[END_REF][START_REF] Stead | Pluripotent cell division cycles are driven by ectopic Cdk2, cyclin A/E and E2F activities[END_REF], we thought important to address this issue more thoroughly.

We therefore devised a new protocol to measure S-phase duration in asynchronous cells, which is based on two short pulses with ethynyl-deoxyuridine (EdU) and BrdU separated by thymidine chases of increasing time. The logic is as follows: without intervening chase, almost all cells in S phase during the first pulse will be labelled with both analogues, hence appear as double-positive cells. By increasing the length of thymidine chase between the two pulses, cells initially labelled with EdU will gradually become unable to incorporate BrdU as they exit S phase. Thus the fraction of EdU + BrdU + cells among EdU + cells will decrease up to a point, which defines S-phase duration (SPD), when the earliest cells in S phase will have completed DNA synthesis (Fig. 3A). The conditions for double pulse-chase and analogues detection were carefully chosen so to allow sufficient incorporation yet chase of each analogue, to avoid their cross-detection during FACS analysis and to remain innocuous for cell proliferation (Bialic, Coulon et al, submitted).

Using this method with exponentially growing mESC and MEF cells, we found that the fraction of double-positive cells decreased linearly with increasing chase times in both cultures, thus validating the approach. The intersection of the regression line with the x-axis showed that S-phase duration was identical (~8.5 h) for mESC and MEF in several biological replicates, a value consistent with estimates made by others. Thus the slower kinetic parameters of DNA replication measured for mESC by DNA combing do not seem to impact on the total length of S phase, which raises the question of how can S phase be the same duration in mESC with a two-fold lower GIFD and slightly reduced FV? We believe the issue is not technical since both methods used to reach our conclusions, GIFD by DNA combing and SPD by EdU-BrdU FACS, are robust, reproducible and quantitative. A novel interpretation related to different modes of S-phase progression is proposed below.

Continuous vs interrupted fork progression in mESC and MEF

Since the parameters we measured are quantitative and were obtained using large sampling and stringent conditions, we felt confident to extrapolate our data to the entire mouse genome. DNA combing in mESCs showed that there are 0.98 active forks per Mb of DNA on average in every S-phase cell during the time of the pulse, which corresponds to 5,488 active forks for the whole diploid mouse genome (2 x 2800 Mb). At the measured average FV of 1.72 kb/min, these 5,488 forks will synthesize 9.44 Mb/min (1.72 x 5,488 / 1000). At this rate, 593 min (9h50) would be required to replicate the entire genome. This time is close, yet longer than the S-phase duration we measured for mESC (8.5 h). The difference might be due, we believe, to an underestimation of the number of forks. Indeed if we also consider mono-colour signals, which likely represent stalled forks and account for 33% of all forks (Suppl. Fig. S1C), and arbitrarily set a FV of 1.24 kb/min for them, then the time required to replicate the whole genome would be 8.5 h (Table I). We conclude that the replication parameters we measured by DNA combing fit rather well, and are sufficient to explain the duration of S phase in mESC.

MEF cells in contrast, with 1.93 active forks per Mb of DNA (10,808 at any time during S phase on average) and a FV of 2.0 kb/min, would be able to synthesize 21.62 Mb every minute or 11,026 Mb of DNA during the 8.5h (510 min) of S phase. This is about twice the size of the mouse genome. Thus there is a ~2-fold excess of forks (and origins) in MEF relative to what would be required to replicate the whole genome during S phase. One explanation would be that forks are not active all the time during S phase (temporal interruption model), with periods of high and low replication activity. We can rule out this hypothesis because our GIFD measurements are done on asynchronous cells and cover all periods of S phase, thus averaging the GIFD for any putative periods of high and low replication activity. Another hypothesis is that fork progression is constrained in MEF with widespread fork pausing events (local interruption model), leading to the activation of intervening dormant origins. A simple modelling of this scenario indeed predicts the two-fold increase in fork density that we measured for MEF (Fig. 4A). Its main consequence would be a decrease in replicon size, something that at first sight should have translated into a ~2-fold lower IOD in MEF compared to mESC. However this is not true if the intervening dormant origins fire with some delay (>30 min) after the forks originating from the primary origins have stalled. If so, the firing of primary and of dormant origins cannot be detected during the same 30 min pulse, and IOD will remain mostly unchanged (Fig. 4A bottom).

To test this model and to get some insight into the distribution of active origins and replication fork progression at the 0.5-1 Mb scale of TADs, we optimized our DNA combing approach to analyse labelled DNA fibres that are larger than 500 kb. For that we used CldU pulses of increasing duration (60 and 120 min) to see how far replication tracts progress and how fast such large domains become fully replicated.

Indeed our model predicts that replication tracts should enlarge faster in mESC due to unhindered fork progression, and that gap size between replication tracts should decrease faster in MEF due to the firing of intervening origins. One difficulty of this approach using long pulses resides in the dazzling heterogeneity of FV and IODs, which can rapidly lead to the merging of neighbouring forks. To reject at least in part such fork fusion events, we limited the size of replication tracts used for this analysis to twice the FV times the pulse duration (i.e. 240 kb for 60 min, 480 kb for 120 min). We found that on average replication tracts increase faster (between 60 and 120 min) in mESC (+62%) than in MEF (+29%), confirming the first prediction of our model proposing that fork progression is constrained in MEF cells (Fig. 4B). Conversely our data show that gap size generally decreases faster in MEF (-56%) than in mESC (-5%) during the same time interval (Fig. 4C). Thus both predictions of our model hold true. The proposal that MEF cells complete domain replication faster than mESC was confirmed in another set of experiments in which we scored the extent of replication of each fibre larger than 300 kb during increasing CldU pulse times (30 min to 180 min). This analysis revealed a trend for large fibres to get more fully replicated after 120 and 180 min of labelling in MEF than in mESC, consistent with the recruitment of additional origins during this time window in MEF cells (Suppl. Fig. S2A,B). We conclude that chromosome replication dynamics is globally different in mESC and MEF, the latter being likely subjected to widespread fork pausing events that trigger the firing of dormant origins, which is probably the main cause for the increased fork density we detected. Thus genome replication can be considered more processive in embryonic stem cells, and more discontinuous or interrupted in differentiated cells, a concept consistent with the rather uniform euchromatin in stem cells and the segmented eu-and heterochromatin in lineage-committed cells.

take place [START_REF] Hiratani | Genome-wide dynamics of replication timing revealed by in vitro models of mouse embryogenesis[END_REF]. Our data suggest that cell differentiation induces a complete reorganization of chromosome replication programmes, with multiple and dynamic changes detected by DNA combing.

GIFD increases during in vitro passaging of fibroblasts

To see whether a low replication fork density is specific to mESC and possibly a marker of pluripotency, we thought to measure the GIFD of immortalized mouse NIH 3T3 fibroblasts and of primary MEFs at different passages after their derivation. To our surprise, both NIH 3T3 cells and early passage MEFs (p1, p3) displayed a GIFD of around 1.0-1.2 forks/Mb, very close to the GIFD of mESC and ~2-fold lower than the MEF of passage 4-5 that are routinely used in the laboratory (Fig. 6A,B). Thus a low GIFD is neither specific to mESC nor a marker of pluripotency. Strikingly the GIFD increased steadily with the number of MEF passages (Fig. 6B), concomitant with the decrease in S-phase fraction of these cells (Fig. 6D). Despite this cell cycle restructuration, the duration of S phase measured by the technique described above remained almost constant (~8 h) in MEF of passages p3 to p9 (Fig. 6E). Thus the decrease in S-phase cells during in vitro MEF culture is not due to a shortening of S phase, but to cell cycle lengthening. It has been shown that the CDK inhibitors p16 INK4a and p21 Cip1 , as well as the p53 activator p19 ARF all accumulate gradually during in vitro culture of primary MEF or normal human fibroblasts, causing a progressive G1 lengthening before senescence cell cycle arrest [START_REF] Alcorta | Involvement of the cyclin-dependent kinase inhibitor p16 (INK4a) in replicative senescence of normal human fibroblasts[END_REF][START_REF] Palmero | Accumulation of p16INK4a in mouse fibroblasts as a function of replicative senescence and not of retinoblastoma gene status[END_REF][START_REF] Zindy | Expression of the p16INK4a tumor suppressor versus other INK4 family members during mouse development and aging[END_REF][START_REF] Zindy | Myc signaling via the ARF tumor suppressor regulates p53-dependent apoptosis and immortalization[END_REF]. By analogy with the GIFD increase in MEF versus mESC discussed above, we interpret the gradual GIFD increase during in vitro cell culture of primary MEF by an increased recruitment of dormant origins, which is potentially triggered by increasing replication fork stalling events occurring during early senescence. Crucially the growing use of dormant origins allows these pre-senescent cells to maintain their S phase to a normal length of about 8h (Fig. 6E). Immortalized NIH 3T3 cells in contrast, as well as several cancer cell line tested, have a low GIFD suggesting that they cannot recruit as many additional origins and, as a consequence, are compelled to increase the length of their S phase to ~11 h or more (Fig. 6A,C and data not shown). These data indicate that early-or latepassage primary cells and immortalized cells all have different spatio-temporal replication programmes, and establish the increasing GIFD as an interesting quantitative marker of early senescence.

Altogether our results illustrate the remarkable plasticity of S phase, both in its internal structure and in its duration, which quickly adapts to the changing physiology of cells and determines their fate.

DISCUSSION

We provide here the first in-depth analysis, at the single-molecule level, of the dynamics of chromosome replication in mouse and human ESC, using DNA combing of long DNA fibres and 3colour detection. Counterstaining the DNA fibre, in addition to detecting IdU and CldU, is essential to check the continuity of DNA molecules, to exclude DNA bundles and to correctly assess fork velocity and inter-origin distances (Técher et al., 2013b;[START_REF] Tuduri | Defining replication origin efficiency using DNA fiber assays[END_REF]. Furthermore the capacity to measure the entire length of hundreds of long DNA molecules was absolutely key to define the global instant fork density (GIFD) as a new DNA replication marker, which assesses the number of replication forks (and origins) that are necessary to replicate the genome fully during the imparted time of S phase (Bialic et al., 2015b). The data presented here strongly suggest that an increased GIFD identifies cells and cell types that are resorted to use dormant origins to complete the replication of their genome. We also designed a non-invasive method to measure S-phase duration (SPD) from asynchronous cell populations, which when coupled to GIFD is very informative on how mammalian cells adjust their spatio-temporal chromosome replication programmes during lineage commitment, pre-senescence or after immortalization. Using these quantitative methods we discovered that pluripotent stem cells (mESC, hESC and iPSC) have a lower GIFD than their differentiated counterparts, and likely have replication forks that progress more steadily through chromosomal domains, without the need to activate many dormant origins. The S phase of differentiated and pre-senescent cells on the contrary may be characterized by more frequent fork stalling events, which simply leave enough time for the neighbouring dormant origins to get activated.

Cell cycle structure and other peculiarities of pluripotent stem cells

Early embryonic development in all metazoans is characterized by rapid cell division cycles and the absence (or truncation) of gap phases between DNA synthesis and mitosis. Although less dramatic than the reduction of cycle length in fly or frog early embryos, the doubling time of mESC is about half (10-12 h) that of MEF (see Suppl. movies M1 and M2). Importantly we show here using a direct method that S phase lasts ~8.5 h in both cell types (Fig. 3), indicating that the cell cycle shortening is mostly at the expense of G1, which lasts only ~2 h in mESC compared to ≥12 h in MEF and lacks Rb-dependent regulation of E2F activity (White et al., 2005). Thus mESC cells spend most of their time (~70%) replicating their chromosomes, not because S phase is longer but because the cell cycle is much shorter. This is also true albeit to a lesser extent for human ESC and iPSC (Fig. 1A,B). The property of cells to keep the duration of S phase quasi constant despite important changes in cell cycle structure or nutrient availability is truly remarkable. Fly and frog embryos, in contrast, have a much shorter S phase pre-MBT than post-MBT. This is due in part to the synchronous, unordered replication of all sequences and to the presence of maternal stockpiles of initiation factors that drastically reduce the spacing between origins [START_REF] Collart | Titration of four replication factors is essential for the Xenopus laevis midblastula transition[END_REF][START_REF] Shermoen | Developmental control of late replication and S phase length[END_REF]. None of these mechanisms seems operating in mESC, which were shown to have globally similar inter-origin distances and spatio-temporal replication patterns compared to MEF [START_REF] Cayrou | Genome-scale analysis of metazoan replication origins reveals their organization in specific but flexible sites defined by conserved features[END_REF][START_REF] Panning | Spatio-temporal organization of DNA replication in murine embryonic stem, primary, and immortalized cells[END_REF]. However replication timing along chromosomes is clearly different in pluripotent stem cells and differentiated cells [START_REF] Ryba | Evolutionarily conserved replication timing profiles predict long-range chromatin interactions and distinguish closely related cell types[END_REF], and DNA replication was shown instrumental for the reprogramming of somatic cells [START_REF] Ganier | Synergic reprogramming of mammalian cells by combined exposure to mitotic Xenopus egg extracts and transcription factors[END_REF][START_REF] Tsubouchi | DNA synthesis is required for reprogramming mediated by stem cell fusion[END_REF]. How DNA synthesis induces pluripotency and what causes the RT changes specific to PSCs is still a mystery and the focus of intense studies.

Pluripotent stem cells have many properties besides fast proliferation rates and short G1 phases that distinguish them from differentiated cells. Their nucleus is smaller but their nucleo-cytoplasmic ratio is high (Suppl Fig. S3 and movies M1,2). Their chromatin is hyper-dynamic with higher rates of histone core, nuclear envelope and lamin B1 turnover, and low levels of lamin A/C [START_REF] Bhattacharya | Spatio-temporal plasticity in chromatin organization in mouse cell differentiation and during Drosophila embryogenesis[END_REF][START_REF] Constantinescu | Lamin A/C expression is a marker of mouse and human embryonic stem cell differentiation[END_REF]Meshorer et al., 2006). Many chromosomal regions of early-or latereplication (RT domains), which correlate with topologically-associated domains (TADs), change during differentiation along with lamin-associated domains (LADs) [START_REF] Hiratani | Genome-wide dynamics of replication timing revealed by in vitro models of mouse embryogenesis[END_REF][START_REF] Peric-Hupkes | Molecular maps of the reorganization of genomenuclear lamina interactions during differentiation[END_REF][START_REF] Pope | Topologically associating domains are stable units of replication-timing regulation[END_REF]. Cyclin E and A-dependent kinase activities, which are tightly cell cycleregulated in somatic cells, are constitutively high throughout the cell cycle in mESC and less-tightly regulated in hESC [START_REF] Neganova | Expression and functional analysis of G1 to S regulatory components reveals an important role for CDK2 in cell cycle regulation in human embryonic stem cells[END_REF]White et al., 2005). Stem cell metabolism and energy requirements are also distinct [START_REF] Teslaa | Pluripotent stem cell energy metabolism: an update[END_REF]. Most of these changes are probably interconnected and finding out what is the primary determinant, if any, of pluripotency will be difficult to address. Reprogramming somatic cells into iPSC has revealed an early change in expression of genes required for cell proliferation and DNA replication, and the capacity to induce DNA replication was shown to be key for reprogramming efficiency [START_REF] Polo | A molecular roadmap of reprogramming somatic cells into iPS cells[END_REF][START_REF] Tsubouchi | DNA synthesis is required for reprogramming mediated by stem cell fusion[END_REF].

Organization of DNA replication in embryonic stem cells

How DNA replication can induce cell reprogramming and a pluripotent chromatin state is unclear despite several hypotheses [START_REF] Champeris Tsaniras | Licensing of DNA replication, cancer, pluripotency and differentiation: an interlinked world?[END_REF]. All PSCs display a specific replication timing (RT) signature, with many domains switching from early-to-late (EtoL) or late-to-early (LtoE) replication compared to differentiated cells [START_REF] Ryba | Evolutionarily conserved replication timing profiles predict long-range chromatin interactions and distinguish closely related cell types[END_REF]. Yet neither the global pattern of replication foci distribution during S phase (Suppl Fig S3; [START_REF] Panning | Spatio-temporal organization of DNA replication in murine embryonic stem, primary, and immortalized cells[END_REF], nor the apparent spacing between adjacent origins, their firing efficiency or fork velocities observed on single DNA molecules seem different in stem cells [START_REF] Cayrou | Genome-scale analysis of metazoan replication origins reveals their organization in specific but flexible sites defined by conserved features[END_REF][START_REF] Schultz | Single-molecule analysis reveals changes in the DNA replication program for the POU5F1 locus upon human embryonic stem cell differentiation[END_REF]this study).

Here we performed the first quantitative study of global DNA replication in embryonic stem cells using single DNA molecules stretched by DNA combing. We define the global instant fork density (GIFD) as a novel kinetic parameter for DNA replication, which reflects the number of replication origins that are recruited genome-wide during S phase in individual cells. We found that GIFD is significantly different in murine or human PSCs compared to differentiated cells (Fig. 2F). In contrast, fork velocity (FV) and inter-origin distance (IOD) do not vary much, consistent with earlier reports [START_REF] Cayrou | Genome-scale analysis of metazoan replication origins reveals their organization in specific but flexible sites defined by conserved features[END_REF][START_REF] Schultz | Single-molecule analysis reveals changes in the DNA replication program for the POU5F1 locus upon human embryonic stem cell differentiation[END_REF]. Using a novel dual pulse-chase protocol, we show that S phase has the same duration in mESC and MEF, about 8.5 h (Fig. 3). By extrapolating our FV and GIFD measurements to the whole genome, we came to the surprising conclusion that MEF cells have a ~2-fold excess of forks relative to what would be necessary to replicate the entire genome during S phase. We interpret this excess by the recruitment of dormant origins in face of more frequent fork pausing events in MEF versus mESC, and provide experimental evidence for this model (Fig. 4). An increased use of dormant origins when replication forks slow down has been documented already 40 years ago in cells of various origins [START_REF] Ockey | The comparative effects of short-term DNA Inhibition on replicon synthesis in mammalian cells[END_REF][START_REF] Taylor | Increase in DNA replication sites in cells held at the beginning of S phase[END_REF] and later confirmed using different fork slowing conditions [START_REF] Gilbert | Replication origin plasticity, Taylor-made: inhibition vs recruitment of origins under conditions of replication stress[END_REF][START_REF] Mcintosh | Dormant origins, the licensing checkpoint, and the response to replicative stresses[END_REF]. Initially we did not favour this hypothesis as the activation of dormant origins should translate into shorter IODs, until we realized that this would not be the case if dormant origins fire later than primary origins (i.e. not during the same pulse time used for DNA combing analysis). Such a delay in the firing of dormant origins has been clearly demonstrated in yeast, either when forks progress normally or when they are slowed down by nucleotide depletion [START_REF] Pasero | Think global, act local--how to regulate S phase from individual replication origins[END_REF][START_REF] Santocanale | Activation of dormant origins of DNA replication in budding yeast[END_REF][START_REF] Vujcic | Activation of silent replication origins at autonomously replicating sequence elements near the HML locus in budding yeast[END_REF]. In mESC in contrast the lower number of active forks that we measured is sufficient to fully replicate the genome within the 8.5 h of S phase, suggesting that these forks progress continuously until they reach another fork without the requirement to activate dormant origins. We therefore propose that DNA replication is more processive and continuous in mESC, whereas it is more interrupted or discontinuous in MEF.

Consequences of interrupted DNA synthesis for chromatin assembly

What could be the consequences for chromatin and the expression/repression of developmentally Indeed some epigenetic marks, especially H3K9me3 and H2K27me3, take much longer than others to be fully re-established after passage of the replication fork, sometime more than a cell cycle (Alabert et al., Another key observation of our study is that immortalized cell lines, such as NIH 3T3 fibroblasts, and early passage MEF have a GIFD as low as that of mESC. These three types of cells share the properties of robust proliferation together with lack of p16 INK4a and p19 ARF expression, which are potent tumour suppressors targeting the Rb and p53 pathways during replicative senescence, respectively.

Strikingly we show that the GIFD increases gradually during the cultivation of MEF from passage 1 to 7 (Fig. 6B), exactly mirroring the accumulation of p19 ARF and p16 INK4a , and the cell cycle slowing of these cells [START_REF] Kim | Expression profiles of p53-, p16(INK4a)-, and telomere-regulating genes in replicative senescent primary human, mouse, and chicken fibroblast cells[END_REF][START_REF] Zindy | Myc signaling via the ARF tumor suppressor regulates p53-dependent apoptosis and immortalization[END_REF]. Thus a low GIFD correlates with full proliferation potential, while high GIFD can be considered as a new marker of replicative senescence. Our proposal that differentiated cells and pre-senescent cells face increased replication fork pausing, which is compensated by a widespread activation of dormant origins responsible for the increased GIFD, is entirely novel to our knowledge and would not have been easily detected by other methods. Fork pausing during differentiation may be dependent on the recruitment of chromatin boundary elements such as the histone demethylases Lsd1,2/SU(VAR)3.3, which are involved in the regulation of gene expression by demarcating the boundaries of eu-and heterochromatin and also required for fork pausing in fission yeast [START_REF] Dimitrova | Histone demethylases in chromatin biology and beyond[END_REF][START_REF] Holmes | Lsd1 and Lsd2 control programmed replication fork pauses and imprinting in fission yeast[END_REF][START_REF] Lan | S. pombe LSD1 homologs regulate heterochromatin propagation and euchromatic gene transcription[END_REF][START_REF] Rudolph | Heterochromatin formation in Drosophila is initiated through active removal of H3K4 methylation by the LSD1 homolog SU(VAR)3-3[END_REF]. Our study, by demonstrating that chromosome replication dynamics is different in embryonic stem cells, differentiated cells and pre-senescent cells, adds a new dimension to the exquisite plasticity of chromosome replication dynamics and proposes a model for the progressive restriction of gene expression during cell differentiation and senescence.

MATERIALS AND METHODS

Cells and cell culture

JM8.N4 mouse ES cell line was cultivated in KnockOut™ DMEM, 15% FBS (ES-tested, PAA), 1×

Non-Essential Amino Acids (1 mM), 1× L-Glutamine (2 mM), 1× Penicillin-Streptomycin, β2-Mercaptoethanol (0.1 mM) and 1000 U/ml Leukemia Inhibitory Factor (LIF, Millipore). mESC were cultivated on 0.1% gelatin-coated (Sigma) tissue culture dishes at 37°C in 5% CO2 and 21% O2. mESC cells were diluted by trypsinization and the medium was changed daily. Mouse embryonic fibroblasts (MEF) were isolated from 13.5 days embryos. Fibroblasts were cultivated in DMEM-GlutaMAX supplemented with 10% FBS (Sigma or Dutscher), Non-Essential Amino Acids and Penicillin-Streptomycin at 37°C in 5% CO2 and 21% O2. MEFs were from passage 4 or 5 unless stated otherwise.

Differentiation of mESC into NPC

For embryoid body differentiation the cells were passaged and unicellular suspension was plated on non-treated cell culture plates in KnockOut™ DMEM, 10% FBS (ES-tested, PAA), 1× Non-Essential Amino Acids (1 mM), 1× L-Glutamine (2 mM) and β2-Mercaptoethanol (0.1 mM). At the dat 4 retinoic acid (5 µM) was added. The medium was changed every two days.

At day 9 embryoid bodies were dissociated with Versene, filtred on a cell strainer and 10 7 cells were plated on 10 cm poly-L-ornithine(PLO)-treated and laminin coated dish in N2 medium. N2 medium was composed of F12/DMEM supplemented with hGF2 (10 ng/ml), EGF (10 ng/ml), apo-transferrin, insulin, putrescin (100 nM), selenite (20 nM), progesterone (20 nM), BSA (50 µg/ml) and 1× Penicillin-Streptomycin.

mNSC cells were isolated from 13.5 days embryos brain and cultivated in N2 medium on PLOtreated and laminin-coated dishes.

Analysis of replication dynamics by DNA combing

DNA preparation, DNA combing and detection was performed as described (Bialic et al., 2015b). anti-IdU clone B44 (Becton-Dickinson) and rat anti-CldU clone BU1/75 (Serotec), followed by fluorescent secondary antibodies as described (Bialic et al., 2015b). Data are from ≥ 3 biological replicas with analysis performed on blinded samples and often scored independently by two persons.

DNA Combing Imaging and Analysis

Pictures of stained, combed DNA were done on a wide-field fluorescence microscope with a 40X

BrdU/7AAD cytometric analysis

For flow cytometry, IdU/CldU-labelled cells were harvested and fixed with ice-cold ethanol added dropwise while vortexing at low speed. Cells were washed with PBS, DNA denatured with HCl 2N/0.5% Triton X-100 for 30 minutes, neutralized with 1 ml 0.1 M borate buffer (pH 8.5) and stained with anti-BrdU antibody (diluted 1:1000, #11-286-C100, Exbio) for 1 h, followed by Alexa 488-conjugated secondary antibody (diluted 1:1000, #A11017 Thermo Fisher) staining for 1 h. Finally, the DNA was stained with 14 µg/ml 7-AAD and cells were treated with 200 µg/ml RNase A for 2 h preceding the acquisition. Right before cytometry, cells were passed through a 70 µm mesh filter to eliminate aggregates. Cytometry was performed on BD FACSCalibur™ (Becton Dickinson).

Determination of S-phase duration

Non-confluent cells were treated for 30 min with 10 µM EdU, pulse-chased for variable time with medium containing 20 µM deoxythymidine and nocodazole 100 ng/ml. Then 30 min pulse with 100 µM BrdU and nocodazole 100 ng/ml (Sigma) was done. For the first point the BrdU pulse followed directly after the EdU pulse and the next ones were done for corresponding dishes of cells after increasing times (2 h, 4 h, 6 h, 8 h and 10 h). Cells were detached, fixed with ice-cold ethanol, denatured with HCl 2N/0.5% Triton X-100, neutralized as for bivariate cytometry. Click-iT reaction was performed to detect EdU and after rinses BrdU was detected with MobU-1 antibody (diluted 1:1000, #11-286-C100, Exbio, does not cross-react with EdU) and Alexa-488 secondary antibody. DNA was stained with 1µg/ml DAPI and after aggregate elimination as above, a BD FACSCanto™ II (Becton Dickinson) apparatus was used.

EdU versus BrdU signals were plotted and the number of EdU-and BrdU-positive cells that were also BrdU-positive was plotted as a function of time. Linear regression was plotted and the slope allowed calculating the S phase duration.

Quantification of gH2AX by western blot analysis

Trypsinized and PBS-washed cells were resuspended in TEB (Triton extraction buffer) containing 0.5% Triton X100, 2 mM phenylmethylsulfonyl fluoride (PMSF), 0.02% NaN3 in PBS at a cell density of

The Study of Fork Pausing Events in MEFs and mouse ESCs

Previous work from the lab has addressed the question of how mouse ESCs establish their pluripotent state and whether chromosome replication dynamics has a role in maintaining pluripotency. Using single-molecule analysis by DNA combing, the data showed MEFs use twice more replication origins compared to mouse ESCs. However, no differences were seen by measuring the fork velocities (FVs) and inter-origin distances (IODs). Despite the difference in GIFD, both mouse ESCs and MEFs spend 8.5 hours to replicate their genome. By extrapolating the S-phase duration (SPD) based on combing data, the results showed that MEFs use twice more origins than required to replicate the genome within the given SPD (Bialic et al., in preparation). Several explanations have emerged from the result that MEFs utilize twice more forks than theoretically required to replicate the whole genome. Since the FVs and the IODs were almost the same in both cell types, then we speculated that the difference in the GIFD seen between MEFs and mouse ESCs might be attributed to differences in fork progression. Because of a higher extent of heterochromatin, MEFs are expected to have a constrained fork progression where forks encounter obstacles (Replication Fork Barriers, DNA secondary structures such as G-quadruplexes, protein complexes), resulting in pausing events. As a result of pausing, more dormant origins would fire, leading to an increase in the fork density.

The approach we previously used to detect intrinsic fork pausing is by detecting the difference in the fork progression during the first and the second pulse of nucleoside analogues, without using drugs (Bianco et al., 2012). If the first and the second pulse duration was the same, the ratio of length track during the first and second pulse should be 1. However, if the fork encounters a barrier or a pause during the second pulse the ratio should be less than 1. Thus, fork the CldU track length (Figure 18,a). This suggests that pausing events cannot be detected using this protocol, or that pausing does not happen in these conditions. Increasing pyridostatin concentration to 2 µM induced fiber breakage, interrupting the CldU track which made it impossible to measure track length accurately. This is consistent with a previous study showing that 2 µM pyridostatin induces DSB and DNA damage (Rodriguez et al., 2012).

We modified the pulse protocol, and we pulsed MEFs and mouse ESCs with IdU for 7 or 15 minutes followed by 60 minutes of CldU pulse. Using this modified protocol, we decreased the chances of having a fork pausing during 7 minutes of pulse while increase it during 60 minutes pulse. The ratios of track lengths of the 60 minutes to 7 minutes or 15 minutes pulses were determined in both MEFs and mouse ESCs. If there were more pausing events during the 60 minutes CldU pulse, the ratio of track length of 60 minutes pulse by 7 or 15 minutes pulse should be higher than when the pausing events are absent. The results show a significant decrease in the track ratios in mouse ESCs compared to MEFs. This actually goes against to what we were expecting (Figure 18,b). Using single-molecule analysis by DNA combing, we were able to assess the distance between origins IOD, so we should also take into consideration that the inter-origins distance 98 and 86 kb in mouse ESCs and MEFs respectively (Figure 19.).

Within 60 minutes pulse the fork travels around 120 kb (given that the fork velocity is around 2 kb in mouse ESCs) thus using these conditions we might be at risk of having firing of new origins during the CldU pulse that might merge with earlier forks. One explanation would be that the possibility of firing of new origins during the 60 minutes CldU pulse in MEFs cannot be excluded. This comes from the difficulty of recognizing new origin firing during a 1-color pulse. If that was true, the CldU track length would be longer due to merging of newly-fired origins with the initial one. In the absence of pausing, the ratio of CldU/IdU pulse should be 8.6 and 4, for 7 and 15 minutes pulse respectively. Mouse ESCs showed ratios of 6.4 and 3.3, suggesting fork slow down by 26 and 18% respectively or hitting replicated regions in mouse ESCs rather than in the MEFs. This led us to test another hypothesis that could explain the increase in GIFD in MEFs.

The Study of Origin Distribution in the Genome of MEFs and mESC

Considering the fact that mouse ESCs display an open euchromatin that becomes more compact and heterogeneous upon differentiation, we hypothesized that the origin clustering and distribution of origins may vary between mouse ESCs and MEFs. Thus, mouse ESCs might have a more even distribution of origins in their genome while MEFs would have more clustered origins. By measuring IODs, our previous results showed no significant difference in the IODs between MEFs and mouse ESC. However, it has been previously shown that the DNA combing technique is limited by the fact that DNA fibers get fragmented during preparation. Thus, we are limited by the size of DNA fibers (Técher et al., 2013a). Given the fact that the distance between origin clusters might vary between 0.5-2 Mb, we speculated that the IOD values we obtained in the analysis of MEFs are the IODs between origins of the same cluster, overlooking the IODs between origins from different clusters due to limited size of DNA fibers. Thus, we decided to improve our DNA combing protocol to be able to include longer DNA fibers in our analysis.

DNA combing has been performed on both MEFs and mouse ESCs. IODs have been measured

in both cell types. In order to exclude that the difference in IODs we see might be due to the difference in DNA fragmentation between both samples, only 500-1200 kb fibers were taken into consideration to measure IODs. Thus, the difference in DNA fiber lengths that have been analyzed in MEFs and mouse ESCs is not significant (Figure 20 This change in IOD in MEFs confirms that we had been missing IODs between origins from different clusters. Furthermore, the absence of this change in mouse ESCs even with longer fibers strengthens our hypothesis that origins are not as clustered as in MEFs. This is consistent with a scenario in which the heterogeneity in the distribution of origins in MEFs (and probably other differentiated cells) causes the forks coming from different clusters to travel more distance, giving time to late origins to be activated, if they were licensed. This would explain the higher fork density in MEFs than in mouse ESCs, along with a similar SPD.

If this hypothesis was true, relaxing the chromatin in MEFs should create a more homogeneous distribution of origins and a reduced GIFD. TSA, a histone deacetylase inhibitor, was shown to induce a reversible decondensation and more homogeneous chromatin distribution

A B

in interphase chromatin (Tóth et al., 2004). In order to avoid cellular arrest caused by TSA treatment, a very low concentration of TSA, 20 nM, was used to treat MEFs for 1, 4.5 or 10 hours followed by IdU and CldU pulses 15 minutes each, a thymidin chase for 2 hours before HMW DNA preparation for DNA combing. GIFD analysis of the above mentioned samples showed a decrease from 1.4 forks/Mb in the control to 0.9, 1 and 0.7 forks/Mb respectively (Figure 21). In order to test this hypothesis, we previously pulsed labeled mouse ESCs with IdU/CldU for 15 min each followed by thymidine chase then cells were fixed and sorted according to DNA content into early, mid and late S phase (Supl. Following sorting each sub-fraction of S phase was collected and prepared for combing.

GIFD was calculated in each of the sub-fractions. The results show that unlike mouse ESCs, MEFs display a variation in the number of active origins throughout the different sub-phases where it starts with 0.99 in S1 and increases to 1.28 in S2 then decreases sharply to 0.54 in S3 (

A B C

In this work we addressed the impact of alteration of S-phase dynamics on cell cycle duration and genome instability. We were able to show that cancer cell lines utilize few numbers of origins to replicate their genome leading to an extension in the duration of S phase and the cell cycle. Despite this extension cancer cells would still enter mitosis with unreplicated DNA and finish replication using MiDAS which is also associated with increased mitotic defects. This alteration in S-phase dynamic might be attributed to origin under-licensing that we were able to rescue by treating with low dose of Palbociclib.

Our results on intermitotic duration by analyzing single cells reveal that almost all of cancer cell lines spend longer time 17-21 hours to enter a second cell cycle compared to the untransformed cell lines. This goes in agreement with previously published data (Bazzocco et al., 2015;Khosravi-Maharlooei et al., 2015) that cancer cell lines show variability in the doubling time and almost all of them spend more than 24 hours to finish a full cell cycle. However, in these two studies the doubling time was assessed using population doubling time by counting the number of cells over several cell cycles without taking into consideration the percentages of proliferating and non-proliferating cells. The reason why this parameter is important comes from the fact that untransformed cells shift between proliferative and quiescent state where the restriction point is active and regulates cellular reentry into another cell cycle. On the other hand, cancer cell lines are more into a continuous proliferative state than quiescent. This would create a bias while calculating the doubling time based on population. Thus, we decided to follow intermitotic time of proliferating cells in both untransformed and cancer cell lines.

In the first part of my results, we were the first to provide the link between the impact of using fewer origins on S-phase length and mitosis. We provided evidence of common features among fourteen different cancer cell lines where they have reduced replication potency compared to untransformed cell lines despite the variability in genome size. All of the tested cancer cell lines displayed a reduction in replication potency accompanied with an extension in S-phase duration. We used a direct method to measure S-phase duration (Bialic et al. in preparation), and we are the first to show that cancer cell lines extend their S phase to be able to finish replication.

This reduction in replication potency might be attributed to origin under-licensing. Origin licensing occurs in two waves in mammalian cells. In vivo licensing assay was used to study MCM loading dynamics during the cell cycle in MCF-7 breast cancer cell line. MCF-7

expressing GFP-MCM2 or GFP-MCM4 were synchronized in G2/M and were released in fresh medium after mitotic shake-off. Cells were analyzed by FRAP in early, mid and late G1. A small fraction of MCM was immobile during early G1 confirming MCM binding to the chromatin.

This immobile fraction increased in late G1 suggesting another wave of licensing took place. In addition, the majority of MCM loading occurred in late G1 when compared to early G1 (Symeonidou et al., 2013). These results suggest that early S phase entry would cause the cells to escape the second wave of licensing where the majority of MCM loading occurs. This might lead to a reduction in the fork density. We were able to rescue this phenotype upon treatment of RBpositive cell lines with palbociclib, reinforcing our that most of cancer cell lines are underlicensed. We were not able to see a difference in the MCM loading on the chromatin; however, this does not exclude that there is a difference in the MCM loading between DMSO and palbociclib-treated HCT116 since we were not able to detect a difference in the MCM loading even in U2OS transfected with CDT1 vector. An ideal experiment to test this hypothesis directly is to reduce licensing in an untransformed cell line through targeting either CDT1 or CDC6 and follow the consequences on S phase dynamics and mitosis compared to the control.

However, despite the extension in S phase, cells still enter M phase with under-replicated DNA and try to finish replication in mitosis, MiDAS. These under-replicated regions were shown to colocalize with FANCD2 foci defining CFS. These sites are known to harbor hotspots of genomic rearrangements and viral integration emphasizing on the role of these sites in inducing genome instability. In an ideal experiment, each cancer cell line should be compared to a control originating from the same tissue and individual. However, we were restricted with the availability of untransformed cell lines known to propagate well in culture. The aim was to have a global idea about the differences in replication dynamics in known and commonly used cell lines in basic cancer research.

Relevant work was published this year. Pennycook and colleagues were able to measure Sphase duration using another approach. T98G glioblastoma cell line was transfected with siE2F6.

E2F6 is a transcription factor from the E2F family known to have an inhibitory activity. Thus knocking down E2F6 should relieve the inhibition on E2F target genes and induce S-phase entry similar to knocking down RB. Both control and transfected cells were synchronized by serum starvation then released in serum and the time of S phase exit was monitored in both samples.

The results showed that siE2F6-transfected cells have shorter S-phase duration compared to the control. While this contradicts our results, the authors claimed that the S-phase entry was not affected whereas the FACS profiles shown indicate that siE2F6-transfected cells enter S phase earlier than the controls. This leads to a bias in estimating the S-phase duration (Pennycook et al., 2020).

In 2017, Matson and colleagues provided evidence that fast MCM loading on chromatin in G1 phase is characteristic of human ESCs (Matson et al., 2017). This MCM loading slows down along with G1 lengthening upon differentiation. The fast loading is attributed to increased expression of CDT1 and to post-transcriptional modifications that lead to CDT1 accumulation in the preceding G2 phase. Although extending G1 phase in human ESCs is correlated with increased differentiation (Coronado et al., 2013a), this differentiation could be due to G1 that altering origin licensing in hematopoietic progenitors affects the ability of these cells to mature or regenerate (Alvarez et al., 2015). Given the fact that G1 phase is shorter in mouse ESCs compared to MEFs, altering Mcm3 at the same level in both cell types might lead to differences in the degree of origin licensing. That might be one of the reasons why lethality is more evident in mouse ESCs upon Mcm3 double knock out. Alternatively, the known excess of MCM complex proteins might become important only in rapidly-proliferating tissue, such as embryonic cells, but not in more mature, less proliferative cells.

Our data on mouse ESCs and MEFs suggest that MEFs utilize 2-fold more origins than required to replicate the genome. We have tested whether the increase in origin density could be attributed to fork pausing events in MEFs. These fork pausing events would allow the firing of dormant origins, explaining the increase in fork density in MEFs. Unexpectedly, our results revealed more pausing events in mouse ESCs compared to MEFs. Then why ESCs would not fire dormant origins? A previous study showed that mouse ESCs have characteristic accumulation of ssDNA gaps along with increased frequencies of fork reversal. These two phenotypes are suppressed upon induction of differentiation. This would explain the decrease in the fork density? in mouse ESCs but not in MEFs. The lack of activation of dormant origins might be attributed to weak checkpoints that mouse ESCs have (Aladjem et al., 1998).

Moreover, downregulation of RAD51, a key player in fork reversal, in mouse ESCs led to an increase in fork speed along with an accumulation of chromosomal fragments. This suggested that fork reversal is an essential mechanism in mouse ESCs to prevent chromosomal breakages.

This might be another reason why checkpoints or dormant origins are not activated in mouse ESC with endogenous replication stress. Furthermore, despite the fact that MEFs display lower levels of replication stress marks, they display higher levels of mitotic defects compared to mouse ESC. This goes further with the suggestion that mouse ESCs are tuned to tolerate replication stress even in mitosis [START_REF] Ahuja | A short G1 phase imposes constitutive replication stress and fork remodelling in mouse embryonic stem cells[END_REF]. The question remains as to how they cope with under-replicated regions upon differentiation. It has been shown in the same study that ESCs lack 53BP1 nuclear bodies, but start to acquire them upon differentiation. This suggests that under-replicated regions in mouse ESCs are channeled to the next cell cycle for repair and they are protected by fork reversal. Whereas upon differentiation these regions are assembled in 53BP1 nuclear bodies until the G1phase before the next S-phase entry.

Another reason behind the difference in fork densities between mouse ESCs and the differentiated counterparts might be attributed to both origins distribution and reprogramming of replication timing. In our study we used IOD as a parameter to study the difference in the distribution of origins in mouse ESCs and MEFs. We first showed that IODs did not change between both cell types. However, when the DNA combing protocol was optimized to analyze longer fibers, we can clearly see that MEFs have longer IODs compared to mouse ESCs. This suggests that we previously missed IODs between different clusters in MEFs, due to fewer Mbrange DNA fibers. In addition, this also emphasizes our hypothesis that the distribution of origins between both cell types is not identical. However, if mouse ESCs had a more homogenous distribution of origins, the GIFD should be higher in the mouse ESCs than MEFs with the given IOD in mouse ESCs. This led us to speculate that there must be a difference in the replication timing program that allows mouse ESCs to use fewer numbers of origins compared to MEFs.

Replication timing analysis and Hi-C were performed at one day intervals on single cells during mouse ESC differentiation. The results showed switching in the replication timing from early to late or late to early among the genome. This switching in the replication timing was shown to be correlated with compartment A to B or B to A switching respectively (Miura et al., 2019).

However, the proportions of early-to-late or late-to-early switching was not calculated during differentiation when compared to mouse ESCs. Thus, we have previously sorted S-phase cells of mouse ESCs and showed that S1, S2 and S3 fractions maintain the same GIFD. However, while sorting MEFs S1 had a low GIFD that increased in S2 then decreased in S3 to a degree even lower than S1. We speculated that the very low GIFD in S3 forces the cells to take a long time to finish replication that might allow the firing of dormant origins if they were licensed. However, the results of three different experiments gave the same trend but with different values so we had a problem of reproducibility. We have already excluded the effect of sorting on this variation between biological experiments that might be attributed to stress induced by laser light. The cells were already pulsed and fixed before sorting.

To finalize with, I mentioned in the introduction that patients with Meier-Gorlin Syndrome Le licensing des origines de réplication est une étape indispensable à leur fonctionnement et qui a lieu en phase M/G1. Il consiste en un chargement de complexes pré-réplicatifs (pre-RC) sur un grand nombre de sites chromosomiques dont certains seront activés comme origines en phase S.

Des travaux sur la levure dans notre laboratoire ont montré que la dérégulation de G1 empêche l'obtention d'un licensing complet. En conséquence de la dérégulation de G1, les cellules traversent la phase S avec moins d'origines actives, retardant l'achèvement de la réplication chromosomique jusqu'après le début de la mitose. Cela entraîne des ruptures de chromosomes, des réarrangements et une mauvaise ségregation (Lengronne and Schwob, 2002a;Tanaka and Diffley, 2002a). En utilisant une analyse single-molecule de la réplication de l'ADN, notre laboratoire a montré que les fibroblastes murins dépourvus des trois paralogues de pRb (rétinoblastome) qui contrôlent la progression G1/S ont une densité globale instantanée en fourches de réplication (GIFD) plus faible ainsi qu'une phase S prolongée (Coulon et al., in preparation).

Le premier objectif de ma thèse était de vérifier si ces phénotypes (faible GIFD, longue phase S) sont fréquents ou prédominants dans les cellules cancéreuses. Jusqu'à présent, quatorze lignées de cellules cancéreuses différentes provenant de neuf tissus épithéliaux différents ont été analysées. En utilisant une nouvelle technique développée au laboratoire, nous avons déterminé que la durée de la phase S (SPD) était très variable et significativement plus longue (10-29h) dans toutes les lignées de cellules cancéreuses testées par rapport aux fibroblastes humains normaux, aux cellules épithéliales mammaires humaines (HMEC) et rétiniennes pigmentaires (RPE), qui ont montré des SPD robustes et plus courtes (7-9h). L'extension de la phase S dans les cellules cancéreuses est probablement due à l'utilisation d'un moins grand nombre d'origines de réplication, car la GIFD mesurée par la technique de peignage moléculaire de l'ADN était généralement inférieure pour les 14 lignées de cellules tumorales, par rapport aux cellules non transformées.

Conti et al ont fourni des preuves de la plasticité de réplication, où la densité d'origine est inversement corrélée avec le taux de progression des fourches afin de conserver une réplication efficace (Conti et al., 2007). En effet, nous avons constaté que certaines lignées de cellules cancéreuses compensent en partie leur faible GIFD en augmentant la vitesse de fourche (FV).

Nous avons défini un nouveau descripteur de la réplication appelé puissance de réplication l'extension de la phase S. Nos résultats montrent que le traitement de l'HCT116 avec une faible dose de palbociclib provoque une diminution de 1,3 heures de la durée de la phase S par rapport à la phase non traitée, confirmant l'influence de la durée de la phase G1 sur celle de la phase S.

Ce projet représente le coeur de ma thèse où j'ai obtenu la plupart de mes résultats.

Objectifs 1) Vérifier si ces phénotypes (faible GIFD, longue phase S) sont fréquents ou prédominants dans les cellules cancéreuses.

2) Déterminer les conséquences de la diminution du nombre d'origines de réplication utilisées et de l'extension de la phase S sur la ségrégation chromosomique et l'instabilité du génome dans les lignées de cellules cancéreuses, en utilisant la vidéomicroscopie.

3) Rétablir la puissance de réplication dans l'une des lignées de cellules cancéreuses testées en prolongeant G1 à l'aide de Palbociclib, un inhibiteur de cdk4/6, et vérifier si cela permettrait de rétablir la longueur normale de la phase S. Afin de voir si ces lignées de cellules cancéreuses ayant une faible GIFD et une phase S prolongée présentent des défauts mitotiques, je suis en train de tester si la réplication et/ou la réparation des chromosomes est achevée pendant la mitose en évaluant la synthèse mitotique de l'ADN ou MiDAS (Bhowmick et al., 2016b;Minocherhomji et al., 2015). (Coronado et al., 2013a). Bien que G1 soit courte, l'efficacité de licensing de l'ADN dans les ESC semble due à la charge rapide des MCM et à l'expression accrue de CDT1 et CDC6 (Matson et al., 2017). En outre lors de la différenciation des ESC, des ESC, certains domaines de timing de réplication deviennent plus vastes. Les domaines qui changent de timing sont appelés développementaux, et ceux qui restent les mêmes constitutifs.

Cela suggère que la régulation du timing de réplication pourrait être impliquée dans la différenciation cellulaire (Dileep et al., 2015a)

First

  I would like to thank Anne-Marie Martinez, Chantal Desdouets, Valeria Naim and Jean-Charles Cadoret for investing their time in evaluating my work where I hope it would meet your expectations. Special thanks go for Valeria and Claude Sardet for following my work during the four years and for their valuable input.

Figure 1 .Figure 2 .Figure 3 .Figure 4 .Figure 5 .Figure 6 .Figure 7 .Figure 8 .Figure 11 .Figure 12 .Figure 13 .Figure 14 .Figure 15 .Figure 16 .Figure 17 .Figure 18 .Figure 20 .Figure 22 . 1 .

 12345678111213141516171820221 Figure 1. Regulation of the cell cycle by RB, cyclins and CDKs in mammalian cells. ......................................... 19 Figure 2. Scheme showing the pathway by which Palbociclib inhibits Cdk4/6 .................................................... 24 Figure 3. Regulation of DNA replication during G1 and S phases. ....................................................................... 31 Figure 4. Arrangement of a replication cluster and chromatin-loop formation .................................................. 34 Figure 5. Replication patterns of early versus late replicating foci in mid and late-S phase and G2 cells......... 35 Figure 6. Replication timing profile of different chromosome segments located in both early and latereplicating compartments ......................................................................................................................................... 37 Figure 7. Different stages of embryonic development in mouse ............................................................................ 45 Figure 8. Replication in silent versus active genes .................................................................................................. 53 Figure 9. A summary of the mechanism by which origin licensing alteration induces genome instability ........ 60 Figure 10. Patterns of FRA3B probe in red of S phase nuclei labeled with BrdU in green ................................ 64 Figure 11. Paucity of initiation events model explaining fragility of FRA3B in human lymphocytes. ............... 65 Figure 12. Model explaining the response of cells to replication stress ................................................................. 66 Figure 13. Mechanisms of classical and alternative NHEJ. ................................................................................... 68 Figure 14. Scheme summarizing the role of 53BP1 in preventing the formation of ultrafine anaphase bridges ..................................................................................................................................................................................... 73 Figure 15. Intermitotic times measured in untransformed and cancer cell lines using time-lapse microscopy ................................................................................................................................................................................... 137 Figure 16. Quantification of DNA content in each cell line by DAPI FACS analysis ........................................ 143 Figure 17. Absence of fork pausing events during the IdU/CldU pulses in both MEFs and mouse ESCs. ...... 204 Figure 18. Detecting fork pausing events using single-molecule analysis by DNA combing ............................. 206 Figure 19. Inter-origins distances measured in mouse ESC and MEFs using single molecule-analysis by DNA combing .................................................................................................................................................................... 207 Figure 20. Single molecule analysis of IOD display significant decrease in IOD in mouse ESC compared to MEFs ......................................................................................................................................................................... 209 Figure 21. Single-molecule analysis of MEFs treated with 20 nM TSA for 1, 4.5 and 10 hours shows a decrease of GIFD in TSA treated samples ............................................................................................................. 210

Figure 1 .

 1 Figure 1. Regulation of the cell cycle by RB, cyclins and CDKs in mammalian cells ( Adapted from Suryadinata et al., 2010).

Figure 2 .

 2 Figure 2. Scheme showing the pathway by which Palbociclib inhibits Cdk4/6 (adapted from (Suryadinata et al., 2010).

Figure 3 .

 3 Figure 3. Regulation of DNA replication during G1 and S phases adapted from (Fragkos et al., 2015).

Figure 4 .

 4 Figure 4. Arrangement of a replication cluster and chromatin-loop formation adapted from (Conti et al., 2007).

Figure 5 .

 5 Figure 5. Replication patterns of early versus late replicating foci in mid and late-S phase and G2 cells (Ma et al., 1998a).

  , b). The organization of chromatin in 3-dimensions comprises long-range chromatin interactions captured by Hi-C method. Such interactions are referred to topologically-associated domains (TADs). TADs define two types of compartments based on their gene expression, compartment A and compartment B. During S phase DNA replication begins within the TADs residing in the nuclear interior in compartment A, correlated with actively transcribed open chromatin. Then it gradually moves to the later TADs in compartment B, correlated with silent compact chromatin then finally to LAD. The progression of DNA replication between RDs of different replication timings forms TTRs which are origin-depleted regions replicated passively by neighboring forks

Figure 6 .

 6 Figure 6. Replication timing profile of different chromosome segments located in both early and latereplicating compartments (Marchal et al., 2019).

Figure 7 .

 7 Figure 7. Different stages of embryonic development in mouse (adapted from (Davidson et al., 2015).

Figure 8 .

 8 Figure 8. Replication in silent versus active genes. A) In silent genes the progressing fork do not encounter RNA polymerase II which acts as an obstacle, thus replication progresses normally without inducing genome instability. B) In long actively transcribed genes transcription starts in G2 and mitosis and even continues till S phase. Upon the onset of DNA replication, the progressing fork encounters RNA polymerase II which acts as a barrier that perturbs the replication fork inducing genome instability (Helmrich et al., 2011).

Figure 9 .

 9 Figure 9. A summary of the mechanism by which origin licensing alteration induces genome instability. The presence of under-licensed origins prevent the cells form finishing replication of the whole DNA template. Whereas origin relicensing case the same DNA segment to b replicated more than one time inducing fork collisions and collapse. Ectopic origin licensing increase the probability of having replication-transcription collisions leading to fork collapse (Petropoulos et al., 2019)

FRA3B,

  one of the most commonly expressed CFS in human lymphocytes, is correlated with replication timing. Fluorescent in situ hybridization (FISH) was used to study the replication timing of FRA3B. The technique is based on denaturing nuclei and incubating them with known gene probes to check for allele copy number. Probe present two times in the nucleus indicates the presence of two alleles (Figure10, a). Whereas the presence of closely apposed doublets on each of the probe signals indicates that the allele has been replicated (Figure10, b and c). The results showed that the probe becomes doublets at the end of S phase compared to the early and late replicating control probes. Treatment of lymphocytes with aphidicolin further delayed the replication, and cells entered G2 with two probe signals but not in doublets. This implied that under replication stress conditions, the cells would exit S phase with late-replicating regions being incompletely replicated. (Le[START_REF] Beau | Replication of a common fragile site, FRA3B, occurs late in S phase and is delayed further upon induction: implications for the mechanism of fragile site induction[END_REF].

Figure 10 .

 10 Figure 10. Patterns of FRA3B probe in red of S phase nuclei labeled with BrdU in green (Le Beau et al., 1998).

Figure 11 .

 11 Figure 11. Paucity of initiation events model explaining fragility of FRA3B in human lymphocytes. Absence of initiation events within the FRA3B core in lymphocytes in unpurturbed S phase drives it to fail to activate the dormant origins in response to the replication stress induced by aphidicolin compared to fibroblasts (Debatisse et al., 2012).

Figure 12 .

 12 Figure 12. Model explaining the response of cells to replication stress. The model shows two adjacent origin clusters one that fires earlier than the other and each has a certain number of fired origins to replicate the region in unperturbed condition. As the forks encounter a barrier or upon slow down, S phase checkpoint is activated which leads to the activation of the adjacent dormant origins within the same cluster and inhibits the firing of origins in the later cluster (Ge and Blow, 2010).

Figure 13 .

 13 Figure 13. Mechanisms of classical and alternative NHEJ. A) Induction of DSB is repaired via: B) Microhomology-mediated end joining which involves the resection of the broken ends associated with microhomologous annealing of both strands. This resultsin loss of DNA sequence. B) Classical NHEJ where KU proteins bind to the broken ends to prevent DNA resection followed by the recruitment of polymerases and ligases. This type of repair results in minimal DNA gain or loss compared to the two other alernatve pathways. C) Polymerase Ѳ-mediated end joining where short regions of homology between DNA strands are extended by polymerase Ѳ. This would result in either insertions or deletions (Rodgers and McVey, 2016).

  through the stabilization of cyclin B and securin. Securin ensures the attachment of sister chromatids. CDK1 in turn phosphorylates monopolar spindle 1 (MPS1) allowing its recruitment to kinetochores. The recruitment of MPS1 to kinetochores allows additional recruitment of SAC effectors. Once chromosomes are aligned properly on the metaphase plate, APC becomes activated and targets cyclin B and securin for degradation allowing the initiation of anaphase. It has been reviewed that DDR in mitosis shares common initial effectors as the DDR induced in interphase cells. However, mitotic DDR has signaling cascades which are quite different from the DDR induced in interphase cells. These signaling cascades in mitotic DDR might be more robust and induce cellular arrest in G1 of the next cell cycle(Ovejero et al., 2020).

Figure 14 .

 14 Figure 14. Scheme summarizing the role of 53BP1 in preventing the formation of ultrafine anaphase bridges. If the cells fail to finish replication before the onset of mitosis, the cells enter the first nitosis with underreplicated regions. The replication in these regions should be then resumed in RAD52-dependent MiDAS pathway. This is considered to be the first chance in resolving DNA replication. If repair was efficient, the cells enter the second interphase. However, if the DNA repair mediated by MiDAS was not efficient, ultrafine chromosome bridges are formed leading to the formation of 53BP1-nuclear bodies in the G1 phase of the second interphase. 53BP1-nuclear bodies in turn recruits RIF1 to presume the replication of these regions and enables the handover of these regions to RAD52-targeted repair. This allows efficient repair in these regions. Whereas defects in RIF1 or 53BP1-nuclear bodies formation enables the handover of these regions to the RAD51-mediated repair which is known to be more toxic than RAD52-mediated pathway (Spies et al., 2019).

Fisher) and 2

 2 mM L-glutamine (25030-024, Thermo Fisher). Immortalized RPE1-hTERT cells were cultured in DMEM-F12 without glutamine (21331-020, Thermo Fisher), supplemented with 10% fetal bovine serum (CVFSVF00-01, Eurobio) and 100U/mL penicillin-(151-40-122, Thermo Fisher) and 2mM L-glutamine (25030-024, Thermo Fisher). Immortalized HMEC-hTERT cells were cultured in serum-free MEGM medium (CC-3150, Lonza) supplemented with bullet kit containing 5 ng/mL epidermal growth factor, 0.5 µg/mL hydrocortisone, 5 µg/mL insulin, 70 µg/mL bovine pituitary extract, 5 µg/mL transferrin, 10 -5 M o-phosphoethanolamine (CC-3151 & CC-4136, Lonza) and 100 U/mL penicillin-streptomycin Thermo Fisher).

  TE buffer was removed, and one agarose plug was equilibrated in 1 mL of 2-(Nmorpholino) ethanesulfonic acid (MES) buffer (Sigma) at 65°C for 30 minutes. DNA solution was further digested in 3U of β-agarase (NEB) overnight at 42°C. Tubes were handled with extreme care after the melting step to prevent DNA breakage. Three unites of agarase were added to the DNA solution and incubated at 42°C for 2 hours.Buffers;T10E1: 10 mM Tris-HCl pH=7.5, 1 mM EDTA. T10N100E50: 10 mM Tri-HCl pH=7.5, 100 mM NaCl, 50 mM EDTA.Proteinase K buffer: 10 mM Tris-HCl pH=7.5, 20 mM NaCl, 0.5% SDS, 100 mM EDTA, 0.2 mg/mL proteinase K 20mg.MES: 50 mM MES, 1 mM EDTA, 100 mM NaCl1.3.3. DNA Combing from DNA Solutionβ-agarase was inactivated by incubating the DNA solution at 65°C for 10 minutes. DNA solution was left to cool down for 5 minutes at RT before combing.Teflon tanks were filled with DNA solution, and 22x22 mm silanized coverslips were dipped for 5 minutes using a homemade DNA combing machine. Silanized coverslips were removed at a constant speed of 300 µm/s. Combed slides were fixed on Whatman paper and left to dry at 65°C for more than 1 hour.1.3.4. Immunodetection of Combed DNADNA coverslips were denatured in freshly prepared 0.5N NaOH for 25 min. DNA coverslips were neutralized in 3 washes of PBS 1X pH=7.5, 3 minutes each. DNA coverslips were blocked in PBS 1X-1% BSA-0.1% tritonX-100. DNA coverslips were incubated in primary antibodies solution containing anti-CldU (rat anti-BrdU, ab6326, Abcam, 1/40) and anti-IdU (mouse anti-BrdU, 347580, Becton Dickinson, 1/10) diluted in PBS 1X-1% BSA-0.05% tritonX-100 in humid chamber at 37°C for 30 minutes. Coverslips were washed 3 times with PBS-0.05% triton X-100, 3 minutes each. DNA coverslips were incubated in secondary antibodies solution containing Alexa488 Goat anti Rat IgG ( A11006, Life Technologies, 1/50) and Alexa 546 Goat anti Mouse IgG (A11030, Life Technologies, 1/50).

  Extracted Chromatin Cells were washed twice with PBS-1X followed by resuspension with 200 µl of 1% BSA + PBS + 0.1% NP-40 containing primary antibody anti MCM2 BM28 diluted 1:200 (610700 mouse monocolonal, BD Biosciences). The cells were incubated for 1 hour at room temperature. Cells were then washed with 1% BSA + PBS + 0.1% NP-40 and incubated with 200 µl of 1% BSA + PBS + 0.1% NP-40 containing secondary antibody Alexa 488 Fab goat anti-mouse (A11017, ThermoFisher, 1/1000) for 1 hour at room temperature. Cells were washed with 1% BSA + PBS + 0.1% NP-40 then incubated in 0.5 µg/mL DAPI (Sigma) solution containing 200 µg/mL RNase A overnight at 4 ºC. mouse ESC cell line was cultured in 0.1%gelatin coated petri dishes. The medium used was KnockOut TM DMEM (10829-018, Thermo Fisher) supplemented with fetal bovine serum (A15-208, ES cell-tested, PAA, GE Health Life Science), 100 U/ml penicillinstreptomycin (151-40-122, Thermo Fisher), 1mM non-essential amino acids (11140-035, ThermoFisher), 2 mM L-glutamine (25030-024, Thermo Fisher), 0.1 mM β-mercaptoethanol(31350-010, ThermoFisher) and 1000 U/ml LIF (Merck Millipore). The cells were maintained at 37ºC, 21% O2 and 5% CO2 incubator. The medium was changed every day and cells were passaged each 2 days.MEFs have been cultured in DMEM-Glutamax (31966-021, Thermo Fisher) supplemented with 10% fetal bovine serum (31966-021, Thermo Fisher), 1 mM 1mM nonessential amino acids (11140-035, ThermoFisher) and 100 U/ml penicillin-streptomycin Thermo Fisher).

4)

  Determine the consequences of decreased replication origin usage and S-phase extension on chromosome segregation and genome instability in cancer cell lines using EdU incorporation on mitotic chromosomes and determining the mitotic defects associated with these phenotypes. Decreased replication potency and extended S phase, due to G1 shortening and replication origin under-licensing, are common features of cancer cells. Baraah Al Ahmad Nachar, Marjorie Drac, Lena Roumegous, Philippe Coulombe, Virginie Georget # , Vincent Coulon* & Etienne Schwob* IGMM, Univ Montpellier, CNRS, Montpellier, France # BioCampus Montpellier, Univ Montpellier, CNRS, INSERM, Montpellier, France Correspondance : Etienne Schwob (schwob@igmm.cnrs.fr) or Vincent Coulon (vincent.coulon@igmm.cnrs.fr) Institut de Génétique Moléculaire de Montpellier, CNRS UMR5535, 1919 route de Mende, 34293 Montpellier cedex 5, France

  cancer cell lines,HCC-1937, HCC-38, Colo205, Colo320, 786-O, BXPC-3, ASPC-1, PC-3, 22RV1, COV318 and MKN-45, were provided by the Montpellier Integrated Cancer Center (SIRIC Montpellier) and cultured according to the instructions given by ATCC. Among these cells, the Cov318 was cultured in DMEM/Glutamax-I medium supplemented with 10% fetal bovine serum (Eurobio) and 100U/ml penicillin-streptomycin, while the remaining ones were cultured in RPMI supplemented with 10% fetal bovine serum (Eurobio) and 100 U/mL penicillin-streptomycin. HeLa, U2OS and HCT116 were cultured in DMEM/Glutamax-I medium supplemented with 10% Hyclone fetal bovine serum (GE Health Care Life Sciences) and 100 U/mL penicillin-streptomycin. Cells were maintained in 37 C humid incubator in the presence of 5% CO2 and 21% O2.

  (25 µM, 15 min) and CldU (200 µM, 15 min), with the exception of U2OS cells which were pulsed for 30 minutes each, then chased with 200 µM thymidine for 1-2 hours. 125,000 cells were used to prepare agarose plugs for DNA purification and molecular combing. The rest of cells (about 10 6 ) were collected and fixed with cold ethanol for determining cell cycle profiles by flow cytometry. Low-melting agarose plugs were incubated twice in proteinase K (0.2 mg/mL, overnight at 37°C), washed 5x with Tris-NaCl-EDTA buffer, then melted in 2-(N-morpholino) ethanesulfonic acid (MES) buffer at pH 5.7 for 30 minutes at 65°C, and digested twice with 3U of β-agarase (NEB), overnight at 42°C and for an additional 2 hours with fresh β-agarase. Silanized glass coverslips were dipped for 5 min in the gDNA solution using a homemade DNA combing machine and then removed at a constant speed of 300 µm/s. DNA fibers were cross-linked to coverslips for 1 hour at 65°C, then denatured in freshly prepared 0.5N NaOH for 25 min, washed, neutralized with PBS then blocked in PBS-1% BSA-0.1% TritonX-100. DNA coverslips were incubated in primary antibodies solution containing anti-CldU (rat anti-BrdU, ab6326, Abcam, 1/40) and anti-IdU (mouse anti-BrdU, Becton Dickinson, 347580, 1/10) followed by secondary antibodies solution containing (Alexa488 Goat anti Rat IgG, A11006, Life Technologies, 1/50) and (Alexa 546 Goat anti Mouse IgG, A11030, Life Technologies, 1/50), then counterstained for DNA (mouse anti-ssDNA, Dshb autoimmune ss-DNA, 1/100) and secondary antibody (goat anti-mouse IgG2a Alexa647, A21241, Life Technologies, 1/50), then mounted with ProlongGold anti-fade reagent. Data analysis was performed double blinded.The acquisition of DNA combing images was done using an epifluorescence Zeiss AxioImager Z1 microscope using a 40x objective. For measuring global instant fork density (GIFD), images were acquired without looking for IdU/CldU signals, based solely on DNA fibers (Cy5 channel). A minimum of 90 Mb of DNA was analyzed. For fork velocities, IdU tracks were measured. DNA combing images were analyzed using IDeFix (Intelligent DNA Fiber Identification and Analysis software) developed in the lab.

Figure 1 :

 1 Figure 1: Cancer cell lines divide more slowly than untransformed cells. A) Inter-mitotic times of individual cells were determined using phase-contrast live cell microscopy over 72h at 10 min interval. Between 65 and 189 inter-mitotic times were scored for each cell line. Median with interquartile range, Wilcoxon test P=2.2x10 -16 between untransformed and transformed groups. B) Examples of bivariate cell cycle profiles for transformed and untransformed cell lines. Gates indicate G1, S and G2+M fractions for each cell line. 22RV1 is a heteroploid prostate cancer cell line. C) Cancer cell lines (in purple) proliferate continuously compared to the untransformed controls (in black). The graph shows the percentage of cells that did not enter the cell cycle (BrdU-) when pulsed with BrdU for the indicated times. The samples were analyzed by FACS (n=2) presented as a mean with SEM.

Figure 2 :

 2 Figure 2: S-phase duration (SPD) is extended in all tested carcinoma cell lines. A) Assay to measure Sphase duration from non-synchronized cell cultures: exponentially-growing cells are first labelled with EdU for 30 min, chased with thymidine for various times, and then labelled again with BrdU; nocodazole is added to trap cells in mitosis to prevent a second S phase. B) Bivariate EdU/BrdU flow cytometry profile of RPE1 cells pulsed with EdU, chased for the indicated times, and re-pulsed with BrdU. C) The fraction of EdU + BrdU + double-positive cells decreases linearly with increasing chase times, and becomes null when the chase time corresponds to the length of S phase. Representative example for RPE1-hTERT. D) The duration of S phase was determined for all fourteen carcinoma cell lines (purple) compared to three untransformed lines (black). Values are mean (n=3); statistical significance by Wilcoxon test (P=

Figure 3 :

 3 Figure 3: Single-molecule analysis of DNA replication in fourteen carcinoma cell lines reveals lower DNA synthesis capacity. A) Representative image of combed DNA molecules isolated from exponentially growing cells labelled with 15 min pulses of IdU and CldU, then immuno-detected for IdU (red), CldU (green) and ssDNA (blue). Scale bar 50µm, DNA combing stretching factor: 1µm = 2 kb. B) Schematic explaining how global instant fork density (GIFD), fork velocity (FV) and replication potency (RP) are calculated. Counted forks are indicated by asterisks; elo, elongating fork; ini, initiation event; ter, termination. GIFD is normalized for the S-phase fraction of each cell line, as indicated (see methods). C) GIFD of untransformed (black) and transformed (purple) cell lines. More than 120 Mb of single DNA molecules was measured for each biological experiments (mean, n=3); no statistical significance by Kruskal-Wallis (P=6.646x10 -2 ) between BJ, HMEC-hTERT and RPE1-hTERT; statistical significance by Wilcoxon test (P=8.193x10 -6 ) between untransformed and transformed group. D) Replication potency for

Figure 4 :

 4 Figure 4: Prolonging G1 phase with low doses of Palbociclib restores normal fork density and S-phase duration in the Rb + HCT116 cancer cell line. A) Low dose Palbociclib increases the fraction of cells in G1 only in Rb + cell lines. IdU-DAPI FACS profile of Palbociclib-treated (PD 0.16 µM, 24 hours) RPE1-hTERT and HCT116 (Rb positive) and HeLa (Rb negative) cell lines. Representative example of 3 biological replicates. B) Low dose Palbociclib slows down but does not block the cell cycle. BrdU-DAPI FACS profile showing that RPE1-hTERT and HCT116 cells treated with 0.16 µM Palbociclib are all BrdU + after24 hours (n = 2). C) Single-molecule analysis of replication fork density by DNA combing.GIFD calculated in DMSO or 0.16 µM Palbociclib-treated RPE1-hTERT, HCT116 and HeLa cells (n=4, 2 biological and 2 technical replicates, 82-591 Mb DNA analyzed per sample). Mean value, Wilcoxon test p = 0.6072, 0.2, 0.0003989 for RPE1-hTERT, HCT116 and HeLa cells, respectively. FV were also determined (Sup. Fig.3). D) S-phase duration was determined using the EdU-dT-BrdU assay in DMSO or Palbociclib-treated HCT116 cells (n=6 biological replicates, Wilcoxon-paired test P=0.0313).

Figure 5 :

 5 Figure 5: Mitotic DNA synthesis (MiDAS) and chromosome segregation defects in cancer cell lines untreated with aphidicolin. A) Exponentially-growing RPE1, U2OS, HCT116 and HeLa cells were pulsed for 5 min with 10 µM EdU, fixed and stained for EdU and phosphoH3, then 1200-1500 pH3 positive mitotic cells were scored on a HCS Opera Phenix microscope. B) DAPI images of P-H3 + cells were used to score for mitotic defects (n = 200). C) Mitotic defects were categorized into lagging chromosomes, chromosome bridges, chromosomes alignment failure, uncondensed DNA. D) Example of a HCT116 cell undergoing prophase displaying numerous EdU foci (red). PhosphoS10-H3 (green) is used to identify the 2-4% mitotic cells among several 100 thousands cells. DNA was counterstained with DAPI in blue. Opera Phenix confocal microscope, 63x objective, Scale bar 20 µm.

Figure 6 :

 6 Figure 6: Replication potency and S-phase duration are inversely correlated and sufficient to cluster transformed cells away from non-transformed ones. A) Plot of replication potency versus S-phase duration for the three untransformed and fourteen transformed cell lines. B) Model for the main differences in cell cycle and DNA replication dynamics between normal and cancer cells. Loss of cell cycle regulators gating the G1/S transition in cancer cells impedes origin licensing and causes S-phase extension undetected by checkpoints. Upon mitotic entry, replication forks collapse and DNA synthesis is

Figure 1 .

 1 Figure 1. Al Ahmad Nachar et al

Figure 15 .

 15 Figure 15. Intermitotic times measured in untransformed and cancer cell lines using time-lapse microscopy. Cells were stained with 100 nM SiR-Hoechst stain 4 hours prior to acquisition. Almost all the tested cancer cell lines show an increase in the inter-mitotic time compared to inter-mitotic times calculated in the absence of SiR-Hoechst (See figure 1. in article 1).

  concentration compared to what have been previously published(Lukinavičius et al., 2015) SiR-Hoechst stain would induce cellular toxicity. The inter-mitotic times measured in untransformed and cancer cell lines using SiR-Hoechst are shown to range between 20.7-28.6 hours be longer than the the untransformed (Figure Figure15). Furthermore, these measurements are shown to be longer compared to the inter-mitotic times of the same cell lines measured in the absence of the stain (Figure1 in article 1)..

Figure 16 .

 16 Figure 16. Quantification of DNA content in each cell line by DAPI FACS analysis, relative to diploid RPE1 cells. Example of G1 peak shift in MKN-45, HCC-38, BXPC-3 and 22RV1 compared to the untransformed diploid RPE1-hTERT cell line stained with 0.5µg/ml (1x), 1.5µg/ml (3x) and 4.5µg/ml (9x) DAPI.

  regulated genes of having more DNA replication pause sites in differentiated cells? Fork pausing by definition would introduce a larger temporal difference in the replication of different loci, those replicated before and after the pause. Time is not without influence on chromatin states. Chromatin assembled during early S is preferentially packaged with acetylated histones H3 and H4, while deacetylated histones composing heterochromatin are preferentially assembled in late S[START_REF] Lande-Diner | Shifts in replication timing actively affect histone acetylation during nucleosome reassembly[END_REF]. Increasing the temporal segmentation of chromosome replication could therefore lead to the more dichotomized euor hetero-chromatin of differentiated cells. Conversely the fast proliferation rate of ESC may provide less time for restoring repressive histone marks, thus leading to a generally more open chromatin in ESC.

  Briefly, asynchronous cells were incubated with 25 µM IdU (Sigma I7125) for 15 min, then 200 µM CldU (Sigma C6891) was added for another 15 min, then chased with 200µM thymidine for 1-2 h. Low Melting Agarose plugs were prepared from 25000 cells per 100 µl plug. Cells were digested in plugs with Proteinase K, rinsed 5 times (one hour each) in TNE 50, and melted for 15 min at 65°C and digested overnight at 42°C in MES buffer pH 5.7 with 1 U of β-agarase. This high molecular weight DNA solution was then used for DNA combing on glass coverslips coated with 7-octenyl trichloro silane provided by the Montpellier DNA Combing Facility. DNA combing was performed on a custom made dip coater at 300 µm/sec. Immunodetection of the DNA fibres and of IdU and CldU tracks was performed with specific monoclonal antibodies, mouse anti-ssDNA clone 16-19 (MAB3034, Merck Millipore), mouse

  immersion objective and filters for Alexa 488, 546, and 647 fluorophores. For GIFD analysis, frames with linear, individual DNA fibres were chosen without looking at the IdU and CldU channels, then all three channels were automatically shot with a MetaMorph software journal. For GIFD, coverslips with low DNA density were chosen to avoid ambiguity of signals and DNA overlapping. A minimum of 100Mb of DNA, and at least 30 unambiguous forks per sample, were analysed. The ratio of the number of forks to the total length of measured DNA was calculated and normalized by the fraction of cells in S phase (measured on the same sample by bivariate flow cytometry). Image analysis and DNA fiber, IdU or CldU tracks measurements were performed on custom made software. For IOD measurements, we resampled the DNA fibre distribution to compare similarly-sized samples, since DNA fragmentation can lead to underestimation of IODs in the most fragmented samples. For FV measurements, the track length was divided by the time of pulse.

Figure 1 :

 1 Figure 1: Pluripotent stem cells have a short G1 and high proportion of cells in S phase. A) Mouse ES cells (line JM8.N4) and MEF isolated from 13.5 d embryos cultivated in DMEM-15%FBS-LIF or DMEM-10%FBS medium at 37°C with 21% O2 were labelled with IdU and CldU (15 +15 min), and analysed by bivariate a-BrdU/7-AAD flow cytometry. Gates indicate the fraction of cells in the G1, S and G2+M phases of the cell cycle. The table reports mean values (±SEM) for G1, S and G2+M fractions in mESC (n=5) and MEF (n=6). B) Human ESC (line H9) and iPSC produced by OSKM Sendaï virus infection of human BJ fibroblasts were grown in E8 medium at 37°C with 5% O2, analysed as in A, and compared to normal human diploid fibroblasts (BJ) grown in DMEM-10%FBS. C) mESC were differentiated into embryoid bodies and neural precursor cells and their cell cycle composition analysed as above. D) Cell cycle remodelling during mESC differentiation into NPC. E) gH2AX and H2AX levels were quantitated by western blot fluorescence in mESC, EB3 and EB6, along with UV-irradiated mESC (254 nm, 20 J/m 2 ). Ratio of gH2AX over H2AX are indicated below.

Figure 2 :

 2 Figure 2: Single-molecule analysis of DNA replication by DNA combing. A) Representative image of combed DNA molecules after IdU-CldU double pulse (15+15 min) and detection for DNA (blue), IdU (red) and CldU (green). Red-Green (RG) tracks indicate ongoing replication forks and their directionality (P1, IdU pulse; P2, CldU pulse). Replication origins are assumed to be at the centre of red tracks surrounded by green (GRG) or sole green (G) tracks. IOD, inter-origin distance. Global fork density is obtained by dividing the total number of RG tracks by the total length of DNA fibres. B) Fork velocities calculated by dividing the length of the CldU track in ongoing forks by the duration of the 2 nd pulse (15 min). Mean FV (±SEM) are 1.72 (±0.03) kb/min for mESC (n=198), 2.0 (±0.07) kb/min for MEF (n=95),

Figure 3 :

 3 Figure 3: S-phase duration in mESC and MEF using split EdU-BrdU dual pulse labelling. A) Principle of the method. Asynchronously growing cell populations are labelled with first EdU (30 min) then BrdU (30 min), or after an intervening thymidine (dT) chase of variable length. Nocodazole (Noc) is added to prevent cells of entering a subsequent cycle. B) Examples of EdU/BrdU FACS dot-plot of MEF indicating the population of double-positive cells (red square) in cultures labelled with EdU and BrdU (30 min each) or after separating the two pulses by 4h and 8h. C) Decrease in double-positive cells in mESC (blue) and MEF (orange) after increased thymidine chase times between the two pulses. Linear regression defines the x-axis intersection point (arrows) as the time required for the earliest S-phase cells to complete DNA replication (S-phase duration). Mean (±SEM) S-phase duration is 8.47 h (± 0.09, n = 3) for mESC and 8.50 h (± 0.22, n=4) for MEF.

Figure 4 :

 4 Figure 4: Recruitment of additional replication origins in MEF compared to mESC. A) Model representing a chromosomal regions containing two replication origins (Ori1,2) and three dormant origins (dOri). Green arrows represent active forks. Red rectangles indicate fork slowing and pausing in MEF cells, leading to the activation of dormant origins and increase in GIFD. The number of forks detected by DNA combing in mESC and MEF is indicated on the right. IODs measured at consecutive times of S phase are depicted below. B) Replication tract size measured on DNA fibres larger than 500 kb after a CldU pulse of 60 or 120 min in mESC and MEF. Mean size (red). C) Distance between two replication tracts on the same fibre (gap size) in the same conditions. D) Cells were labelled with CldU for 30, 60, 120 or 180 min and replication tract size analysed on DNA fibres larger than 300 kb red, mean). E)Fraction of DNA fibres larger than 400 kb that are more than 70% replicated in mESC and MEF after increasing CldU pulse times.

Figure 5 :

 5 Figure 5: Replication dynamics by DNA combing during the differentiation of mESC into NPC. mESC were differentiated and analysed by DNA combing at days three (EB3), six (EB6) and nine (EB9) of embryoid body formation, along with NSCs isolated from embryo's brain (mNSC) and neural progenitor cells (mNPC) differentiated in vitro. A) Mean (±SEM) inter-origin distances for mESC (152.3 kb ±18.3, n=19), EB3 (131.0 kb ±12.1, n=33), EB6 (103.4 kb ±12.5, n=23), EB9 (128.8 kb ±14.1, n=27) and mNPC (117.7 kb ± 8.5, n=12. B) DNA fibre length distribution for the fibres used in A. C) Mean (±SEM) fork velocity (FV) for mESC (1.84 kb/min ±0.04, n=124), EB3 (1.57 kb/min ±0.05, n=106), EB6 (1.99 kb/min

Figure 18 .

 18 Figure 18. Detecting fork pausing events using single-molecule analysis by DNA combing . A) Pyridostatin treated MEFs show no significant pausing events compared to the untreated. 65-103 green tracts were

Figure 19 .

 19 Figure 19. Inter-origins distances measured in mouse ESC and MEFs using single molecule-analysis by DNA combing (Bialic et al., in preparation).

  , a). Interestingly, this time the IODs in MEFs were 58% higher (median=166 vs 105 kb, respectively) compared to mouse ESCs. The IODs measured in mouse ESCs are similar to IODs analyzed previously on shorter fibers whereas the IODs measured in MEFs were increased to 166 kb (Figure 19, Figure 20 b).

Figure 20 .

 20 Figure 20. Single molecule analysis of IOD display significant decrease in IOD in mouse ESC compared to MEFs. A) Mann Whitney P >0.01 Median with interquartile range for fiber length between MEFs and mouse ESC. B) 45-68 IODs on 500-1200 kb fibers were analyzed. Mann Whitney P <0.01 Median with interquartile range for IODs between MEFs and mouse ESC.

Figure 21 .

 21 Figure 21. Single-molecule analysis of MEFs treated with 20 nM TSA for 1, 4.5 and 10 hours shows a decrease of GIFD in TSA treated samples. 80-270 Mb of total DNA have been analyzed per sample.

Figure 1 ,

 1 Bialic et al). The results showed a constant GIFD throughout the different sub-phases of S phase implying that the same number of origins is used in each of these sub-phases. In order to get insight whether MEFs might utilize same number of origins throughout the different sub-phase of S phase, MEFs were also pulsed labeled with IdU/CldU for 15 min each followed by thymidine chase then cells were fixed and sorted according to DNA content into early, mid and late S phase(Figure 22.)

Figure 22 .

 22 Figure 22. Sorting of S phase fraction into early, mid and late S phase based on DNA content on MEFs using Flow Cytometry.

Figure 23 )

 23 Figure 23). Another two different biological experiments were done on MEFs where I obtained

Figure 23 .

 23 Figure 23. GIFD calculated in sorted MEFs based on DNA content into S1, S2 and S3 sub-phases of S phase. A) represents 2 technical experiments on a single biological experiment. B) and C) each represents an independent biological experiment.

  prolongation itself or to the stress associated with cell cycle alteration, as several stresses have been shown to trigger ESC differentiation. If this correlation was true, one should take into consideration this fact while targeting some cancers with CDK4/6 inhibitors. Our results provide evidence that treating RB-positive cancer cell lines with a low dose of palbociclib would rescue origin density and S-phase duration through prolonging G1 phase, which might improve licensing. However, this effect on prolonging G1 phase might induce differentiation of adult stem cells (if they behave as ESCs) around the tissue treated with palbociclib, potentially exhausting the stem cell compartment. One should be cautious concerning palbociclib targets. Whether the importance of origin licensing is more evident in stem cells than differentiated cells was addressed in 2015. Knocking out both Mcm3 alleles in MEFs was compatible with DNA replication, but it resulted in accumulation of g-H2AX foci, indicating DNA damage. However, knocking out both Mcm3 alleles in mouse embryos resulted in pre-implantation lethality. Knocking out a single Mcm3 allele in mouse embryos was correlated with increased tumors frequencies and invasiveness. The authors assessed hematopoietic tumors in mid-late gestation embryos and revealed the presence of immature erythroblasts with micronuclei indicating genomic instability. Furthermore, erythroid lineage cells derived from Mcm3-deficient embryos lost their ability to mature when transplanted in irradiated recipient mice. This indicates

(

  MGS) display inherited mutations in one of the licensing components. However, these patients have decent life expectancy and do not display increased risk of cancer compared to nonpatients. The reason that might explain this might be prenatal lethality of mutants with strong phenotypes similar to the Mcm3 double knock out mentioned in Alvarez and colleagues work above. However, patients with weaker phenotypes (hypomorphs) would survive and display developmental abnormalities. In addition, MGS being a very rare disease makes it more difficult to have enough cases to check for susceptibility to cancerogenesis.Cancer cells are renowned for their unlimited proliferative capacity. In order to give more insights about the very first line mechanisms that pushes cancer cells to go awry we addressed first how alteration in DNA replication dynamics induces genome instability. We used direct methods to study S-phase dynamics at the single molecule and the single cell levels in both untransformed and cancer cell lines. Most of cancer field research focuses on rereplication and overreplication as mechanisms to explain replication defects in cancer. Very few studies shed the light on the importance of full origin licensing in maintaining genome stability in mammalian cells. We provided evidence that cancer cell lines utilize low number of origins to replicate the genome. As a consequence, SPD and doubling time are extended which goes against to what is published in most of the literature. Despite the extension in the SPD, cancer cell lines fail to finish replication and enter mitosis with under-replicated regions. These regions finish replication by a recent published mechanism, MiDAS. We proposed that under-licensing in G1 is the main mechanism where early S phase entry causes the cells to escape the second wave of origins licensing. As a result, cancer cell lines enter S phase with few licensed origins thus it would utilize fewer number of origins and induce the phenotypes mentioned above. In order to rescue origin licensing, we treated RB-positive HCT116 colon cancer cell line with low dose of palbociclib to prolong G1 giving more chances for the cells to benefit from the second wave of origin licensing that occurs at the end of G1 phase. The results showed an improvement in both GIFD and SPD confirming our hypothesis. This would allow us to highlight on the mechanism by which low dose of palbociclib is used to treat cancers.Similarly, embryonic stem cells are characterized by their fast proliferation and short cell cycle. Understanding how mouse ESC proliferate fast but still managing to sufficiently replicate their genome. It has been already shown that ESCs are characterized by having short G1 and short cell cycle. However, they also have increased levels of chromatin-bound CDT1 and CDC6 and fast MCM loading to the chromatin which explains how these cells maintain genome stability despite having short G1. Again, this highlights on the importance of origin licensing in maintaining proper replication of the genome. Thus, we also studied S-phase dynamics in both mouse ESCs and MEFs and followed the changes upon differentiation. Our previous data showed that mouse ESCs spend as long as MEFs to replicate their genome while mouse ESCs use 2-fold less number of origins when compared to MEFs. The difference is mainly in the distribution and the replication timing of origins. We were able to exclude the hypothesis that MEFs encounter more pausing events resulting in the activation of dormant origins which would increase the GIFD. Studying the physiology of mouse ESCs permits us to compare it to the replication dynamics of cancer cell lines and probably find an efficient treatment.La plupart des tumeurs solides, sinon toutes, présentent une altération génétique ou épigénétique d'un ou plusieurs régulateurs de la transition G1/S du cycle cellulaire et présentent une instabilité du génome. Cependant, on ne sait pas encore précisément comment ces deux événements sont liés, mais l'altération du phénomène de licensing est une hypothèse intéressante.

(

  RP=GIFD*FV) pour mesurer cette compensation. La RP a été calculée entre 1,71 et 2,07 pour les cellules non transformées et entre 0,4 et 1,3 pour les lignées de cellules cancéreuses. Cela indique que l'efficacité de la réplication est fortement diminuée dans les cellules cancéreuses, et que leur phase S plus longue est une conséquence de la difficulté à répliquer le génome.Les conséquences de l'extension de la phase S sur le cycle cellulaire a été analysée par time-lapse microscopy. Le temps de doublement a été calculé pour huit lignées cellulaires cancéreuses. Il a été démontré récemment que le stress de réplication induit par de faibles doses d'aphidicoline déclenche une réplication tardive et la résolution des intermédiaires de réplication seulement en la mitose, accompagné d'une synthèse d'ADN tardive (MiDAS)(Bhowmick et al., 2016b; Minocherhomji et al., 2015). La pénétrance de la MiDAS spontanée a également été évaluée dans des cellules normales et cancéreuses en l'absence d'aphidicoline, en utilisant l'acquisition et l'analyse automatisées d'images sur un microscope confocal Opera Phenix. Entre 1200 et 1500 cellules mitotiques ont été analysées pour chaque lignée cellulaire. Nos résultats suggèrent que les cellules cancéreuses présentant une réplication tardive subissent une recombinaison mitotique de l'ADN à une fréquence plus élevée, ce qui est probablement responsable de leur instabilité chromosomique. Les défauts mitotiques et leurs fréquences ont également été analysés dans les lignées cellulaires testées et les résultats étaient conformes à ceux que nous avions obtenu avec le MiDAS. Les trois lignées de cellules cancéreuses testées présentaient une fréquence plus élevée de défauts mitotiques se manifestant par un défaut d'alignement des chromosomes sur la plaque équatoriale, des chromosomes en retard et des ponts chromosomiques.Des données antérieures du laboratoire sur la levure sic1∆ ou les mutants cdc6-1 et sur les MEF Rb TKO (Triple KnockOut) murins laissent fortement présager que les défauts de licensing des origines pourraient être la cause principale de la faible GIFD et de la longue phase S des cellules cancéreuses. Afin de tester cette hypothèse, de faibles doses de Palbociclib, un inhibiteur spécifique de CycD-Cdk4/6(Fry et al., 2004), ont été utilisées pour restaurer une phase G1 plus longue dans les cellules cancéreuses afin d'améliorer l'origin licensing et de corriger la durée de la phase S. Le traitement de la lignée cellulaire cancéreuse HCT116 (RB-positive) et de la RPE1-hTERT (non transformée) avec une faible dose de palbociclib a augmenté la GIFD par rapport aux témoins non traités. Toutefois, cet effet n'a pas été observé lorsque la lignée cellulaire HeLa (RB négative) a été traitée avec une faible dose de palbociclib. Nous avons ensuite testé si le fait de restaurer la GIFD dans la lignée cellulaire cancéreuse HCT116 permettrait d'empêcher

4)

  Déterminer les conséquences de la diminution du nombre d'origines de réplication utilisées et de l'extension de la phase S sur la réplication mitotique, la ségrégation chromosomique et l'instabilité du génome dans les lignées de cellules cancéreuses en utilisant l'incorporation d'EdU sur les chromosomes mitotiques et en déterminant les défauts mitotiques associés à ces phénotypes.RésultatsI)Une faible puissance réplicative induit une extension de la phase S dans plusieurs lignées de cellules cancéreuses testées (objectifs 1-2) Une phase G1 tronquée et la présence d'un stress de réplication étant des caractéristiques fréquentes des cellules cancéreuses, on s'attend à ce qu'elles passent plus de temps en phase S afin de répliquer le génome. Cependant, cette idée n'a jamais été validée de manière approfondie. En utilisant une méthode nouvellement développée au laboratoire et basée sur l'incorporation séquentielle d'un double pulse d'EdU et de BrdU, j'ai mesuré la durée de la phase S dans quatorze lignées de cellules cancéreuses différentes et bien caractérisées provenant de neuf tissus épithéliaux (issues du Broad Institute, NCI et généreusement fournies par le SIRIC Montpellier), en comparaison avec des lignées non transformées BJ, HMEC-hTERT et RPE1-hTERT. Les mesures de SPD ont montré que toutes les lignées de cellules tumorales testées jusqu'à présent présentent une phase S plus longue (entre 10 et 29 heures), bien au-delà des 7-9 heures déterminées pour les cellules contrôles non transformées. Afin de tester si cette extension de la phase S est due à l'utilisation de moins d'origines de réplication, j'ai mesuré la GIFD en utilisant la technique de peignage moléculaire de l'ADN. Les résultats montrent une tendance générale pour la plupart des cellules cancéreuses indiquant une GIFD inférieure à celle des contrôles normaux. Pour vérifier si les cellules cancéreuses compensent la diminution de la GIFD, la vitesse des fourches (FV) a été mesurée. La GIFD des lignées de cellules cancéreuses se situait entre 0,49 et 1,24, tandis que celle des témoins non transformés se situait entre 1,19 et 1,91. Quant aux vitesses de fourches, elles sont comprises entre 0,6 et 1,52 pour les lignées de cellules cancéreuses et entre 1 et 1,55 pour les lignées non transformées. La puissance réplicative a été calculée comme un facteur de la GIFD et de la FV. Un facteur de 1,71-2,07 a été calculé pour les contrôles, en revanche il est de 0,4-1,3 pour les lignées de cellules cancéreuses, indiquant une capacité moins grande à répliquer l'ADN. J'ai également montré une corrélation linéaire entre la puissance réplicative et la SPD dans la plupart des lignées de cellules cancéreuses. Les cellules non transformées ont les durées de phase S les plus courtes et les puissances réplicatives les plus hautes, tandis que les lignées de cellules cancéreuses ayant les plus longues SPD ont la plus faible puissance réplicative. J'ai également confirmé que notre mesure de la puissance réplicative est indépendante des différences de quantité d'ADN entre ces lignées de cellules cancéreuses. II) Conséquences d'une faible GIFD et d'une phase S prolongée pour la mitose et le cycle cellulaire (objectif 3)

  Contrairement aux études mentionnées, nous évaluons la synthèse endogène d'ADN mitotique en absence d'aphidicoline, qui ralentit la réplication de l'ADN. J'ai utilisé le criblage automatisé avec Opera Phenix pour tester la présence de MiDAS dans les lignées HCT116, HeLa et U2OS et les comparer à la lignée cellulaire RPE1-hTERT non transformée. Entre 1200 et 1500 cellules mitotiques ont été analysées pour l'incorporation d'EdU dans chacune des lignées cellulaires testées. Les résultats montrent que le pourcentage de cellules mitotiques incorporant de l'EdU est de 5,5%, 1,14% et 0,7% dans U2OS, HCT116 et HeLa respectivement, comparé à 0,4% dans la lignée cellulaire RPE1-hTERT. III) Restaurer le faible pouvoir de réplication et la longue SPD par une prolongation de la phase G1 dans les lignées cellulaires cancéreuses (objectif 4) Sur la base de nos résultats dans les modèles de levure et de TKO, nous avons fait l'hypothèse que cette réduction de la GIFD dans les cellules cancéreuses est due à un licensing déficient pendant la phase G1. PD 0332991 ou Palbociclib, inhibiteur spécifique du cdk4/6, inhibe la phosphorylation de RB et entraîne l'arrêt des cellules en G1. Nous émettons l'hypothèse que prolonger G1 donnerait aux cellules plus de temps pour mettre en place le licensing, ce qui pourrait rétablir la puissance de réplication. Nous avons traité les HCT116 (RB positives), les HeLa (RB négatives) et les RPE1-hTERT (non transformées) avec une faible dose de palbociclib. Les résultats montrent une augmentation de la GIFD dans les lignées cellulaires des HCT116 et RPE1-hTERT. Comme prévu, aucun changement n'a été observé dans la lignée cellulaire HeLa traitée par palbociclib. Cela a confirmé que l'inhibition de la phosphorylation des RB restaure la densité de fourches. Nous avons également vérifié si ce sauvetage de la GIFD permettrait de rétablir la durée normale de la phase S. La lignée cellulaire HCT116 a été traitée avec la même dose de palbociclib puis les cellules ont été testées pour la SPD en utilisant le protocole mentionné ci-dessus. Les résultats ont montré que le rétablissement de la GIFD dans les HCT116 entraîne une légère diminution mais non significative de la SPD. Conclusion Dans ce travail, nous mettons en évidence un paradoxe intéressant à propos du cancer. Nous avons développé des méthodes directes molécule-unique et cellule-unique pour étudier la dynamique de la phase S dans les lignées cellulaires non transformées et cancéreuses. Nous avons montré que les cellules cancéreuses utilisent moins d'origines pour répliquer le génome. En conséquence, la SPD et le temps de doublement sont prolongés, ce qui va à l'encontre de ce que l'on pense habituellement des cellules cancéreuses. Malgré l'extension de la phase S, les cellules cancéreuses ne parviennent pas à terminer leur réplication et entrent en mitose avec des régions sous-répliquées. Ces régions finissent leur réplication par un mécanisme récemment publié appelé MiDAS. Nous proposons que l'under-licensing en G1 soit le principal mécanisme induisant les phénotypes mentionnés ci-dessus. Afin de restaurer le licensing des origines, nous avons traité la lignée cellulaire HCT116 RB-positive avec une faible dose de palbociclib pour prolonger la phase G1. Les résultats ont montré une amélioration à la fois du GIFD et du SPD, confirmant notre hypothèse proposant que la briéveté de G1 empêche un licensing suffisant dans les cellules cancéreuses. Ce travail permet donc de comprendre un mécanisme par lequel les cellules cancéreuses peuvent acquérir une instabilité génétique, et propose une stratégie thérapeutique basée sur de faibles doses de palbociclib. J'ai également contribué à une étude initiée par une ancienne doctorante de mon laboratoire qui travaillait auparavant sur la dynamique de réplication dans les cellules souches pluripotentes. Les cellules souches embryonnaires (ESC) sont caractérisées par un cycle cellulaire particulier où elles passent moins de temps dans les phases G1 et G2 que les cellules somatiques. Il a été démontré que le fait d'avoir des G1 courtes était important pour le maintien de la pluripotence. La modification de la cinétique de G1 induit un allongement de G1 qui s'accompagne d'une différenciation

3 ) 1 ) 2 )

 312 . Cependant, le lien entre le remodelage du cycle cellulaire, la régulation de la réplication de l'ADN et le développement doit être étudié plus profondément. L'objectif principal de ce projet est d'obtenir plus d'informations sur la régulation de la dynamique de réplication des ESC par rapport à leur état différencié.Les principaux résultats de ce projet au moment de mon intervention étaient les suivants :L'analyse single-molecule par peignage de l'ADN a été utilisée pour vérifier si une dynamique de réplication particulière est corrélée à la pluripotence des cellules souches. Les résultats ont montré peu de changement dans les vitesses de fourches et les distances interorigines (IOD) entre les ESC murines et les fibroblastes embryonnaires murins (MEF) différenciés. La première approche consiste à optimiser le protocole de peignage de l'ADN en augmentant la longueur des fibres, ce qui pourrait permettre de mesurer les IOD entre les clusters. La seconde consiste à diminuer la compaction de la chromatine dans les MEF, ce qui pourrait permettre une distribution plus uniforme des origines. L'établissement d'une distribution uniforme des origines dans les MEF devrait réduire le nombre d'origines nécessaires dans les MEF pour répliquer le génome. Une autre possibilité expliquant l'augmentation de la GIFD dans les MEF pourrait être attribuée à une différence dans la distribution des origines utilisées au début, au milieu et à la fin de la phase S. Le fait d'avoir plus d'origines en phase S précoce et moins d'origines en phase S tardive permettrait également d'activer les origines dormantes si elles étaient licensed. Dans le cas contraire, les cellules devraient passer plus de temps à répliquer les régions tardives, malgré l'augmentation antérieure du nombre d'origines utilisées.. Afin de tester cette hypothèse, les MEF ont été triées en cellules de phase S précoce, moyenne et tardive sur la base de leur profil de contenu en ADN. La GIFD a été déterminée et comparée dans chacune des fractions mentionnées. des événements de la pause de la fourchette dans MEFs et mESCs (Objectif Nous avons calculé le rapport entre la deuxième impulsion et la première, où la durée des deux impulsions était de 15 minutes. De cette façon, nous devrions être en mesure de détecter les pauses pendant la deuxième impulsion. Si des pauses étaient présentes lors de la seconde impulsion, le rapport devrait être inférieur à 1. Cependant, les MEF et les mESC ont montré des rapports autour de 1 indiquant que la pause n'a pas été détectée pendant 15 minutes d'impulsion. Le traitement des MEF avec 500 nM de pyridostatine, un stabilisateur des Gquadruplexes, pendant 2 à 3 heures n'a pas non plus permis de détecter une pause dans les MEFs. II) Etude de la distribution des origines dans le génome des MEF et des mESC (Objectif En mesurant les IOD, nos résultats précédents n'ont montré aucune différence significative des IOD entre les MEF et les mESC. Cependant, il a été précédemment démontré que la technique de peignage de l'ADN est limitée par le fait que les fibres d'ADN se fragmentent pendant la préparation. Nous avons décidé d'améliorer notre protocole de peignage de l'ADN afin de pouvoir conserver de longues fibres d'ADN dans notre analyse. Le peignage de l'ADN a été effectué à la fois sur les MEF et les mESC. Les IOD ont été mesurées dans les deux types de cellules. Les IOD dans les MEF étaient 58% plus élevées (médiane=166 vs 105 kb, respectivement) par rapport aux mESC. Les IOD mesurées dans les mESC sont similaires aux IOD analysées précédemment sur des fibres plus courtes alors que les IOD mesurées dans les MEF ont augmenté à 166 kb. Cela correspond à un scénario dans lequel l'hétérogénéité de la distribution des origines dans les MEF (et probablement d'autres cellules différenciées) fait que les fourches provenant de différents clusters parcourent une plus grande distance, ce qui donne le temps aux origines tardives d'être activées, si elles ont été licensed. Une dose de 20 nM de Trichostatin A, un inhibiteur des histones désacétylases, devrait ouvrir la chromatine et permettre une distribution plus homogène des origines. La TSA a été utilisée pour traiter les MEF pendant 1, 4,5 ou 10 heures, suivi d'un double pulse a montré une diminution de 1,4 fourchettes/Mb dans le contrôle à 0,9, 1 et 0,7 fourchettes/Mb dans le GIFD respectivement. Cela confirme notre hypothèse selon laquelle les différences de GIFD entre les MEF et les mESC pourrait reposer sur une différence dans la distribution des origines. III) Le proto la GIFD au cours de la phase S dans les MEF (Objectif 3). Nous avons pensé que la régulation temporelle pourrait jouer un rôle, de même que le fait que les mESC ont une chromatine plus ouverte que les cellules différenciées. Ainsi, les mESC pourraient avoir des régions de réplication plus précoces que les MEF, ce qui leur donnerait l'avantage de disposer de plus de temps pour répliquer le génome. Les mESC et les MEF ont été marquées, fixées et triées par cytométrie selon leur contenu en ADN pour obtenir les phases S précoce, moyenne et tardive . Les résultats ont montré une GIFD constante tout au long des différentes sous-phases de la phase S, ce qui implique que le même nombre d'origines est utilisé dans chacune de ces sous-phases. Les résultats montrent que, contrairement aux mESC, les MEF présentent une variation du nombre d'origines actives au cours des différentes sous-phases, avec 0,99 en S1, 1,28 en S2 puis diminue fortement pour atteindre 0,54 en S3. Deux autres expériences biologiques différentes ont été réalisées sur des MEF où j'ai obtenu la même tendance ; cependant, je n'ai pas réussi à reproduire le même nombre d'origines utilisées dans chacune des sous-phases. Malgré cet échec, les différences entre les nombres d'origines utilisées dans les différentes sous-phases impliquent que les régions qui se répliquent tardivement dans les MEF utilisent moins d'origine, ce qui ralentit le déroulement de la phase S. Conclusion Dans ce travail, nous fournissons des arguments expérimentaux montrant une différence dans la dynamique de réplication entre les mESC et des cellules plus différenciées (MEF). Nous avons montré que les mESC prennent autant de temps que les MEF pour répliquer complètement 239 leur génome, alors qu'elles ont moins d'origines actives. Ce paradoxe peut être expliqué par des différences dans la distribution et le temps de réplication des origines. Comprendre comment les mESC prolifèrent rapidement tout en parvenant à répliquer complètement leur génome est un enjeu important, tant pour la biologie des cellules souches que pour celle d'autres modèles hyperprolifératifs (cancer).

1.2. Untransformed Cell Lines

	1. Project 1:
	1.1. Cell Culture
	1.1.1. Cancer Cell Lines
	Eleven cancer cell lines, HCC-1937, HCC-38, Colo205, Colo320, 786-O, BXPC-3,
	ASPC-1, PC-3, 22RV1, COV318 and MKN-45, were provided by the Montpellier Integrated
	Cancer Center (SIRIC Montpellier). Among these cells, the Cov318 was cultured in
	DMEM/Glutamax-I medium (31966-021, Life Technologies) supplemented with 10% fetal
	bovine serum (CVFSVF00-01, Eurobio) and 100U/ml penicillin-streptomycin (151-40-122,
	Primary human fibroblasts (BJs) were cultured in DMEM/Glutamax-I medium (31966-021,
	Life Technologies) supplemented with 10% fetal bovine serum (Eurobio) and 100 U/mL
	penicillin-streptomycin (151-40-122, Thermo Fisher), 1 mM pyruvate (31966-047, Thermo

Thermo Fisher) whereas the rest were cultured in RPMI (R2405, Sigma) supplemented with 10% fetal bovine serum (CVFSVF00-01, Eurobio) and 100 U/mL penicillin-streptomycin (151-40-122, Thermo Fisher). HeLa, U2OS and HCT116 were cultured in DMEM/Glutamax-I medium (31966-021, Life Technologies) supplemented with 10% Hyclone fetal bovine serum (SV30160.03, GE Health Care Life Sciences) and 100 U/mL penicillin-streptomycin

 Thermo Fisher)

. Cells were maintained in 37°C humid incubator in the presence of 5% CO2 and 21% O2.

1.

  -growing cells are pulsed sequentially with EdU and BrdU at one time point. For the other time points, the EdU and BrdU pulses are interrupted by increasing times of thymidine chase. When EdU and BrdU are pulsed sequentially, most of the cells in S phase incorporate both nucleotides. As the time of thymidine chase increases, the fraction of cells incorporating both analogues decreases (due to cells entering or exiting S phase) until it reaches zero. This time when EdU-positive cells stop incorporating BrdU corresponds to the duration of

	incubated in 0.5 µg/mL DAPI (Sigma) solution containing 200 µg/mL RNase A overnight at 4
	ºC.
	S phase.
	1.6.3.2. Double Labeling in Cell Culture
	Asynchronous cells growing exponentially in culture were pulsed with 10 µM EdU
	(Invitrogen) for 30 minutes. For the 30 minutes time point, EdU-containing medium was
	removed and replaced by medium containing 100 µM BrdU (Sigma) and 100 ng/ml nocodazole
	it reaction containing 2 mM CuSO4 (Sigma), 10 mM ascorbic acid (Sigma) and 0.02 mM di-
	sulfocyanine 5 azide (FP-1JV6320, Interchim) for 45 minutes at 37 ºC. Cells were washed with
	PBS 1X, 1% BSA, 0.5% Tween 20 for 5 minutes, then incubated in mouse anti-BrdU solution
	(MoBu-1, 11-286-C100, Exbio, 1/1000) diluted in PBS 1X, 1% BSA, 0.5% Tween 20 for 1 hour
	at 37 ºC. Cells were washed with PBS 1X then incubated in Alexa 488 Fab goat anti-mouse
	(A11017, ThermoFisher, 1/1000) for 1 hour at 37 ºC. Cells were washed with PBS 1X then

(Sigma) for 30 minutes. For the other time points, EdU-containing medium was removed and replaced by medium containing thymidine …… BrdU and Nocodazole for 30 minutes. Cells were trypsinized and fixed with 96% cold ethanol.

1.6.3.3. Immunodetection

Cells were washed twice with an equal volume of PBS. Cells were treated with 2N HCl, 0.5% Triton X-100 for 30 minutes to denature DNA allowing the access of anti-BrdU antibodies to their epitope. Cells were washed with PBS, neutralized with sodium borate buffer pH8.5. Cells were blocked in PBS 1X, 1% BSA, 0.5% Tween 20 for 5 minutes. Cells were incubated in Click-

table III .

 III Summary of forks analyzed for fork velocities in untransformed and transformed cell lines.

	A A				HCT116-DMSO
		0.5 h		2 h		4 h		6 h
	BrdU	Q1 4.91%	Q2 47.3%	Q1 11.2%	Q2 43.3%	Q1 22.4%	Q2 34.4%	Q1 28.4%	Q2 27.5%
		Q4	Q3	Q4	Q3	Q4	Q3	Q4	Q3
		46.8%	0.482%	38.9%	6.34%	26.1%	16.7%	18.7%	24.6%
		8 h	EdU Ed	9 h		+
		Q1 32.4%	Q2 18.7%	Q1 31.9%	Q2 15.9%	EdU
						/
						+
						BrdU
		Q4 17.0%	Q3 31.6%	Q4 15.7%	Q3 36.3%	Ratio EdU +
							Time between pulses (h)
					HCT116-0.16 µM PD
		0.5 h		2 h		4 h		6 h
	BrdU
			EdU
		8		9 h

h Q1 2.83% Q2 35.3% Q3 0.793% Q4 61.2% Q1 11.9% Q2 29.8 % Q3 6.21% Q4 52% Q1 39.8% Q2 21.9% Q3 13.4% Q4 25% Q1 43.6% Q2 16% Q3 19.2% Q4 20.8%

	Q1 50.1%	Q2 9.4%	Q1 47.3%	Q2 6.51%
	Q4 14.7%	Q3 25.6%	Q4 17.5%	Q3 28.6%

Table I :

 I Whole-genome extrapolation of replication parameters measured by DNA combing

	Figure 6: Increased GIFD in MEF at different passages after derivation from embryos. A) Global instant
	Bi-colour for density (GIFD) was measured in NIH 3T3 cells and compared to mESC. Mean (±SEM) is 1.13 Mean GIFD Genome size Total number Mean FV DNA synth./min Time needed to replicate forks (forks/Mb) (Mb, 2n) of forks/pulse (kb/min) (Mb) full genome
	fork/Mb (±0.01, n=3, total 330 Mb). B) GIFD increases during in vitro culture. GIFD was measured in mESC 0.98 5600 5488 1.72 9.44 593 min (9.89 h)
	MEF at passages p1 (1.05 ±0.1 fork/Mb, n=3), p3 (1.22 ±0.09 fork/Mb, n=5), p5 (1.73 ±0.45 fork/Mb, MEF 1.93 5600 10808 2.00 21.62 259 min (4.32 h)
	n=3) and p7 (2.41 ±0.47 fork/Mb, n=4). C) S-phase duration measured by EdU-BrdU dual pulse chase is
	All forks Mean GIFD 8.47 h (±0.11) for mESC (n=3) and 10.52 h (±0.73) for NIH 3T3 (n=3). D) The fraction of cells in S Genome size Total number Mean FV DNA synth./min Time needed to replicate (forks/Mb) (Mb, 2n) of forks/pulse (kb/min) (Mb) full genome
	mESC phase decreases during passages of MEF: p1, 32.2 % ± 0.85 (n=3); p3, 30.7 % ± 2.35 (n=4); p5, 21.3 % ± 1.59 5600 8904 1.24 11.04 507 min (8.45 h) MEF 2.88 5600 16128 1.44 23.22 241 min (4.02 h)
	0.7 (n=4); p7, 16.2 % ± 1.2 (n=4). E) S-phase duration in MEF at different passages after in vitro culture.
	±0.04, n=124), EB3 (1.57 kb/min ±0.05, n=106), EB6 (1.99 kb/min
	±0.06, n=55), EB9 (1.73 kb/min ±0.06, n=67), mNPC (2.02 kb/min ±0.06, n=120) and mNSC (1.52 kb/min
	±0.06, n=72). Mann-Whitney test **** p<0.0001, ** p<0.01. D) Median (SD) global instant fork density
	is 1.90 fork/Mb (0.23) for MEF (n=3, 366.7 Mb), 1.03 fork/Mb (0.23) for mESC (n=4, 532.3 Mb), 1.13
	fork/Mb (0.28) for EB3 (n=3, 383.9 Mb), 0.69 fork/Mb (0.21) for EB6 (n=4, 726.9 Mb), 0.54 fork/Mb
	(0.21) for EB9 (n=3, 493.4 Mb), 1.43 fork/Mb (0.30) for mNPC (n=3, 323.7 Mb) and 1.34 fork/Mb (0.11)
	for mNSC (n=2, 308.8 Mb). Unpaired t test.

Suppl. figure1. Al Ahmad Nachar et al

Acknowledgements Acknowledgments

We thank A. Trullo for developing software to help tracking cells analyzed by time-lapse microscopy, the Montpellier Ressources Imagerie (MRI) and DNA combing (MDC) facilities for precious help, Vjekoslav Dulic for providing a first set of live cell imaging inter-mitotic times, and all lab members for useful suggestions and critical reading of the manuscript. We are grateful for Thierry Gostan for all of the statistical analysis. We thank IGMM and CRBM facilities and personel for sharing help and reagents. We are grateful to the IRCM Cell culture unit of Montpellier SIRIC (INCa-DGOS-Inserm 6045) for providing us with all of the validated cancer cell lines. BA was supported by a PhD fellowship from La Ligue Nationale Contre le Cancer. ES lab is funded by grants from ANR, ARC, SIRIC Montpellier and the Region Occitanie with help from EU-FEDER grants. Acknowledgements We thank J. de Vos, P. Nararro and P. Pasero for useful comments and support during this work, S. Khochbin for Atad2 KD cells, the Montpellier RIO Imaging (MRI) and DNA combing (MDC) facilities for precious help, as well as D. Gilbert, M. Lagha and B. Le Tallec for critical reading of the manuscript. MB was supported by PhD fellowships from the French Ministry of Education and Research (MESR) and by the Ligue contre le Cancer. The labs of JML and ES are funded by grants from xxx, yyy (JML), Agence Nationale de la Recherche and Cancéropole Grand Sud-Ouest (ES).

restore a normal S phase to pre-cancerous cells, thus presumably also decreasing their genome instability.

It is likely that these low doses of Palbociclib will not generate resistance, and we therefore suggest this new therapeutic strategy to be tested.

LIST of ABBREVIATIONS

53BP1

p53 binding protein1 7-AAD 7-aminoactinomycin D

Abstract

Why cancer cells have unstable genomes is still poorly understood. Seminal work has shown that precocious CDK activation in G1 and reduced replication origin licensing cause S-phase extension and gross chromosome rearrangements in yeast. Since most cancer cells have alterations in various G1/S cell cycle regulators that could hamper origin licensing, we analysed chromosome replication dynamics in fourteen human epithelial cancer cell lines using newly developed techniques. Our results show that cancer cells take longer to replicate their genome (10-29h) compared to untransformed cells (7-9h), which is mainly due to a lower density of replication forks measured on single DNA molecules. Combining fork density and fork velocity, we define replication potency as the amount of DNA a cell is capable of synthesizing per minute, and found that it was lower for all tested cancer cell lines. High-content microscopy revealed mitotic DNA synthesis and chromosome segregation defects in cancer cells not treated with replication drugs, likely a consequence of mitotic entry with under-replicated DNA. Notably, the low fork density and long S phase of HCT116 cells were suppressed by extending G1 with low doses of the Cdk4/6 inhibitor Palbociclib. Our results identify decreased origin firing and S-phase extension as new hallmarks of cancer cells potentially responsible for their high genome instability.

Supplementary Material

Suppl. Movie 1: Time-lapse imaging of RPE1-hTERT by phase contrast widefield inverted microscope for 45 hours (1 frame every 10 min).

Suppl. movie 2: Time-lapse imaging of RPE1-hTERT by fluorescence widefield inverted microscope for 80 hours (1 frame every 10 min). RPE1-hTERT were incubated with 100 nM SiR-Hoechst vital stain 4 hours prior to acquisition.

Supplementary Tables

Suppl. table I Suppl. figure 3: Single-molecule analysis of fork speed (FV) in RPE1-hTERT, HCT116 and HeLa cells.

83-354 fork velocities were analyzed per experiment (n=4, mean).

Suppl. figure 4: An example of EdU/BrdU FACS dot-plot of HCT116 cancer cell line treated with A) DMSO, B) 0.16 µM PD for 24 hours then released in a medium containing EdU for 30 min, chased 0.5, 2, 4, 6, 8, and 9 h with thymidine and nocodazole followed by (30 min) BrdU pulse. The fraction of doublepositive cells was plotted against increasing thymidine chase, provides S-phase duration.

Suppl. figure 5: Quantification of MCM loading in G1. A) HCT116 cells treated with DMSO or 0.16 µM PD for 24 hours. G1-I and G2-M gates were considered as negative controls. The mean fluorescence intensity was determined in each of the G1 subgates, subtracted from the G1-I or G2-M gates intensity and were compared in both conditions B) U2OS overexpressing empty, geminin and CDT1 vectors. G2-M gate was considered to be the negative control. The mean fluorescence intensity was determined in G1 gate, subtracted from the G2-M and was compared between conditions.

Suppl. figure 3. Al Ahmad Nachar et al

RPE1-hTERT HCT116 HeLa

Suppl. Table I. Fourteen well-characterized cancer cell lines of nine different epithelial tissues.

Suppl. table II.

Determining the DNA content in G1 by DAPI fluorescence intensity measured by FACS using three different concentrations. The G1 peak of six cancer cell lines was compared to the untransformed diploid RPE1-hTERT cell line.

Suppl. table III. Time needed to replicate the genome based on an extrapolation of the DNA combing data. An example of the calculations done based on DNA content ratio determined in table 4.

Given that:

Number of forks per pulse: Genome size x GIFD DNA synthesized/min: Genome size x GIFD x FV Extrapolated SPD: Genome size/ DNA synthesized/min = Genome size/Genome size x GIFD x FV = 1/Replication potency.

Domain organization

A commonly held view of chromosomal replication organization is that spatially-constrained 0.4-0.8

Mb replication domains (RD) of uniform replication timing and containing 3-6 synchronously firing origins are separated by temporal transition regions (TTR) of equal size either devoid or with a much lower density of origins. Given a FV of ~2 kb/min and a IOD of 100-120 kb within RDs, this model predicts that RDs should get fully replicated within 30-60 min, whereas TTRs may require 3-6 hours for their replication. Although such domains with numerous synchronous origins and rapid replication completion may exist, we believe that they are not very common. Indeed using increasing pulse times (30 to 180 min) we found that most replication tracts remain smaller than 200 kb even after 120 or 180 min of labelling, and that only a small fraction of fibres > 400 kb are more than 70% replicated after a 60 min CldU pulse (Fig. 4D,E). Interestingly this fraction of near-fully replicated fibres increases faster in MEF than in mESC, indicating once more that replication domains are organized differently in the two cell types. We conclude that the majority of RDs either do not contain 3-6 but rather 1-2 active origins, as previously suggested [START_REF] Berezney | Heterogeneity of eukaryotic replicons, replicon clusters, and replication foci[END_REF][START_REF] Yurov | The units of DNA replication in the mammalian chromosomes: evidence for a large size of replication units[END_REF], or that replication forks commonly pause for a significant time before merging.

Replication dynamics during the differentiation of mESC in NPC

Comparing different cell types does not convey the dynamic nature of the transitions that come with the progression from one cellular state to another. To this end we monitored by DNA combing the replication parameters during the differentiation of mESC into neural precursor cells (NPC). mESC were differentiated in vitro first by LIF removal, then by retinoic acid addition, and analysed during 9 days, at stages EB3, EB6 and EB9 of embryoid body formation [START_REF] Bibel | Differentiation of mouse embryonic stem cells into a defined neuronal lineage[END_REF]. Interestingly we found that mean local inter-origin distance decreased from 152 kb in mESC to 131 kb at EB3 and 103 kb at EB6, before rising again thereafter (Fig. 5A). These differences in IODs are not due to biased sampling as mean fibre length was similar in all cases (Fig. 5B). The transient decrease in IOD suggests that cells may reorganize globally their replication programme and/or go through a period of replication stress during the early stages of differentiation, which would cause the firing of dormant origins and the reduction of IODs.

The latter interpretation is supported by the 2-3-fold increase in gH2AX levels seen at the same stages (Fig. 1E). Accordingly, mean fork velocity dropped significantly at the EB3 stage (1.57 kb/min compared to 1.84 kb/min before the differentiation), and fluctuated thereafter (Fig. 5C). Surprisingly the global instant fork density (GIFD), which was already low in mESC (1.03 fork/Mb), decreased even further at EB6 (0.69 fork/Mb) and EB9 (0.54 fork/Mb) (Fig. 5D), at the time cells adopt a novel cell cycle structure characterized by a long G1 phase (Fig. 1D). The drop in FV and IOD happens already at EB3, when the first replication timing changes occur, before transcriptional changes associated to lineage commitment 2015). The establishment and maintenance of these strong repressive histone marks may thus depend on the lengthening of the cell cycle that comes with differentiation. Programmed fork pausing can also impose the polarity of replication within a given locus, which can have cell fate consequences as nicely exemplified by mating type switch control in fission yeast [START_REF] Dalgaard | Does S. pombe exploit the intrinsic asymmetry of DNA synthesis to imprint daughter cells for mating-type switching?[END_REF]. Interestingly the Oct4 and Nanog genes were shown to change replication polarity between ESC (where they are expressed) and differentiated cells (where they are repressed) [START_REF] Schultz | Single-molecule analysis reveals changes in the DNA replication program for the POU5F1 locus upon human embryonic stem cell differentiation[END_REF]. Deciphering the links between replication dynamics (origin choice, fork polarity, timing) and the expression of developmental genes is an interesting avenue for future studies.

DNA replication during differentiation

Using an in vitro model of early embryogenesis by differentiation of mESC in embryoid bodies (EB)

and neural precursor cell (NPC), we found that the cell cycle time and G1 length progressively increase from EB3 to EB9, causing a proportional decrease in the fraction of cells in S phase (Fig. 1C,D). At the earliest steps of differentiation (EB3) corresponding to post-implantation epiblast stages, we detected an increase in histone H2AX phosphorylation (Fig. 1E) as well as a moderate decrease in replication fork velocity (FV) and inter-origin distances (IOD), compared to the starting mESC (Fig. 5A,C). Together these three parameters suggest the existence of a mild replication stress, which may contribute to the EtoL replication timing changes and global genome reorganization that occur at this stage [START_REF] Hiratani | Genome-wide dynamics of replication timing revealed by in vitro models of mouse embryogenesis[END_REF]. Indeed replication stress is known to delay the firing of late origins via an intra-S checkpoint operating through Chk1-mediated phosphorylation and inhibition of the initiation factors Treslin/Sld3 and Dbf4 [START_REF] Guo | Interaction of Chk1 with Treslin negatively regulates the initiation of chromosomal DNA replication[END_REF][START_REF] Zegerman | Checkpoint-dependent inhibition of DNA replication initiation by Sld3 and Dbf4 phosphorylation[END_REF]. During both Drosophila and Xenopus early embryogenesis, transient Chk1 activation at pre-MBT stages is responsible for the progressive lengthening of S phase by imposing late replication to some sequences [START_REF] Shermoen | Developmental control of late replication and S phase length[END_REF][START_REF] Shimuta | Chk1 is activated transiently and targets Cdc25A for degradation at the Xenopus midblastula transition[END_REF]. Moreover mice and fly embryos lacking the upstream ATR kinase die from mitotic catastrophe precisely (mouse E6.5/EB3, fly NC13) after the first replication timing changes took place [START_REF] Blythe | Coordinating Cell Cycle Remodeling with Transcriptional Activation at the Drosophila MBT[END_REF][START_REF] Brown | ATR disruption leads to chromosomal fragmentation and early embryonic lethality[END_REF][START_REF] Farrell | From egg to gastrula: how the cell cycle is remodeled during the Drosophila mid-blastula transition[END_REF]. Our data on fork slow down and increased H2AX phosphorylation at EB3 are consistent with a model of replication stress-induced remodelling of the replication program at early stages of differentiation of ESC. What triggers this early stress in Drosophila or Xenopus embryogenesis is still debated, but the proposal that it might be a drop in Cdk1 activity required for late origin firing fits the sharp decline of cyclin A2 and B1 and increase of p21/p27 CKI seen at this early stage of mESC differentiation [START_REF] Collart | Titration of four replication factors is essential for the Xenopus laevis midblastula transition[END_REF][START_REF] Farrell | From egg to gastrula: how the cell cycle is remodeled during the Drosophila mid-blastula transition[END_REF]Fujii-Yamamoto et al., 2005b).

DNA Replication plasticity during in vitro cell culture and immortalization

10 7 cells/ml. After 10 min lysis on ice the samples were centrifuged at 65000g for 10 min to spin down the nuclei. The pellet was washed in TEB and centrifuged. Finally, the pellet was resuspended in 0.2N

HCl and the histones were acid-extracted overnight at 4°C. After centrifugation, the pellet was retrieved and proteins quantified. Control cells were irradiated with UVC 254 nm, 20 J/m 2 . Proteins were separated in 15% polyacrylamide gel and then transferred onto nitrocellulose membrane. The membrane was blocked in TBST containing 5% non-fat milk. The antibodies used for detection were: anti-gH2AX

(Millipore, 05-636, 1:1000), anti-H2AX (Abcam, ab11175, 1:5000) and secondary anti-mouse and antirabbit antibodies coupled to Alexa800 and Alexa680 fluorophores. Signals were detected using LI-COR Odyssey Imaging System.

EdU foci staining and analysis

Asynchronous cells growing on coverslips were pulse-labeled with 10 µM EdU for 5 min. Cells were fixed with paraformaldehyde 3.2% for 12 min, permeabilized with Triton X-100 0.2% and incubated with 1% BSA/0.5% Tween 20. EdU foci were revealed by staining with Alexa 536-azide using Click-iT chemistry. DNA was stained with DAPI and the coverslips were mounted with Prolong® gold anti-fade reagent (Invitrogen).

SUPPLEMENTAL MATERIAL

Supplementary methods

Time-lapse video-microscopy

The cells were plated a day before imaging on 6-well plates. Acquisitions were done using Zeiss Axioimager Z1 microscope with Coolsnap HQ2.2 camera and 20x objective. Images were taken every 10 min for 50 h.

Supplementary figure legends

Suppl. Following this logic, mouse ESC showed a significant slight decrease in the CldU/IdU ratio compared to MEFs. However, the CldU/IdU ratio was not less than 1 in both MEFs and mouse ESC indicating that fork pausing is not detected during the 15 minutes pulse. In order to test whether fork pausing events can be detected using DNA molecular combing, MEFs were treated with pyridostatin, a stabilizer of G-quadruplexes, for 2-3 hours. The stabilization of Gquadruplexes increases the chance of forks to pause as they encounter these barriers. MEFs were treated with 500 nM or 2 µM pyridostatin-containing medium followed by 15 minutes IdU pulse then 60 minutes of CldU pulse. The 60-minute pulse was used to increase the chances of detecting the pausing events that the forks might encounter. Measuring CldU track length in 500 nM pyridostatin-treated MEFs and DMSO-treated controls revealed no significant difference in