Jean-Charles Cadoret

Evi Soutoglou

Annabel Quinet

Melissa Leriche

Clara Bonnet

Jagannath Jana

Gita Chhetri

Sabrina Mennour

Sylvain Martineau

Vincent Pennaneach

Didier Busso

Xavier Veaute

Pascale Bertrand

Sarah Lambert

Kumar Somyajit

Patricia Uguen

Ste ´phan Vagner
email: stephan.vagner@curie.fr

Break-Induced Replication BirA Biotin

Le Hir for accepting to read, comment and evaluate my work

INTRODUCTION

Au cours de sa vie, l'ARN est associé avec des protéines de liaison à l'ARN (RBP) pour former des complexes ribonucléoprotéiques. Cette interaction dynamique permet de contrôler le métabolisme de l'ARN messager (ARNm) et est crucial pour de nombreux processus cellulaires impliqués dans la régulation post-transcriptionnelle de l'expression des gènes tels que la transcription, l'ajout d'une coiffe en 5', l'épissage alternatif, la polyadénylation de l'extrémité 3', la translocation du noyau au cytoplasme, la localisation subcellulaire de l'ARN, la traduction et enfin sa stabilité.

Parmi les ARNm cibles des RBP, il y a des gènes de réponse aux dommages de l'ADN (DDR) ce qui positionne les RBP comme des acteurs dans la maintenance de l'intégrité du génome. En plus de ce rôle lié à leur fonction de liaison à l'ARN, les RBP sont localisées aux sites de lésions et aident ainsi dans les voies du DDR.

Les RBP participent à prévenir les dommages de l'ADN, en empêchant par exemple la formation des hybrides ARN:ADN (R-loop) et à résoudre les contraintes topologiques, en interagissant avec la topoisomérase TOP1. Elles sont également directement impliquées dans la signalisation, en s'associant à l'ADN endommagé ou à d'autres facteurs du DDR. Par ailleurs, même s'il y a de plus en plus d'évidences que l'ARN est produit au niveau des dommages et joue un rôle dans la signalisation de réparation, il n'est pas clairement défini si la fonction des RBP dans le DDR dépend de leur activité de liaison à l'ARN. En outre, plusieurs protéines du DDR ont été suggérées comme ayant des capacités de liaison à l'ARN, mais le rôle de cette éventuelle activité reste à être démontré. De part toutes ces interactions, les RBP sont donc des acteurs de la stabilité du génome et leur expression aberrante ou l'altération de leurs fonctions peuvent être associées à des pathologies, incluant les cancers.

Ces dernières années, environ 1400 RBP ont été identifiées dans les cellules humaines.

Une RBP est communément définie comme étant une protéine liant directement l'ARN à travers un ou plusieurs domaines de liaison à l'ARN (RBD), induisant un changement dans le destin de l'ARN qu'elle lie. Ces domaines classiques de liaison à l'ARN sont le motif de reconnaissance de l'ARN (RRM), le domaine d'homologie K (KH), le doigt de zinc (ZnF) et le domaine de liaison à l'ARN double-brin (dsRBD), pour ne citer que les mieux caractérisés. Plusieurs RBD peuvent être retrouvés dans une seule RBP, augmentant ainsi l'affinité et la spécificité des interactions protéine-ARN. Par ailleurs, de récentes études ont révélé l'existence de RBP dîtes non-conventionnelles qui ne présentent pas de domaine spécifique. Ces RBP sont souvent composées de régions désordonnées capables de lier l'ARN. L'absence de structure de ces régions leur permet de reconnaître des séquences et des structures d'ARN qui ne sont pas accessibles à des RBD plus structurés. Les régions désordonnées sont classées selon la répétition du motif : Serine/Arginine (SR), Arginine/Glycine (RGG ou GAR) et les patchs basiques Arginine ou Lysine (R/K patches).

Ces régions désordonnées, et d'une manière globale les interactions ARN-protéine, ont pu être identifiées et caractérisées grâce aux approches à grande échelle développées au cours de la dernière décennie. Aujourd'hui, il existe un éventail de méthodes à adapter en fonction de la problématique à résoudre. Il existe des techniques centrées Comme décrit plus haut, les RBP jouent un rôle dans le métabolisme de l'ARNm et également dans le métabolisme de l'ADN, tels que la réplication ou la réparation des dommages où les RBP interagissent plutôt avec des facteurs de ces mécanismes. La réplication de l'ADN est un processus conservé et finement régulé, permettant à une cellule de dupliquer correctement son matériel génétique avant de le transmettre lors de la phase de division. Ce mécanisme bidirectionnel et semi-conservatif implique de nombreuses protéines, qui ne sont, par ailleurs, pas toujours bien déterminées.

La réplication de l'ADN commence au niveau des origines de réplication avec le chargement du complexe de pré-réplication en fin de phase G1. À la transition G1/S, ce complexe sera activé pour former alors le complexe de pré-initiation. Ce complexe protéique permet de dérouler l'ADN double-brin menant au recrutement de facteurs de réplication pour former le réplisome. Le réplisome commence alors à convertir le complexe de pré-initiation en deux fourches de réplication progressant dans des directions opposées. Deux brins sont alors déterminés : le brin précoce synthétisé de manière continue, et le brin retard synthétisé de manière discontinue et constitué de fragments d 'Okazaki. Pour initier la synthèse de l'ADN sur ces brins, et notamment sur le brin retard, des amorces ARN-ADN sont nécessaires. L'amorce ARN en 5', entre 7 et 12 ribonucléotides, est produite par les primases et l'amorce ADN, entre 10 et 25 désoxyribonucléotides, est générée par l'ADN polymérase α considérée comme sujette aux erreurs. À la suite de ces amorces, l'ADN polymérase δ synthétise, sur le brin retard et de manière fidèle, les fragments d'Okazaki jusqu'à 100-200 nucléotides.

Lorsque l'ADN polymérase δ rencontre le fragment d'Okazaki en aval, elle déplace l'amorce ARN en une structure simple-brin appelée le « flap ». La maturation de ce flap est un processus critique pour le bon déroulé de la réplication de l'ADN et tous les acteurs impliqués dans la maturation ne sont pas encore bien décrits. Le flap obtenu est un substrat reconnu et clivé par l'endonucléase FEN1 laissant un espace entre deux fragments d'Okazaki que l'ADN ligase 1 (LIG1) peut ensuite utiliser pour joindre les fragments et ainsi obtenir un ADN double-brin fonctionnel. Dans certains cas, encore mal définis, il y a un retard de clivage de FEN1 induisant une voie noncanonique de maturation du brin retard : la voie du flap long. L'ADN polymérase δ continue alors le déplacement de l'amorce et les protéines de réplication A (RPA) se lient à la partie ADN simple-brin pour recruter et stimuler l'activité de la nucléase DNA2. DNA2 raccourcit alors le long flap, permettant d'obtenir de nouveau un substrat clivable par FEN1. Le passage entre un flap court et long n'est pas complètement compris mais des hypothèses émergent. Il a cependant été suggéré que le long flap serait favorisé dans des régions transcriptionellement actives, pour permettre à l'ADN polymérase δ de corriger les éventuelles erreurs de l'ADN polymérase α et ainsi diminuer le taux de mutation.

Enfin, à la suite de la maturation des fragments d'Okazaki, la réplication de l'ADN se termine, lorsque deux fourches convergentes se rencontrent. Les tensions topologiques sont alors résolues, les espaces restants sont remplis et ligaturés et les complexes de réplication sont dissociés de la chromatine. Chaque étape de la réplication doit donc être réalisée sans défaut et sans stress pour maintenir la stabilité du génome et la progression dans le cycle cellulaire.

En effet, des altérations dans la maturation des fragments d'Okazaki conduisent à une instabilité génétique et sont souvent associées à des pathologies. Ces défauts induisent un espacement (« gap ») d'ADN simple-brin, entre deux fragments d'Okazaki non maturés, qui est reconnu par la protéine PARP1 résultant alors en une accumulation des chaînes poly(ADP-ribose) (PAR) en phase S. Cibler PARP1 par des inhibiteurs (PARPi) est souvent utilisé en clinique, notamment pour cibler les cancers déficients pour les gènes BRCA. En effet, la perte de BRCA augmente ces gaps qui sont alors visés par les PARPi. La perte de 53BP1 dans ce contexte induit une résistance à ces drogues.

Par ailleurs, la cellule est constamment sujette à un stress, par des agents endogènes (dommage de l'ADN, gaps d'ADN simple-brin, hybrides ARN:ADN, structure Gquadruplex de l'ADN) ou exogènes (drogues ciblant le métabolisme des nucléotides, l'activité de polymérases, les topoisomérases), ce qui se traduit par le ralentissement ou l'arrêt de la fourche. La fourche doit alors être réparée avant de redémarrer pour éviter de s'effondrer, créant un état irréversible où la fourche ne peut pas reprendre la réplication. Plusieurs mécanismes de réparations existent : la tolérance aux dommages de l'ADN, la régression de la fourche et le redémarrage médié par la recombinaison homologue (HR). Par ailleurs, même si les facteurs de la HR sont les acteurs majeurs pour répondre au stress réplicatif, certaines protéines de la voie de ligature des extrémités non homologues (NHEJ) peuvent aussi jouer un rôle.

Lorsqu'il y a une cassure double-brin, il y a deux voies de réparation possible. Si c'est en phase S alors la voie HR est favorisé, à l'inverse si c'est en phase G1 la voie NHEJ est choisie. Une protéine clé dans le choix de ces voies est la protéine 53BP1. 53BP1 est codée par le gène TP53BP1 sur le chromosome 15 du génome humain et a été historiquement identifiée en 1994 comme partenaire de la protéine p53, un facteur de transcription considéré comme le « gardien du génome ». Le complexe 53BP1-p53 permet de réguler les fonctions de p53 pour répondre de manière adaptée à l'environnement cellulaire.

Par la suite, 53BP1 est devenue principalement connue pour son rôle dans la signalisation des voies de réparation de l'ADN et notamment dans le choix de ces voies. 53BP1 favorise la voie NHEJ en réponse à une cassure double-brin spontanée, mais aussi dans des contextes où la cassure est programmée : au niveau des télomères et pour la recombinaison V(D)J et isotypique nécessaires à la diversité des anticorps.

L'activité anti-résection de 53BP1 est cruciale pour son rôle dans le choix des voies.

En effet, la première étape de la voie HR est une étape de résection par les nucléases et 53BP1 protège donc l'ADN de cette dégradation, par ses interactions avec différentes protéines, tels que RIF1, DYNLL1 et Shieldin. Ces interactions protéiques sont médiées par la structure de la chromatine puisque 53BP1 reconnaît des marques d'histones : H4K20me2 via ses domaines Tudor et H2AK15ub via son domaine UDR.

En plus de son rôle dans la réparation de l'ADN, 53BP1 participe à la résolution du stress réplicatif. Lorsque la fourche de réplication est bloquée, 53BP1 est recrutée à l'ADN naissant pour éviter sa résection. La perte de 53BP1 induit donc une diminution du taux de réplication et de la survie cellulaire, ainsi qu'une accumulation d'aberrations chromosomiques. De plus, 53BP1 est impliquée dans la protection des fourches régressées. Similaire à sa fonction anti-résection dans le choix des voies de réparation des cassures double-brin, 53BP1 favorise une voie sans clivage pour le redémarrage des fourches bloquées.

Les interactions que 53BP1 établies, même si elles ne sont pas toutes connues, sont médiées par ses domaines protéiques. C'est une protéine de 1972 acides aminés dont la structure peut être divisée en trois parties : Une région N-terminal contenant 28 sites Sérine/Thréonine/Glutamine impliqués dans les interactions avec RIF1 pour l'activité anti-résection de 53BP1, une région centrale, qui est la région minimale de formation de foyers aux sites de lésion, contenant un domaine d'oligomérisation (OD), un domaine GAR, deux domaines Tudor, un signal de localisation nucléaire (NLS), un domaine de recrutement dépendant de l'ubiquitine (UDR) et une partie C-terminal composée de deux domaines BRCT essentiels pour la liaison avec son partenaire p53.

FA

Fanconi Anemia

FBH1

F-Box DNA Helicase 1

FEN1

Flap structure-specific Endonuclease 1

FMRP

Fragile X Mental Retardation Protein

FXS

Fragile X Syndrome

GAR

Glycin-Argining Rich

GFP

Green Fluorescent Protein

GINS

Go-ichi-ni-san complex

Gloe-seq

Genome-wide ligation of 3'-OH ends followed by sequencing

G-phase

Gap phase (G1; G2)

G-quadruplex

Alternative splicing

Alternative splicing is a critical process which contributes to protein diversity. The heterogenous nuclear ribonucleoprotein (hnRNP) and Serine/Arginine proteins are the main players in this mechanism. Splicing factors bind to specific sequences on pre-mRNA molecules to regulate exon skipping, intron retention or the use of alternative splice sites [START_REF] Chen | Mechanisms of alternative splicing regulation: insights from molecular and genomics approaches[END_REF]. For instance, in 2020, a study conducted by the Vagner laboratory have demonstrated that the splicing factors zinc finger ran-binding domain-containing protein 2 (ZRANB2) and the SYF2 pre-mRNA splicing factor (SYF2)

both associate with the epithelial cell transforming 2 (ECT2) mRNA to favour the production of a variant that includes the exon 5. Interestingly, in breast cancer, depletion of this ECT2 variant increases sensitivity to doxorubicin, a drug that inhibits topoisomerase II and induces DNA damages [START_REF] Tanaka | ZRANB2 and SYF2-mediated splicing programs converging on ECT2 are involved in breast cancer cell resistance to doxorubicin[END_REF].

Polyadenlyation

RBPs are responsible for the process of polyadenylation, which consists of the addition of stretches of Adenines at the 3' end of mRNAs. First the pre-mRNA is cleaved by the cleavage and polyadenylation specificity factor (CPSF) and the cleavage stimulatory factor (CstF). The poly(A) tail is then added by the poly(A) polymerase (PAP) allowing the stabilisation of the mRNA [START_REF] Proudfoot | Integrating mRNA Processing with Transcription[END_REF]. Alteration in the polyadenylation affects the transcript at several levels including its export in the cytoplasm, its translation, and its decay rate. These mechanisms (cleavage and polyadenylation) are regulated differently depending on the cellular context. In damaged cells, CstF is sequestered in complexes with various factors, including RNA polymerase II (RNA Pol II), the tumour suppressors BARD1/BRCA1 and PARN, which inhibits 3' processing. However, Adrien Decorsière et al. have shown that following DNA damage, the RBPs hnRNP H and F prevent this sequestration to maintain the 3' maturation of p53 mRNA. These RBPs interact with a G-quadruplex structure downstream of the polyadenylation site of p53 resulting in the recruitment of Cstf and PAP [START_REF] Decorsière | Essential role for the interaction between hnRNP H/F and a G quadruplex in maintaining p53 pre-mRNA 3'-end processing and function during DNA damage[END_REF]. Since, the p53 protein is considered as "the guardian of the genome", the p53 mRNA must be tightly regulated by RBPs at all levels, mentioned above, to properly respond to potential cellular stresses [START_REF] Haronikova | The p53 mRNA: an integral part of the cellular stress response[END_REF].

mRNA stability

Along with the poly(A) tail, the 3' untranslated region (UTR) plays an important role in transcript stability. Among the various 3'UTR-interacting proteins, the human antigen R (HuR) is known to bind a multitude of mRNAs to enhance their translation.

For instance, in response to genotoxic agents, HuR stabilises the poly(ADP-ribose) glycohydrolase (PARG) mRNA, thereby upregulating PARG expression to facilitate DNA repair through the removal of poly(ADP-ribose) (PAR) chains and the modulation of the interaction between poly(ADP-ribose) polymerase 1 (PARP1) and chromatin [START_REF] Chand | Posttranscriptional Regulation of PARG mRNA by HuR Facilitates DNA Repair and Resistance to PARP Inhibitors[END_REF]. The p53 mRNA is also targeted by HuR. In colorectal carcinoma cells, HuR binds to p53 through the AU-rich elements (AREs), in a UV-dependent manner to stabilise it and thus favour its translation [START_REF] Mazan-Mamczarz | RNA-binding protein HuR enhances p53 translation in response to ultraviolet light irradiation[END_REF].

Moreover, in response to exposure to ionizing radiation HuR can dissociate from its target mRNA hence promoting the cell survival [START_REF] Masuda | Global dissociation of HuR-mRNA complexes promotes cell survival after ionizing radiation[END_REF].

Translation

Once the mRNA is stabilised, translation can begin, and this post-transcriptional mechanism is also controlled by RBPs. The translation of p53 mRNA is regulated by numerous RBPs, such as murine double minute 2 (MDM2). Under normal conditions, the E3 ubiquitin ligase MDM2 interacts with the p53 protein to induce its degradation.

Following DNA damage, MDM2 gets phosphorylated by the Serine/Threonine kinase Ataxia-telangiectasia mutated (ATM), enabling its interaction with p53 mRNA, and thereby stimulating its translation. Indeed, phosphorylated MDM2 binds p53 mRNA through its RING domain and this association results in an increase in synthesis and a decrease in MDM2-dependent degradation of p53 [START_REF] Gajjar | The p53 mRNA-Mdm2 interaction controls Mdm2 nuclear trafficking and is required for p53 activation following DNA damage[END_REF]. The murine double minute 4 (MDMX), an MDM2 homolog also phosphorylated by ATM, facilitates this interaction by serving as a chaperone for p53 mRNA and providing an MDM2 binding platform [START_REF] Haronikova | The p53 mRNA: an integral part of the cellular stress response[END_REF]. Furthermore, disturbances in the translation process are a major event leading to the development of cancer (For review [START_REF] Fabbri | The plasticity of mRNA translation during cancer progression and therapy resistance[END_REF]. When oncogenic signalling pathways are active, this can induce the activation of the eukaryotic translation initiation complex eIF4F (composed of eIF4A, eIF4E and eIF4G) and thus promoting translation. RBPs can then act downstream of these signalling pathways to control the translation of specific mRNAs. For example, HuR increases the stability of eIF4E transcripts in certain cancers, inducing the expression, among others, of MYC. The oncoprotein MYC is involved in cell proliferation, and it enhances the expression of components of the eIF4F complex.

Localisation

Many RBPs are also involved in the transport of RNA molecules to specific subcellular compartments. For instance, the fragile X mental retardation protein (FMRP)

associates with G-quadruplexes forming in the 3'UTR of its mRNAs target to promote their localisation throughout neuronal projections. FMRP interacts with RNA through its Glycine-Arginine-rich domain (GAR domain composed of a RGG motif) and the deletion of this domain results in a defect in transcript transport. These defects are explained by a reduced ability of FMRP to bind mRNAs, and not by a reduced capacity of the protein to be transported into neurites [START_REF] Goering | FMRP promotes RNA localization to neuronal projections through interactions between its RGG domain and G-quadruplex RNA sequences[END_REF]. Moreover, FMRP is an essential protein for proper brain development and is involved, among other functions, in neuronal activity and synaptic plasticity. Importantly, the absence of FMRP leads to Fragile X Syndrome (FXS), a genetic disorder that often causes cognitive impairment, behavioural disorders and physical abnormalities. This syndrome is one of the most common forms of hereditary intellectual disability.

Additionally, a mutation found in patients with FXS involves an insertion within the RGG box, causing an ORF frameshift, disrupting the RGG box and leading to the production of a short C-terminal sequence. The novel C-terminal peptide encodes a functional nuclear localisation signal altering the subcellular localisation of FMRP [START_REF] Okray | A novel fragile X syndrome mutation reveals a conserved role for the carboxy-terminus in FMRP localization and function[END_REF].

As described above RBPs act as co-and post-transcriptional regulators, and may participate in genome stability by targeting DNA damage response (DDR) genes (Dutertre et al., 2014). Furthermore, several DDR proteins have been characterised as having RNA-binding capabilities (Fig. 3) (Audoynaud et al., 2021). For example, the NHEJ factor Ku is an RBP that directly binds to RNA with a hairpin structure possessing a bulge. One target of Ku is the p53 mRNA and Ku interacts with the 3'UTR region of p53 to repress its translation under normal conditions. When DNA is damaged, this interaction is inhibited to allow the expression of p53 [START_REF] Lamaa | A novel cytoprotective function for the DNA repair protein Ku in regulating p53 mRNA translation and function[END_REF]. Another example is the catalytic subunit of DNA-dependent protein kinase (DNA-PKcs) which interacts with the long non-coding RNA LINP1 to promote the NHEJ pathway. LINP1 also acts as a scaffold in the Ku/DNA-PKcs complex [START_REF] Zhang | Long noncoding RNA LINP1 regulates repair of DNA double-strand breaks in triple-negative breast cancer[END_REF]. One last example of a DDR protein identified as an RBP is the PARP1 protein.

PARP1

Direct players in DNA repair signalling

Apart from RBPs with defined functions in RNA biology, these past years have seen the discovery of an increasing number of RBPs localised at DNA damage sites and therefore playing a more direct role in DDR (Fig. 3). These RBPs constitute a new class of DDR players and are now known as DNA Damage RNA-binding proteins (DDRBPs) (Dutertre and Vagner, 2017). However, although it is becoming increasingly evident that RNA itself is involved at damage sites (Audoynaud et al., 2021;Michelini et al., 2017Michelini et al., , 2018;;[START_REF] Zong | RNA: a double-edged sword in genome maintenance[END_REF], it is not yet clearly defined whether the function of RBPs in DNA repair signalling depends on their RNA-binding activity.

Physical agents are the ionizing radiations and ultraviolet (UV) rays. Chemical agents are typically agents used in chemotherapy, such as alkylating agents (methyl methanesulfonate), cross-linking agents (cisplatin) and topoisomerase inhibitors (etoposide) but they can also be environmental pollutants such as cigarette smoke [START_REF] Ciccia | The DNA Damage Response: Making It Safe to Play with Knives[END_REF]. Endogenous sources include replication stress (errors, stalled forks), products of cellular metabolism (free radicals) and spontaneous chemical modifications (depurination, deamination) [START_REF] Tubbs | Endogenous DNA Damage as a Source of Genomic Instability in Cancer[END_REF].

These endogenous and exogenous lesions generally result in base modifications, mismatches, inter-or intra-strand crosslinks and single-or double-strand breaks.

RBPs prevent DNA damage by controlling R-loop formation, DNA topology and DNA metabolism

RBPs intervene to help prevent the occurrence of the lesions described above (Fig. 3) (Dutertre et al., 2014). As mentioned, an important endogenous source of DNA damage is the collision between transcription and replication leading to stalled replication forks during the S-phase. The nascent RNA transcribed by the RNA polymerase II hybridizes with the template DNA strand thus forming RNA:DNA hybrids, displacing the complementary DNA strand and creating a three-strand structure called an R-loop. The formation of R-loops is favoured at Guanine-rich DNA repeats and topologically constraint regions and can be controlled by multiple RBPs [START_REF] Aguilera | R Loops: From Transcription Byproducts to Threats to Genome Stability[END_REF]. For instance, depletion of the RBP Serine/Argininerich splicing factor 1 (SRSF1 or ASF/SF2) results in an accumulation of R-loops and a hypermutation phenotype. SRSF1 interacts with the nascent pre-mRNA to promote its splicing, facilitated by the carboxy-terminal domain of RNA Pol II which recruits SRSF1 to the transcript, preventing it from associating with the template DNA and thus inhibiting the formation of R-loops [START_REF] Li | Inactivation of the SR protein splicing factor ASF/SF2 results in genomic instability[END_REF]. SRSF1 also interacts with DNA topoisomerase I (TOP1) to inhibit the occurrence of R-loops [START_REF] Tuduri | Topoisomerase I suppresses genomic instability by preventing interference between replication and transcription[END_REF]. TOP1 resolves topological constraints by relaxing DNA, and interactions between RBPs and TOP1 contribute to safeguarding the integrity of the DNA [START_REF] Czubaty | Proteomic analysis of complexes formed by human topoisomerase I[END_REF].

Several RBPs are also being identified as playing roles in DNA metabolism such as DNA replication and telomere biogenesis, which can be a source of DNA damage.

Indeed, telomeres are a favourable site for the formation of R-loops. The long noncoding telomeric repeat-containing RNA (TERRA) is prone to form this RNA:DNA hybrids resulting in telomere instability. The NonO and SFPQ proteins which bind TERRA, suppress RNA:DNA hybrids formation and replication defects at telomeres to ensure their integrity [START_REF] Petti | SFPQ and NONO suppress RNA:DNA-hybrid-related telomere instability[END_REF]. In addition, the hnRNP family is also strongly involved in telomere regulation. The length of telomeres is regulated by hnRNP C and U by associating with telomerase reverse transcriptase (TERT). The hnRNP F protein interacts with TERRA to control its localisation, thereby impacting the telomere length as well. The hnRNP A1 protein interacts with TERRA, enabling the regulation of telomere coating by the Shelterin complex during DNA replication [START_REF] Nishida | RNA Binding Proteins and Genome Integrity[END_REF]. HnRNP A1 also interacts with the flap structure-specific endonuclease 1 (FEN1) to stimulate its activity and therefore to assist in the maturation of Okazaki fragments during the replication process (see section II-3) [START_REF] Chai | Interaction and stimulation of human FEN-1 nuclease activities by heterogeneous nuclear ribonucleoprotein A1 in alphasegment processing during Okazaki fragment maturation[END_REF].

Furthermore, in response to DNA damage, the RNA helicase UPF1 associates with DNA polymerase δ to mediate DNA replication and DNA repair in S-phase [START_REF] Azzalin | The Double Life of UPF1 in RNA and DNA Stability Pathways[END_REF].

RBPs can be directly involved in DNA repair pathways

Despite all the existed mechanisms that prevent DNA damage, lesions can still occur.

The repair of these lesions is essential for the cell survival and to avoid the accumulation of DNA damage. If the repair is optimal, the cell will proceed with its regular division cycle but if the damage is not properly repaired, it will disrupt the cell cycle. There are several repair pathways with specific actors that depend on the type of lesions and the cell cycle phase (Fig. 3). For example, the removal and replacement of damaged bases are handled by the base excision repair (BER) pathway, mismatched bases by the mismatch repair (MMR) pathway, intra-strand crosslinks by the nucleotide excision repair (NER) pathway, while inter-strand crosslinks, as well as double-strand breaks, are repaired of by the homologous recombination (HR) pathway in S-phase or by the non-homologous end joining (NHEJ) pathway in G1phase [START_REF] Hakem | DNA-damage repair; the good, the bad, and the ugly[END_REF][START_REF] Huang | DNA damage repair: historical perspectives, mechanistic pathways and clinical translation for targeted cancer therapy[END_REF].

In the past few years, the co-localisation between DNA damage sites and certain RBPs has been observed (Dutertre et al., 2014). RBPs are recruited to the damaged DNA in a manner that may or may not be dependent on their RNA-binding activity. Indeed, once recruited, RBPs can interact, independently of RNA, with DDR factors, DNA or PAR chains. PAR chains generated by poly(ADP-ribose) polymerase (PARP) are necessary for recruiting certain RBPs, mainly from the splicing factors family such as RNAbinding motif protein X-linked (RBMX). Indeed, Britt Adamson and colleagues demonstrated in 2012 that RBMX, also known as heterogenous nuclear ribonucleoprotein G (hnRNP G), accumulates at double-strand break sites in a PARPdependent manner to promote HR [START_REF] Leriche | A genomewide homologous recombination screen identifies the RNA-binding protein RBMX as a component of the DNA-damage response[END_REF]. Moreover, RBMX is capable of binding to damaged DNA to protect the ends [START_REF] Shin | p53 promotes the fidelity of DNA end-joining activity by, in part, enhancing the expression of heterogeneous nuclear ribonucleoprotein G[END_REF]. Indeed, RBPs can be recruited to damage sites by interacting with the DNA to regulate the signalling of repair pathways. For example, the RBP Y-box binding protein 1 (YB-1) interacts with mismatched DNA and can interfere with the MMR repair pathway [START_REF] Chang | YB-1 disrupts mismatch repair complex formation, interferes with MutSα recruitment on mismatch and inhibits mismatch repair through interacting with PCNA[END_REF].

Another example includes the pre-mRNA processing factor 19 (PRP19) E3 ubiquitin ligase whose ubiquitination favours its binding to double-strand DNA to regulate DNA repair pathways [START_REF] Idrissou | The PRP19 Ubiquitin Ligase, Standing at the Cross-Roads of mRNA Processing and Genome Stability[END_REF]. PRP19 is recruited to single-strand DNA, formed following damage, by interacting directly with Replication Protein A (RPA) to facilitate the recruitment of other DDR factors [START_REF] Maréchal | PRP19 Transforms into a Sensor of RPA-ssDNA after DNA Damage and Drives ATR Activation via a Ubiquitin-Mediated Circuitry[END_REF]. RBPs can also be recruited to damage sites through direct protein-protein interaction. For instance, the splicing factor heterogenous nuclear ribonucleoprotein C (hnRNP C) is part of the complex containing the breast cancer gene 1 and 2 (BRCA1/2) and the partner and localizer of BRCA2 (PALB2) to assist in HR [START_REF] Anantha | Requirement of heterogeneous nuclear ribonucleoprotein C for BRCA gene expression and homologous recombination[END_REF].

These examples demonstrate that, in addition to their function in the expression of DDR coding genes, RBPs have a more direct role in the repair pathways that often relies on protein-protein interactions. However, it remains unclear whether it also depends on their RNA-binding activity.

In summary, RNA-protein interactions are crucial for many cellular processes including mRNA processing, regulation of gene expression, and the maintenance of telomere integrity and genome stability. Mutations in RBPs, such as FMRP whose mutations are associated with FXS, can therefore alter RNA-RBP interactions and, consequently, cause human diseases. For this reason, the aberrant expression or altered function of RBPs in pathologies are being increasingly studied [START_REF] Hong | RNA Binding Protein as an Emerging Therapeutic Target for Cancer Prevention and Treatment[END_REF][START_REF] Mohibi | Cancer the'RBP'eutics-RNA-binding proteins as therapeutic targets for cancer[END_REF].

RNA-binding domains

As mentioned above, RBPs are major regulators of gene expression due to their direct interaction with mRNAs. Recent approaches have enabled the classification of RBPs into two categories: canonical and unconventional. Canonical RBPs possess one or more RBDs, while unconventional RBPs may present disordered regions or others unconventional ways to interact with RNAs. Furthermore, most RBDs are conserved and homologous to human RBPs are found in other organisms, such as yeast (Gerstberger et al., 2014a). Among the unconventional RBPs, some are also involved in biological processes distinct from mRNA processing [START_REF] Baltz | The mRNA-bound proteome and its global occupancy profile on protein-coding transcripts[END_REF][START_REF] Castello | Insights into RNA Biology from an Atlas of Mammalian mRNA-Binding Proteins[END_REF][START_REF] Hentze | A brave new world of RNAbinding proteins[END_REF]. Indeed, 27% of identified RBPs in human and 40% in yeast have no known function related to RNA biology and these RBPs are called "enigmRBP" [START_REF] Beckmann | The RNA-binding proteomes from yeast to man harbour conserved enigmRBPs[END_REF]. Many of these RBPs exhibit enzymatic activities and are involved in classical metabolic pathways [START_REF] Beckmann | The expanding universe of ribonucleoproteins: of novel RNA-binding proteins and unconventional interactions[END_REF][START_REF] Castello | Insights into RNA Biology from an Atlas of Mammalian mRNA-Binding Proteins[END_REF][START_REF] Perez-Perri | The RNA-binding protein landscapes differ between mammalian organs and cultured cells[END_REF].

There are several characterised types of RBDs, each exhibiting unique structural and functional properties and a few examples of RBDs are succinctly described in Table 1 [START_REF] Corley | How RNA-Binding Proteins Interact with RNA: Molecules and Mechanisms[END_REF]. It is noteworthy that RBPs can possess both canonical and noncanonical domains. Additionally, a considerable number of RBPs still lack a precisely defined RBD.

RNA-binding domains

Canonical domains

Multiple RBDs can be found within a single RBP. This increases the affinity and specificity of the RNA-proteins interaction. Currently, the best characterised RBDs include the RNA recognition motif (RRM), the K-homology (KH) domain, the zinc finger (ZnF) domain and the double-strand RNA-binding domain (dsRBD).

RNA recognition motif

The RRM is the most abundant domain in higher eukaryotes (Gerstberger et al., 2014b). This common type of RBDs is found in many RBPs, mainly in splicing factors such as the hnRNP A1 protein. In general, the RRM is a 90 amino acids domain that encompasses two conserved sequences known as RNP1 and RNP2. These sequences are composed of 8 and 6 amino acids, respectively. The RRM adopts a characteristic structure, called β1α1β2β3α2β4 topology, involving a four-strand antiparallel β-sheet supported by two α-helices. The RNA-binding sequences RNP1 and RNP2 are positioned in the central β-sheet β3 and β1, respectively. They expose three conserved aromatic residues on the surface of the β-sheet, which serve as the main binding surface for RNA molecules [START_REF] Cléry | From Structure to Function of RNA Binding Domains[END_REF][START_REF] Daubner | RRM-RNA recognition: NMR or crystallography…and new findings[END_REF]. Due to their structure, RRM domains interact with 2-8 nucleotides of single-strand RNAs, and the accumulation of RRMs in an RBP enhances the specificity of the RNA-protein interaction [START_REF] Maris | The RNA recognition motif, a plastic RNAbinding platform to regulate post-transcriptional gene expression[END_REF].

K-homology domain

Historically, the KH domain was discovered in the hnRNP K protein in the 1990s and has been identified in numerous other RBPs ever since, such as translation regulators.

It is a domain composed of 70 amino acids that has the ability to bind both singlestrand RNAs and single-strand DNAs [START_REF] Valverde | Structure and function of KH domains[END_REF]. Generally, the RNAbinding surface of the KH domain includes two α1 and α2 helices, connected by a GXXG loop (X being any amino acid) and a β-sheet. The conserved GXXG motif can recognize 4 nucleotides [START_REF] Cléry | From Structure to Function of RNA Binding Domains[END_REF]. Similar to RRM domains, KH domains are often repeated to enhance the efficiency of RNA-binding [START_REF] Corley | How RNA-Binding Proteins Interact with RNA: Molecules and Mechanisms[END_REF].

Zinc finger domain

While zinc finger proteins are most known for their role as DNA-binding transcription factors, there are certain classes of zinc finger proteins that also function as RNAbinding proteins, like the methyltransferase-like (METTL) enzymes. The zinc finger domain contains 30 amino acids and forms a ββα topology coordinated by a zinc ion.

There are several types of zinc finger domain capable of binding RNA, classified based on the amino acids that interact with this ion (e.g. CCHC, CCCH, CCCC or CCHH zinc fingers) [START_REF] Cléry | From Structure to Function of RNA Binding Domains[END_REF][START_REF] Hall | Multiple modes of RNA recognition by zinc finger proteins[END_REF].

Double-strand RNA-binding domain

Double-strand RNA-binding domains are characterised by a conserved αβββα topology and typically consist of around 70 amino acids. They are found in various proteins across different organisms, such as RNAses or adenosine deaminases acting on RNA (ADAR), and play important roles in RNA processing, RNA editing and RNA interference. It is often considered that these domains recognize the shape of RNA rather than the sequence. However, it has been shown that in vivo, proteins containing the dsRBD bind to specific double-strand RNAs [START_REF] Masliah | RNA recognition by double-stranded RNA binding domains: a matter of shape and sequence[END_REF].

There are also other classical sequence-specific RBDs that have been characterised over the years, and not described here, such as the dead box helicases domain and the Piwi/Argonaute/Zwille (PAZ) domain [START_REF] Corley | How RNA-Binding Proteins Interact with RNA: Molecules and Mechanisms[END_REF].

Intrinsically disordered regions

Among the large family of RBPs, more than half do not possess known RBDs. For comparison, the RRM domain, which is the most common, is only present in 13% of RBPs [START_REF] Beckmann | The expanding universe of ribonucleoproteins: of novel RNA-binding proteins and unconventional interactions[END_REF]. These non-canonical RBDs identified in RBPs do not have the same structural features as the canonical RBDs, but they still play important roles in RNA-binding and processing.

Recent techniques for studying RBDs have revealed that most RBPs actually possess intrinsically disordered regions (IDRs) [START_REF] Castello | Comprehensive Identification of RNA-Binding Domains in Human Cells[END_REF]). An IDR is a region of at least 30 amino acids that lacks stable tertiary structure [START_REF] Uversky | Intrinsically Disordered Proteins and Their "Mysterious[END_REF]. This instability is due to low sequence complexity of IDRs. Indeed, IDRs are depleted in hydrophobic residues and enriched in amino acids that promote disorder, such as charged or polar residues (Glycine, Proline, Lysine, Arginine, Aspartate, Glutamate, Serine), as well as Tyrosine, which is an amino acid found in surfaces interacting with

RNAs. There are different types of IDRs based on the repetition of the disordered motif: Serine/Arginine (SR repeats), Arginine/Glycine (RGG repeats) and Arginine-or Lysine-rich patches (R/K basic patches).

The lack of a defined structure in IDRs enables them to recognize RNA sequences and structures that are not accessible to more structured RBDs. Furthermore, the binding to RNAs stabilizes the disordered structure of these regions [START_REF] Järvelin | The new (dis)order in RNA regulation[END_REF][START_REF] Varadi | Computational approaches for inferring the functions of intrinsically disordered proteins[END_REF]. RBPs with IDRs typically have multiple binding sites that interact with RNAs in a flexible and dynamic manner. However, a very recent study [START_REF] Ray | RNA-binding proteins that lack canonical RNA-binding domains are rarely sequence-specific[END_REF] appears to show that many unconventional RBPs do not bind to RNAs in a specific manner, which could be precisely associated with the abundance of IDRs within the protein.

SR repeats

Serine and Arginine-rich repetitions are predominantly found in SR proteins, known as splicing factors for the pre-mRNA, such as Serine/Arginine-rich splicing factor 1 (SRSF1). The splicing efficiency depends on the number of repeats and phosphorylation of the SR motif. These proteins contain an RRM domain at the Nterminal end, which recognizes the RNA, and an SR-rich domain at the C-terminal end, which plays a role in protein-protein and RNA-protein interactions. In addition proteins containing SR motifs are also involved in various processes, such as mRNA export, translation and genome stability maintenance. For example, SRSF3 acts as a post-transcriptional regulator by controlling internal ribosome entry site (IRES)mediated translation [START_REF] Jeong | SR Proteins: Binders, Regulators, and Connectors of RNA[END_REF].

RGG repeats

The RGG repetitions, also known as GAR, are a motif frequently found in RBPs, but their RNA-binding properties are still poorly defined. The interaction between RNA and RGG repeats varies depending on the sequence and structure of the target RNA.

For example, the RGG motif of the FMRP interacts with its target, the sc1 RNA, through the recognition of a G-quadruplex structure [START_REF] Phan | Structure-function studies of FMRP RGG peptide recognition of an RNA duplex-quadruplex junction[END_REF]. In addition, proteins with RGG repeats play key roles in many cellular processes such as transcription, DNA damage signalling and regulation of apoptosis. For example, meiotic recombination 11 (MRE11), which is the first player in the signalling of DNA repair pathways, possesses a GAR domain and this motif is essential for the recruitment of MRE11 to DNA damage sites [START_REF] Yu | The MRE11 GAR motif regulates DNA double-strand break processing and ATR activation[END_REF].

The activity of the RGG motif in these mechanisms is often regulated by Arginine methylation by protein arginine methyltransferase (PRMT) (Thandapani et al., 2013).

This post-translational modification influences the localisation, activity and function of proteins with GAR domains, as well as the interaction of these proteins with their partners [START_REF] Chong | RGG/RG Motif Regions in RNA Binding and Phase Separation[END_REF]. For instance MRE11 is methylated by the enzyme PRMT1, important for its localisation at DNA damage sites and also for the DNA damage checkpoint response mediated by MRE11 (Boisvert et al., 2005a[START_REF] Boisvert | Methylation of MRE11 regulates its nuclear compartmentalization[END_REF][START_REF] Déry | A Glycine-Arginine Domain in Control of the Human MRE11 DNA Repair Protein[END_REF]. The 53BP1 protein, a crucial protein in double-strand break repair, also possesses an RGG motif (described below in III-2) that is methylated by PRMT enzymes and this methylation is important for stabilizing 53BP1 [START_REF] Boisvert | The GAR motif of 53BP1 is arginine methylated by PRMT1 and is necessary for 53BP1 DNA binding activity[END_REF][START_REF] Hwang | PRMT5 promotes DNA repair through methylation of 53BP1 and is regulated by Srcmediated phosphorylation[END_REF])

R/K basic patches

Basic patches are composed of 4 to 8 Lysines or, less frequently, Arginines forming a highly positive surface that mediates molecular interactions. These motifs are very common among RBPs. Basic patches can be found anywhere in the protein structure and often form islands that flank globular domains, indicating the cooperation between structured and disordered regions [START_REF] Järvelin | The new (dis)order in RNA regulation[END_REF]. An example of a protein with a basic patch is the poly(A) RNA polymerase D5 (PAPD5), involved in the degradation of aberrant rRNA. At its C-terminal end, PAPD5 possesses a basic patch, containing Lysines, that is directly involved in its RNA-binding activity [START_REF] Rammelt | PAPD5, a noncanonical poly(A) polymerase with an unusual RNA-binding motif[END_REF].

In summary, disordered regions as well as canonical RNA-binding domains play an important role in biological processes through their RNA-binding activity. The domains detailed above constitute only a fraction of the existing RBDs. Research on RNA-protein interactions continues to expand, and system-wide approaches (see section I-3) have led to the discovery of new domains and domains that are yet to be annotated. For example, the YT521-B homology (YTH) was identified in 2010 as a novel RBD that binds modified RNAs (m 6 A mark) [START_REF] Zhang | The YTH domain is a novel RNA binding domain[END_REF]. Overall, these additional RBDs enrich the repertoire of RBDs in RBPs and highlight the diverse ways in which RBPs interact with RNAs.

Methods to monitor RNA-protein interactions

The precise mechanisms by which an RBP recognizes and selectively interacts with its 2).

Historically, RNA-protein interactions have been described using methods based on UV-C irradiation at 254 nm, creating irreversible covalent bond within living cells, known as cross-linking. Cross-linking enables the interactions to be maintained and the RNA-protein complexes to be purified under stringent conditions. The UV-C crosslinking requires direct contact ("zero" distance) between the protein and the nucleic acids and does not favour protein-protein cross-linking (Greenberg, 1979;Pashev et al., 1991). UV-C cross-linking depends on the structure of the RNA and the protein, which implies that overall UV-C cross-linking has low efficiency. Indeed, UV-C has a preference for crosslinking uridines [START_REF] Hockensmith | Laser crosslinking of proteins to nucleic acids. I. Examining physical parameters of protein-nucleic acid complexes[END_REF] and also certain amino acids (Cysteine, Lysine, Phenyalanine, Tryptophane and Tyrosine). In addition, even though UV-C crosslinks RNA-protein interactions more efficiently (mainly singlestrand RNA), it has also been used to crosslink interactions between protein and single-strand DNA (Steen et al., 2001).

Furthermore, there are other crosslinking methods used to study RNA-protein interactions, such as crosslinking by formaldehyde treatment which is not specific to RNA and can also crosslink protein-protein interactions [START_REF] Niranjanakumari | Reversible crosslinking combined with immunoprecipitation to study RNA-protein interactions in vivo[END_REF], or photoactivatable-ribonucleoside-enhanced crosslinking (PAR-crosslinking)

with incorporation of 4-thiouridine (4-SU), an uridine analogue [START_REF] Hafner | Transcriptome-wide Identification of RNA-Binding Protein and MicroRNA Target Sites by PAR-CLIP[END_REF].

In the case of PAR-crosslinking, 4-SU is incorporated into nascent RNA and UV-A at 365 nm is used to crosslink interactions between protein and RNA containing 4-SU.

These crosslinking alternatives have disadvantages (protein-protein crosslinking for formaldehyde and cellular toxicity for PAR-crosslinking) which makes UV-C the preferred choice in many techniques for studying RNA-protein interactions.

RNA-centric techniques

RNA-centric methods are used to investigate proteins that bind to a particular RNA.

These methods consist of initiating the study with a specific RNA of interest, followed by the characterisation of the associated RBPs, mainly through mass spectrometry or western blot analysis.

Methods with cross-linking

These methods involve a cross-linking step either UV or formaldehyde. In UV-based

approaches, examples include RNA affinity purification (RAP), peptide-nucleic-acidassisted identification of RBP (PAIR), MS2 in vivo biotin-tagged RAP (MS2-BioTRAP)

and tandem RNA isolation procedure (TRIP). These techniques share the common step of cross-linking, but they differ in the methods used for capturing RNA and purifying of RNA-protein complexes. For example, the RAP method uses biotinylated probes containing an antisense oligonucleotide to the target RNA, enabling purification using streptavidin-coated beads. On the other hand, the PAIR technique uses a peptide nucleic acid probe. Furthermore, there are also methods based on formaldehyde cross-linking, such as capture hybridization analysis of RNA targets (CHART) or chromatin isolation by RNA purification (ChIRP) [START_REF] Ramanathan | Methods to study RNA-protein interactions[END_REF].

Methods without cross-linking

Additionally, there are methods available that do not require a cross-linking step. For example, the RNA proximal protein interaction detection (RaPID) method relies on the (dCas13) fused to a biotin-labelling enzyme specifically to an RNA of interest. Upon addition of biotin, the biotin-labelling enzyme biotinylates proteins in its proximity [START_REF] Gräwe | RNA-Centric Methods: Toward the Interactome of Specific RNA Transcripts[END_REF].

Methods to study the RBPome

All the methods described above start by targeting an RNA of interest. However, in large-scale studies for the discovery of RBPs and better characterisation of the RBPome, there is no specific RNA. Therefore, it was necessary to develop other techniques capable of capturing a group of RNA.

In 2012, two research teams described the RNA interactome capture (RIC) method, which enabled hundreds of RBPs to be identified in HeLa and HEK human cell lines [START_REF] Baltz | The mRNA-bound proteome and its global occupancy profile on protein-coding transcripts[END_REF][START_REF] Castello | Insights into RNA Biology from an Atlas of Mammalian mRNA-Binding Proteins[END_REF]. This method is based on a cross-linking with UV light (UV-C at 254 nm or UV-A at 365 nm). After irradiation, the polyadenylated RNA (poly(A) RNA) is purified using oligo(dT) columns. Finally, the RBP bound to these poly(A) RNAs are analysed using mass spectrometry. In 2016, Thomas Conrad and colleagues developed the serial interactome capture (serIC), a derivative of the RIC, which allows the investigation of RNA-protein interactions specific to the nucleus [START_REF] Conrad | Serial interactome capture of the human cell nucleus[END_REF].

The main limitation of RIC and its derivatives is that these techniques focus on poly(A)

RNA associated with RBPs. In 2018, Xing Chen's group developed a click-chemistryassisted RNA interactome capture (CARIC) strategy, which allows the identification of RBP independent of the polyadenylation [START_REF] Huang | Transcriptome-wide discovery of coding and noncoding RNA-binding proteins[END_REF]. The same year, Matthias

Hentze's group developed the complex capture (2C) (Fig. 4) technique to address this limitation (Asencio et al., 2018). After UV-C cross-linking, cells are lysed and the lysate is loaded on a silica column. Thanks to its physico-chemical properties, the silica column retains RNA, whether free or associated with proteins. It is then possible to analyse, through western blot or mass spectrometry, the RBPs associated with both poly(A) and non-poly(A) RNA. A year later, David Tollervey's team developed a similar technique to 2C called total RNA-associated protein purification (TRAPP), which use silica beads to identify RBPs in the yeast Saccharomyces cerevisiae and the bacterium E. coli [START_REF] Shchepachev | Defining the RNA interactome by total RNA-associated protein purification[END_REF].

Protein-centric techniques

Protein-centric approaches are used to identify the RNA targets of a protein of interest. These approaches mostly rely on immunoprecipitation techniques followed by the detection of RNA interacting with the protein of interest, typically through sequencing or RT-qPCR.

Immunoprecipitation-dependent methods

A commonly used approach to analyse RBPs in cells is the RIP (RNA Immunoprecipitation). This method involves immunoprecipitating a protein of interest along with the associated RNA. The RNAs are then precipitated, purified and sequenced. The identified RNAs need to be further validated by RT-qPCR. To preserve RNA-RBP complexes, immunoprecipitation is performed using low-stringency washes, which may potentially induce non-specific interactions.

To overcome this specificity issue, CLIP (Cross-linking and Immunoprecipitation) was developed by the team of Robert B Darnell in 2003 and is now the most widely used method for identifying and characterising in vivo RNA-RBP interactions [START_REF] Ule | CLIP Identifies Nova-Regulated RNA Networks in the Brain[END_REF] (Fig. 5). This technique is based on the cross-linking between protein and RNA using UV-C light, allowing the use of highly stringent washes unlike RIP. After crosslinking, cells are lysed, followed by RNAse treatment. The protein of interest, along with the associated RNA, is immunoprecipitated using a specific antibody against the protein of interest. The protein-RNA complexes are then radioactively labelled at the 5' end of the RNA and migrated on a denaturing SDS-PAGE. The complexes are transferred onto a nitrocellulose membrane, allowing the visualization of the RNA-RBP complex through autoradiography. In cases like CLIP-seq, the RNA bound to the RBP of interest, can also be extracted. For this purpose, the membrane is cut, proteins are digested and the bound RNAs are purified. After creating a cDNA library, the nucleic acids can be sequenced to identify the RNA target interacting with the RBP of interest [START_REF] Ule | CLIP: A method for identifying protein-RNA interaction sites in living cells[END_REF].

Immunoprecipitation-independent methods

One limitation of CLIP, or RIP, is that they are based on immunoprecipitation of the protein, which requires the use of a specific antibody. Recently immunoprecipitationindependent techniques have emerged. For example, in the targets of RNA-binding proteins identified by editing (TRIBE) technic, the target protein is fused to the catalytic domain of the RNA editing enzyme ADAR. ADAR deaminates neighbouring adenosines and the resulting deaminated bases can be identified through sequencing [START_REF] Mcmahon | TRIBE: Hijacking an RNA-Editing Enzyme to Identify Cell-Specific Targets of RNA-Binding Proteins[END_REF]. In addition to RNA editing, RNA tagging methods exist, where the RBP is fused to an enzyme that covalently adds uridines to the end of the RNA, which can also be identified through sequencing [START_REF] Lapointe | Protein-RNA networks revealed through covalent RNA marks[END_REF].

In summary, the methods used to study RNA-protein associations provide valuable insights into characterising interactions and understanding their roles within the cell.

These powerful tools are developed at all levels: in silico, in vitro, in cellulo and in vivo.

New methods or derivatives of original methods are constantly emerging to search for more specificity. For example, a very recent study has introduced a new technique specifically designed to identify single-strand RBPs: the kethoxal assisted singlestrand RNA interactome capture (KASRIC) [START_REF] Zhao | Transcriptome-wide identification of single-stranded RNA binding proteins[END_REF]. In addition, alongside these experimental approaches, numerous bioinformatics methods have been developed to predict RNA-protein interactions [START_REF] Corley | How RNA-Binding Proteins Interact with RNA: Molecules and Mechanisms[END_REF]. For instance, RNAbindPlus is a web server designed to predict RNA-binding residues within a given protein sequence. On the contrary, RBPmap searches for RBP-binding motifs within a given RNA sequence. These in silico tools help to determine the binding sites between protein and RNA. The development of the modelling methods and their reliability takes time and is not always straightforward to implement [START_REF] Wei | Protein-RNA interaction prediction with deep learning: structure matters[END_REF].

However, an increasing number of computational techniques are emerging, and recently, AlphaFold and RoseTTAFold that enable the accurate prediction of protein and nucleic acid structure, has revolutionized the field of structural biology [START_REF] Baek | Accurate prediction of protein structures and interactions using a three-track neural network[END_REF][START_REF] Jumper | Highly accurate protein structure prediction with AlphaFold[END_REF].

II -DNA replication: Focus on the lagging strand

DNA replication is an essential, highly conserved and tightly regulated process that follows orderly steps occurring in the S-phase of the cell cycle. This semi-conservative process ensures the accurate duplication of the genetic material, from the mother cell to the daughter cell containing one original and one newly synthesized strand, prior cell division. The fidelity of this mechanism is therefore a crucial step in maintaining genome integrity. Indeed, this process involves many proteins, and any errors or defects can lead to genomic instability and disease (Burgers and Kunkel, 2017;[START_REF] Chai | Interaction and stimulation of human FEN-1 nuclease activities by heterogeneous nuclear ribonucleoprotein A1 in alphasegment processing during Okazaki fragment maturation[END_REF][START_REF] Gaillard | Replication stress and cancer[END_REF][START_REF] Zeman | Causes and consequences of replication stress[END_REF].

In eukaryotes, DNA replication is bidirectional, with replication occurring at two replication forks moving in opposite directions. One strand, the leading strand, is synthesized continuously in the direction of replication fork movement. The other strand, the lagging strand, is synthesized discontinuously in the opposite direction, creating Okazaki fragments that are later joined to form a continuous strand. The replication of the lagging strand is a mechanism that is still not fully understood in human. DNA replication process can be divided into three main stages: initiation at replication origins, synthesis of DNA at replication forks known as elongation, and termination at the site where two converging forks meet [START_REF] Dewar | Mechanisms of DNA replication termination[END_REF][START_REF] Fragkos | DNA replication origin activation in space and time[END_REF][START_REF] Kang | Eukaryotic DNA replication: Orchestrated action of multi-subunit protein complexes[END_REF]Zaher et al., 2018). Each stage will be described below, with a particular focus on the lagging strand.

Initiation

To start DNA replication, an origin of replication (ORI) is required. In mammals, several ORIs are distributed along the chromosomes. Their nature is poorly defined, as ORIs do not seem to have consensus sequences. However, a genome-wide study in 2008 localized ORIs in regions close to CpG islands [START_REF] Cadoret | Genome-wide studies highlight indirect links between human replication origins and gene regulation[END_REF]. Only 20 to 30% of ORIs will be activated during the S-phase. Indeed, several ORIs are dormant and used to rescue stalled replication forks during replicative stress [START_REF] Courtot | 53BP1 Integrates DNA Repair and p53-Dependent Cell Fate Decisions via Distinct Mechanisms[END_REF].

The firing phase follows the licensing phase and enables DNA synthesis to be activated. During the G1/S-phase transition, the MCM2-7 complex is phosphorylated by the Dbf4-dependent kinase (DDK) and cyclin-dependent kinase (CDK). This leads to the activation of the MCM7 double hexamer and the binding of the cell division cycle 45 protein (CDC45) and GINS factors (from the Japanese "go-ichi-ni-san"

meaning "5-1-2-3", after the four related subunits of the complex Sld5, Psf1, Psf2 and Psf3) to the MCM complex, forming the CMG complex [START_REF] Heller | Eukaryotic origindependent DNA replication in vitro reveals sequential action of DDK and S-CDK kinases[END_REF]. The formation and activation of the CMG complex are also favoured and mediated by the phosphorylation of the RecQL4 helicase, DNA polymerase ε and treslin, by CDKs. The origins of replication are then activated, and the pre-replication complexes are converted into pre-initiation complexes. These complexes will unwind the doublestrand DNA and replication factors will be recruited to form the replisome, such as replication protein A (RPA) which bind the single-strand DNA, replicative polymerases, replication factor C (RFC) and the proliferating cell nuclear antigen (PCNA) that acts as a processivity factor for DNA polymerases. Then, the replisome starts converting the pre-initiation complex into two replication forks moving in opposite directions (Fig. 7).

Elongation

In the 1960s, Okazaki and colleagues demonstrated the existence of a leading strand and a lagging strand [START_REF] Okazaki | Mechanism of DNA chain growth. I. Possible discontinuity and unusual secondary structure of newly synthesized chains[END_REF]. The leading strand is synthesized in the same direction as the fork progression, while the lagging strand is synthesized in the opposite direction, hence discontinuously [START_REF] Kang | Eukaryotic DNA replication: Orchestrated action of multi-subunit protein complexes[END_REF].

The first step of DNA synthesis is common to both strands and begins with the action of Primase-DNA polymerase α (PRIM-Polα) complex. The primase, containing a catalytic subunit PRIM1 and a regulatory subunit PRIM2, initiates DNA synthesis by generating a 5' RNA primer of about 7 to 12 ribonucleotides in length. Subsequently, the DNA polymerase Polα, incorporates 20 to 25 nucleotides, in the 5' to 3' direction,

following this RNA primer. The RNA-DNA primer enables replication elongation on both strands [START_REF] Jain | Eukaryotic DNA polymerases[END_REF][START_REF] Sun | Okazaki fragment maturation: DNA flap dynamics for cell proliferation and survival[END_REF]) (Fig. 8).

On the leading strand, the PRIM-Polα complex is then replaced by DNA polymerase ε (Polε) to continue DNA synthesis. On the other hand, on the lagging strand, DNA polymerase δ (Polδ) synthesizes, discontinuously as the polarity of the parental strand is inverted, approximately 100-200 nucleotides, forming the Okazaki fragments (Burgers and Kunkel, 2017;[START_REF] Mcelhinny | Division of Labor at the Eukaryotic Replication Fork[END_REF]. To form a functional doublestrand DNA, these Okazaki fragments need to be processed and RNA primers to be removed. Several mechanisms for maturing Okazaki fragments exist and are described below.

During each division of a human cell, tens of millions of Okazaki fragments are produced and ligated. DNA combing analyses, based on double incorporation of thymidine analogues followed by fiber stretching (Quinet et al., 2017a), have shown that forks travel approximately at 1-3 kb per min, depending on cell type (Conti et al., 2007). This means that it takes only a few seconds to synthesize a single Okazaki fragment [START_REF] Burgers | Solution to the 50-year-old Okazaki-fragment problem[END_REF][START_REF] Zheng | Okazaki fragment maturation: nucleases take centre stage[END_REF]. is therefore considered to be an error-prone polymerase, that particularly generates insertions and deletions (Guilliam et al., 2015). Polα synthesizes approximately 10%

of the genome during DNA replication, and the errors potentially generated are therefore deemed "acceptable". Furthermore, considering all polymerases together, it is estimated that there is one error per 10 4 -10 5 nucleotides incorporated [START_REF] Ganai | DNA Replication-A Matter of Fidelity[END_REF]. Moreover, Polδ can displace the strand, an activity necessary for the maturation of Okazaki fragments, whereas Polε has a very limited capacity and can only move one or two nucleotides. The potential strand-displacement activity of Polα is not well defined. It seems that this polymerase has the ability to displace the DNA strand but only in the presence of manganese ions (Mn 2+ ) [START_REF] Boldinova | Strand Displacement Activity of PrimPol[END_REF] (Table 3).

Fidelity

Error-prone Error-free Error-free

Strand displacement

Conditional Yes Limited

Okazaki fragments maturation

Flap processing pathways

The mechanism of Okazaki fragments processing and all the players involved are still not well described. However, in vitro and recent in vivo data have enabled to model this process, especially in yeast. It is accepted that the removal of the primer and ligation of the Okazaki fragments can occur through two pathways: the short flap and the long flap pathway (Fig. 9) [START_REF] Rossi | Reconstituted Okazaki fragment processing indicates two pathways of primer removal[END_REF][START_REF] Sun | Okazaki fragment maturation: DNA flap dynamics for cell proliferation and survival[END_REF]Zaher et al., 2018).

Short flap pathway

Okazaki fragment synthesis is an extremely rapid process, meaning that Polδ, in complex with PCNA, frequently encounters the downstream Okazaki fragment, forcing it to displace the RNA primer. The RNA primer displacement induces a 5' single-strand RNA structure, known as the flap. This flap is short and is approximatively 2 to 10 nucleotides. Polδ does not have an endonuclease activity, therefore the Flap endonuclease 1 (FEN1) cleaves the flap nucleotide by nucleotide, inducing a DNA nick.

The two adjacent Okazaki fragments are joined by the DNA ligase 1 (LIG1) to produce a functional double-strand DNA (Raducanu et al., 2022).

Long flap pathway

In some cases, cleavage by FEN1 is delayed, resulting in the formation of a long flap, whose length is greater than 20 nucleotides. The exact reason for the delayed cleavage by FEN1 is still not well defined. It could be due to a FEN1 protein defect or the presence of RPA. Indeed, RPA binds to the single-strand DNA part of the flap, preventing the action of FEN1. The DNA replication helicase/nuclease 2 (DNA2) is then recruited and its nuclease activity is stimulated by RPA. DNA2 cleaves the long flap in the middle to shorten it and thus obtaining a 5' short flap of less than 10

The long flap pathway would thus be favoured in transcriptionally active regions. In 2010, Lata Balakrishnan and colleagues demonstrated in vitro that the histone acetyltransferase p300 plays a role in the switch between the long and short flap pathways. Specifically, p300 acetylates FEN1, leading to its inhibition, and also acetylates DNA2, stimulating its activity. Thereby these two simultaneous actions of p300 promote the long flap pathway. Furthermore, p300 is known to be enriched in an active chromatin environment to assist the transcription machinery in accessing DNA. In addition to this role, p300 would therefore ensure replication fidelity in these transcriptionally active regions, by promoting the long flap pathway (Balakrishnan and Bambara, 2011;Balakrishnan et al., 2010).

Defects in Okazaki fragments maturation

It is essential for the cell to properly mature the Okazaki fragments. Indeed, alterations in Okazaki fragments processing lead to genetic instability and are often associated with human pathologies or cancers. For example, mutations in FEN1, leading to a dysfunction in its nuclease activity, are associated with autoimmunity, chronic inflammation or cancers with aberrant chromosome numbers (Zheng et al., 2007(Zheng et al., , 2011[START_REF] Zheng | Multiple roles of DNA2 nuclease/helicase in DNA metabolism, genome stability and human diseases[END_REF].

In case of a defect, the cell has several mechanisms for the recognition and the processing to maintain trouble-free replication. These defects can be induced by the loss of maturation factors or by genotoxic agents, resulting in an ssDNA gap between two unligated Okazaki fragments. PARP1 recognises these gaps on the lagging strand and binds to the resulting single-strand DNA, leading to PARylation of PARP1 and surrounding proteins. In human cells, Hana Hanzlikova and colleagues demonstrated that this increase in S-phase PAR, led by the depletion of FEN1 or LIG1, is not related to DNA damage or replication stress. Moreover, the PARP activity facilitates the recruitment of the single-strand break repair protein XRCC1 and its partner LIG3, resulting in a backup processing of Okazaki fragments. Therefore the accumulation of PAR in S-phase reflects a defect in the lagging strand maturation (Hanzlikova et al., 2018).

PARP1 inhibitors (PARPi) are commonly used in the clinic, such as olaparib which induces PARP1 trapping on DNA and therefore inhibits the ssDNA break repair signalling. These inhibitors can also disrupt nascent DNA maturation, particularly when FEN1 is depleted (Vaitsiankova et al., 2022). For example, the synthetic lethality using PARPi is frequently employed to target BRCA-deficient cancers. Indeed, loss of BRCA leads to an increase in the ssDNA replication gaps targeted by PARPi (Cong et al., 2021;[START_REF] Panzarino | Replication gaps underlie BRCA deficiency and therapy response[END_REF]. In these genetic backgrounds, the loss of 53BP1 leads to resistance against PARPi. To overcome this resistance, depleting LIG3 restores the PARPi-sensitive phenotype (Paes Dias et al., 2021).

Termination

When two converging forks meet, the termination step can begin (Fig. 10).

The first step is to resolve the topological tensions. The topological stress is resolved by the production of pre-catenanes and the dissolution of these catenated replication products by the action of topoisomerases.

Then it is necessary to fill and ligate the remaining gaps. The CMG complex from the leading strand of one fork passes over the 5' end of the last Okazaki fragment on the lagging strand of the converging fork and moves along the double-strand DNA [START_REF] Dewar | Mechanisms of DNA replication termination[END_REF].

The last step is the dissociation of the CMG complex from chromatin, by polyubiquitination of the MCM7 subunit. This dissociation notably allows for the avoidance of inappropriate protein recruitment and thus promotes a correct progression of cell cycle. The proteins responsible for poly-ubiquitination are not clearly identified in mammals [START_REF] Kang | Eukaryotic DNA replication: Orchestrated action of multi-subunit protein complexes[END_REF]. Overall, the termination process is not yet fully understood, and many questions remain to be answered.

Replication stress

DNA replication is therefore a complex and highly regulated process. It is important for the cell to maintain faultless, stress-free replication before continuing its progression through the cell cycle. Indeed, defects during DNA replication are a major source of genetic instability, which is also a hallmark of cancer [START_REF] Gaillard | Replication stress and cancer[END_REF][START_REF] Hanahan | Hallmarks of cancer: The next generation[END_REF][START_REF] Macheret | DNA Replication Stress as a Hallmark of Cancer[END_REF].

Cause of replication stress

The cell is constantly subjected to stress that must be resolved. When this stress occurs during the S-phase of the cell cycle, it results in a slowdown or halt of replication forks. When a fork is stalled, it temporarily paused to remove the stress, and then it either restarts or collapses. There are several sources of replicative stress, both endogenous and exogenous to the cell (Fig. 11).

Endogenous sources include DNA damage (e.g. double-strand breaks, single-strand DNA gaps), incorporation of ribonucleotide, unusual DNA structures (e.g. Gquadruplexes) and transcription-replication conflicts (e.g. RNA:DNA hybrids) [START_REF] Zeman | Causes and consequences of replication stress[END_REF]. A deficiency in replication factors, the presence of fragile sites, an insufficiency of the pool of deoxyribonucleotide triphosphates (dNTPs) or a lack of activation of replication origins can also be a source of stress [START_REF] Briu | Replication Stress, Genomic Instability, and Replication Timing: A Complex Relationship[END_REF][START_REF] Willaume | A Link between Replicative Stress, Lamin Proteins, and Inflammation. Genes (Basel)[END_REF].

Exogenous sources correspond to genotoxic agents, used in chemotherapy or in research laboratories. There are several types of these drugs, such as molecules that block the synthesis of deoxyribonucleotide by targeting the nucleotide metabolism (e.g. hydroxyurea) or activity of polymerases (e.g. aphidicolin), those that cause DNA damage or interfere with DNA repair signalling (e.g. cisplatin, olaparib) and those that interfere with topoisomerases (e.g. etoposide, camptothecin) [START_REF] Vesela | Common Chemical Inductors of Replication Stress: Focus on Cell-Based Studies[END_REF]. The cell must therefore implement appropriate responses to manage these sources of stress effectively.

Once the fork is stabilized, several mechanisms can be employed to restart it (Fig. 12), such as the activation of dormant origins, the regression of the fork or the activation of DNA damage tolerance pathways [START_REF] Neelsen | Replication fork reversal in eukaryotes: from dead end to dynamic response[END_REF][START_REF] Pasero | Nucleases Acting at Stalled Forks: How to Reboot the Replication Program with a Few Shortcuts[END_REF]Quinet et al., 2017b). Conversely, if there is a defect in fork stabilization, the fork collapses as proteins from the replisome dissociate, leading to a double-strand break. Therefore, the players of homologous recombination come into play to repair the DNA.

DNA damage tolerance

To avoid the consequences of a blocked fork, there are mechanisms of DNA damage tolerance. These mechanisms stabilize the fork and allow it to continue progression, even in the presence of lesions (Fig. 12). Three mechanisms are defined. First is template switching (TS), which uses the newly synthesized sister chromatid and is thus an error-free mechanism. In contrast, translesion synthesis (TLS) is an errorprone pathway based on the ability of a DNA polymerase to replicate while bypassing a DNA lesion. TLS is often associated with a risk of mutagenesis. Finally, the last mechanism is the repriming mechanism by PRIM-Polα complex, which involves reinitiating replication downstream of the lesion using its DNA primase activity, leaving a ssDNA gap to be filled later [START_REF] Berti | Replication stress: getting back on track[END_REF][START_REF] Pasero | Nucleases Acting at Stalled Forks: How to Reboot the Replication Program with a Few Shortcuts[END_REF][START_REF] Quinet | To skip or not to skip: choosing repriming to tolerate DNA damage[END_REF][START_REF] Willaume | A Link between Replicative Stress, Lamin Proteins, and Inflammation. Genes (Basel)[END_REF].

Homologous recombination-mediated replication fork restart

A stalled replication fork can result in a double-strand break (DSB). There are several pathways for the repair of double-strand breaks, depending on the cell cycle (Fig. 14).

Homologous recombination (HR) is favoured in post-replication and the nonhomologous end joining (NHEJ) pathway is promoted in pre-replication. Unlike NHEJ, HR is a high-fidelity DSB repair pathway that functions in G2/S-phase because this pathway requires the availability of a homologous copy of the DSB-containing region,

i.e. the sister chromatid. In this pathway, DSBs are recognized by the MRE11/RAD50/NBS1 complex (MRN). MRE11 is responsible for the resection of the DSB end. To achieve this, MRE11 binds RAD50, which is associated with DNA, to maintain the DNA resected by MRE11. It also binds NBS1, which recruits the ATM kinase and the CtIP protein. The recruitment of CtIP will activate the endonuclease activity of MRE11 and thus initiate the resection [START_REF] Daley | Biochemical mechanism of DSB end resection and its regulation[END_REF][START_REF] Sun | Structural basis of homologous recombination[END_REF].

To extend the resection, other enzymes are required such as the exonuclease EXO1.

The resulting single-strand DNA end is bound and stabilized by RPA. The BRCA1/BRCA2 complex will allow the exchange of RPA by RAD51 [START_REF] Carreira | The BRC repeats of BRCA2 modulate the DNA-binding selectivity of RAD51[END_REF]. Subsequently, a RAD51 filament is obtained and stretched to look for sequence homology [START_REF] Renkawitz | Mechanisms and principles of homology search during recombination[END_REF]. Once the homology is found, a double Holliday junction is formed. The repaired DNA will then contain or not crossovers [START_REF] Willaume | A Link between Replicative Stress, Lamin Proteins, and Inflammation. Genes (Basel)[END_REF].

A defect in HR decreases the replication fork progression speed [START_REF] Daboussi | A homologous recombination defect affects replication-fork progression in mammalian cells[END_REF]. Indeed, HR is necessary to protect and repair the replication forks. When the fork is stalled, RAD51 bound to single-strand DNA protects the nascent DNA from excessive degradation by nucleases like MRE11. BRCA2 also plays a role in protection as it stabilizes RAD51 to the DNA [START_REF] Liu | Two replication fork remodeling pathways generate nuclease substrates for distinct fork protection factors[END_REF].

When the stalled fork fails to restart or when the processing of the fork is prolonged, it becomes a substrate for structure-specific endonucleases which generate singlestrand DSBs [START_REF] Dehé | Control of structure-specific endonucleases to maintain genome stability[END_REF]. Subsequently, these breaks undergo repaired by the break induced replication (BIR) pathway, which is a highly mutagenic HR mechanism [START_REF] Costantino | Break-induced replication repair of damaged forks induces genomic duplications in human cells[END_REF].

In conclusion, DNA replication is a common process to all organisms that enables the transmission of genetic material. Disturbances to this process, whether from endogenous or exogenous agents, pose a challenge for the cell that must be addressed correctly to maintain the integrity of the genome. Mechanisms such as DNA damage tolerance, fork reversal and homologous recombination help to stabilize, repair and restart the replication forks. Furthermore, replication errors generated by replicative polymerases are repaired by the mismatch repair (MMR) pathway, which is not detailed in this introduction. In this process, the lesion is first recognized, then the incorrect DNA sequence is excised and finally the DNA is resynthesized without errors. Overall, replication mechanisms, and notably the replication of the lagging strand through Okazaki fragments, in both stressed and unstressed conditions are complex and not yet fully understood. However, an increasing number of new factors are regularly being identified as replication players.

p53 transcriptional response

Although the interaction between 53BP1 and p53 has been known for decades, the role of 53BP1 in p53 signalling remained undefined until recently. p53 is a transcription factor that control a multitude of biological mechanisms such as cellcycle arrest, apoptosis, DNA repair and cellular stress responses [START_REF] Amelio | Context is everything: extrinsic signalling and gain-of-function p53 mutants[END_REF][START_REF] Vousden | Blinded by the Light: The Growing Complexity of p53[END_REF]. 53BP1 mediates p53-dependant cell cycle progression by regulating cell-cycle checkpoint. This 53BP1-p53 pathway is mediated by the ubiquitin specific peptidase 28 (USP28) (Fig. 15). For instance, in the event of centrosome loss or prolonged mitosis, 53BP1 interacts with the deubiquitinase USP28 that can deubiquitinate p53 to stabilize it. Therefore, p53 induces the transcription of target genes (e.g. p21) to arrest cell growth [START_REF] Fong | 53BP1 and USP28 mediate p53-dependent cell cycle arrest in response to centrosome loss and prolonged mitosis[END_REF][START_REF] Lambrus | A USP28-53BP1-p53-p21 signaling axis arrests growth after centrosome loss or prolonged mitosis[END_REF].

Moreover, in response to ionizing radiation, Raquel Cuella-Martin and colleagues demonstrated that 53BP1 also promotes the p53-dependent transcriptional response (Cuella-Martin et al., 2016). Furthermore, another actor involved in the function of 53BP1 in p53 signalling is the AHNAK protein (also known as desmoyokin meaning "giant" in Hebrew). AHNAK interacts with 53BP1 to suppress p53 activity.

Consequently, depletion of AHNAK increases 53BP1-p53 interaction, leading to the activation of p53 and inducing apoptosis in human cancer cells [START_REF] Ghodke | AHNAK controls 53BP1-mediated p53 response by restraining 53BP1 oligomerization and phase separation[END_REF].

This regulatory role of 53BP1 on p53 is independent of its role in DNA repair.

End-joining processes

One of the most studied roles of 53BP1 today is its involvement in the repair of DSBs.

DSBs are cytotoxic lesions formed when both DNA strands are broken and the cell must repair this break to survive. The repair can occur optimally through DSB repair mechanisms, enabling the cell to continue its normal cycle of division. However, if the break is not properly repaired, it can lead to genetic instability and chromosomal abnormalities giving rise to numerous diseases, including cancers [START_REF] Jackson | The DNA-damage response in human biology and disease[END_REF].

DSBs can occur in different context: from an exogenous source (e.g. ionizing radiation, UV, genotoxic chemicals) or from a spontaneously endogenous source (e.g. replicative stress) [START_REF] Chapman | Playing the End Game: DNA Double-Strand Break Repair Pathway Choice[END_REF]. The major DSBs repair pathways are HR (homologous recombination) and c-NHEJ (canonical NHEJ).

The c-NHEJ pathway is a conserved process that promotes direct ligation of break ends. It is a fast mechanism but also error prone. Indeed, Zhiyong Mao and colleagues showed in 2008, using chromosomally integrated fluorescent reporter, that NHEJ lasts about 30min, compared with 7h for HR (Mao et al., 2008a). NHEJ is active throughout the cell cycle, favoured in G1-phase and at the G2/M transition [START_REF] Karanam | Quantitative Live Cell Imaging Reveals a Gradual Shift between DNA Repair Mechanisms and a Maximal Use of HR in Mid S Phase[END_REF][START_REF] Mao | DNA repair by nonhomologous end joining and homologous recombination during cell cycle in human cells[END_REF]. Inevitably, this speed leads to errors. For instance, the NHEJ pathway can be associated with insertions, deletions, substitutions at the break sites, and translocations if DSBs from different parts of the genome are joined [START_REF] Chang | Non-homologous DNA end joining and alternative pathways to double-strand break repair[END_REF].

After double-strand break, the first step in the NHEJ pathway is the recognition of the DNA damage site. This recognition is enabled thanks to the binding of the Ku70/80 heterodimer [START_REF] Soutoglou | Positional stability of single double-strand breaks in mammalian cells[END_REF]. Since most DSBs have incompatible ends, nuclease activity is required. Ku70/80, which binds to DNA without sequence preference, serves as a platform for the recruitment of the catalytic DNA-PK subunit (DNA-PKCs). The association of Ku/DNA-PKcs induces the autophosphorylation of DNA-PKcs and the phosphorylation of target proteins such as the endonuclease Artemis. Artemis will therefore generate regions of microhomologies (< 4 nucleotides)

to facilitate end joining. Then, polymerases will be recruited to fill in the doublestrand DNA. Finally, the DNA Ligase 4 (LIG4) and the X-ray repair crosscomplementing protein 4 (XRCC4) complex join the DNA ends [START_REF] Lieber | The Mechanism of Human Nonhomologous DNA End Joining *[END_REF][START_REF] Riballo | A Pathway of Double-Strand Break Rejoining Dependent upon ATM, Artemis, and Proteins Locating to γ-H2AX Foci[END_REF]) (Fig. 16).

53BP1 is not considered a core protein of the NHEJ pathway, however it is involved upstream of this pathway and promotes its signalling in certain context. 53BP1 also mediates programmed NHEJ in the context of telomere fusion, V(D)J recombination and CSR (Class Switch Recombination). For example, the oligomerisation of 53BP1 is required for CSR signalling [START_REF] Bothmer | Regulation of DNA End Joining, Resection, and Immunoglobulin Class Switch Recombination by 53BP1[END_REF]. The depletion of 53BP1 also induces a defect in V(D)J recombination with extensive degradation [START_REF] Difilippantonio | 53BP1 facilitates long-range DNA endjoining during V(D)J recombination[END_REF]. Moreover, in a deprotected telomere context, the recruitment of 53BP1

to damaged chromosome end is essential to facilitate repair by NHEJ [START_REF] Dimitrova | 53BP1 promotes nonhomologous end joining of telomeres by increasing chromatin mobility[END_REF].

Double-strand break repair choice

53BP1 has therefore become known for its significant function in DSB repair through the NHEJ pathway, and also for its role in the choice of DSB repair signalling [START_REF] Daley | 53BP1, BRCA1, and the choice between recombination and end joining at DNA double-strand breaks[END_REF][START_REF] Lei | Multifaceted regulation and functions of 53BP1 in NHEJ-mediated DSB repair (Review)[END_REF]) (Fig. 16). 53BP1 possesses an anti-resection activity, thus protecting damaged ends from resection by nucleases, an essential step for HR.

The antagonist of 53BP1 is BRCA1, associated with its partner BARD1, which promotes this DNA resection. The deletion of BRCA1 is a driver event in the development of breast cancers as it induces HR deficiency. The same applies to BARD1. In these genetic backgrounds, the loss of 53BP1 rescues HR, leading to resistance of cancer cells to anti-cancer agents [START_REF] Becker | BARD1 reads H2A lysine 15 ubiquitination to direct homologous recombination[END_REF][START_REF] Bouwman | 53BP1 loss rescues BRCA1 deficiency and is associated with triple-negative and BRCA-mutated breast cancers[END_REF][START_REF] Bunting | 53BP1 inhibits homologous recombination in Brca1-deficient cells by blocking resection of DNA breaks[END_REF][START_REF] Jaspers | Loss of 53BP1 causes PARP inhibitor resistance in Brca1-mutated mouse mammary tumors[END_REF].

The NHEJ pathway is the primary choice for repairing DSBs as it can be activated during any phase of the cell cycle, triggered by phosphorylation of 53BP1. In the Sphase, BRCA1 promotes the HR pathway by facilitating the dephosphorylation of 53BP1 through the phosphatase PP4C leading to the release of RIF1 [START_REF] Isono | BRCA1 Directs the Repair Pathway to Homologous Recombination by Promoting 53BP1 Dephosphorylation[END_REF]. In fact, as soon as 53BP1 is phosphorylated by ATM, it recruits RIF1 and the phosphatase PP1, thus forming a barrier to DNA end resection in G1-phase by inhibiting the recruitment of MRN and CtIP [START_REF] Chapman | RIF1 Is Essential for 53BP1-Dependent Nonhomologous End Joining and Suppression of DNA Double-Strand Break Resection[END_REF][START_REF] He | DYNLL1 binds to MRE11 to limit DNA end resection in BRCA1deficient cells[END_REF].

Moreover, 53BP1 can also interact with DYNLL1 that physically inhibits the resection machinery [START_REF] He | DYNLL1 binds to MRE11 to limit DNA end resection in BRCA1deficient cells[END_REF]. 53BP1 therefore prevents early resection, however if DNA resection has already been initiated, 53BP1 can also act to inhibit extended resection by interacting with RIF1, PTIP and Shieldin. Subsequently, Shieldin recruits CTC1-STN1-TEN1(CST)/Polα/Primase inducing PRIM-Polα-dependent 3' end fill-in DNA synthesis [START_REF] Gupta | DNA Repair Network Analysis Reveals Shieldin as a Key Regulator of NHEJ and PARP Inhibitor Sensitivity[END_REF][START_REF] Mirman | 53BP1-shieldindependent DSB processing in BRCA1-deficient cells requires CST-Polα-primase fill-in synthesis[END_REF][START_REF] Noordermeer | The shieldin complex mediates 53BP1-dependent DNA repair[END_REF].

The choice of DSB repair pathways also relies on chromatin structure and histone marks. 53BP1 recognizes histone marks H4K20me2, methylated by MMSET, and H2AK15ub, ubiquitinated by the RNF8-RNF168 pathway, at double-strand breaks, enabling it to play its role in inhibiting resection [START_REF] Fradet-Turcotte | 53BP1 is a reader of the DNA-damage-induced H2A Lys 15 ubiquitin mark[END_REF][START_REF] Pei | MMSET regulates histone H4K20 methylation and 53BP1 accumulation at DNA damage sites[END_REF]. In S-phase, however, the H4K20me2 motif is less present at chromatin because it is diluted after the synthesis of new nucleosomes incorporating into replicated DNA. The recognition of H4K20me0 by BRCA1/BARD1 therefore prevents the anti-resection activity of 53BP1 and promotes HR [START_REF] Nakamura | H4K20me0 recognition by BRCA1-BARD1 directs homologous recombination to sister chromatids[END_REF]) (Fig. 15).

Replication stress response

Recent proteomic studies have identified NHEJ factors as being active during DNA replication, when replicative stress is induced (Dungrawala et al., 2015;[START_REF] Mukherjee | RIF1 promotes replication fork protection and efficient restart to maintain genome stability[END_REF]Ribeyre et al., 2016). This is indeed the case for 53BP1 which also plays a role in the S-phase during the response to replication stress (Fig. 15).

In 2004, Sagar Sengupta and colleagues demonstrated that following HU treatment, which reduces the dNTPs pool and thus induces replicative stress, 53BP1 is recruited to nascent DNA. Once recruited, 53BP1 facilitates the accumulation of BLM and p53 at stalled forks, in a CHK1-and ATR-dependent manner [START_REF] Sengupta | Functional interaction between BLM helicase and 53BP1 in a Chk1-mediated pathway during S-phase arrest[END_REF].

Moreover, the recruitment of 53BP1 to the fork is a fast process, approximately 5min after the induced stress (Dungrawala et al., 2015).

The role of 53BP1 at the stalled fork is actually also based on its anti-resection activity. In 2018, Joonyoung Her and colleagues observed that in primary B cells, the depletion of 53BP1 resulted in the degradation of nascent DNA upon HU treatment.

However, the interactions that 53BP1 establishes to mediate its functions in DNA replication are still unclear. Her et al. propose that 53BP1 interacts with histones

H4K20me2 and H2AK15Ub at the stalled fork to activate ATR-mediated CHK1 phosphorylation and thus stabilize the fork. Moreover, 53BP1 protects the fork by limiting access to the MRE11 exonuclease, thereby inhibiting its activity (Her et al., 2018).

Furthermore, following a replicative stress, the loss of 53BP1 reduces replication rates, the cell survival and increases chromosomal aberration (Tripathi et al., 2007;Villa et al., 2018).

53BP1 is also involved in the fork reversal mechanism. There are 2 pathways to protect the regressed fork end: one is 53BP1-dependent and the other is independent of 53BP1. 53BP1 protects specifically the substrate generated by the F-Box DNA helicase 1 (FBH1), associated with BRCA2 and RAD51 [START_REF] Liu | Two replication fork remodeling pathways generate nuclease substrates for distinct fork protection factors[END_REF].

Additionally, similar to the mechanism of choosing the appropriate DSB repair pathways, 53BP1 competes with BRCA1 in selecting the pathway for the restart of stalled replication fork. 53BP1 in association with RIF1 promotes a pathway without cleavage, whereas BRCA1 favours a slower pathway with cleavage, known as the break-induced replication (BIR) pathway [START_REF] Xu | 53BP1 and BRCA1 control pathway choice for stalled replication restart[END_REF].

Taken together, these studies show that 53BP1, like its anti-resection role in DSB repair, controls nascent DNA degradation by acting as a nuclease barrier and thus preventing resection of stalled replication forks after induction of replication stress.

Structure

Human 53BP1 is a protein, with no enzymatic activity, of 1972 amino acids. It contains several domains (Fig. 17) that can be divided into 3 main parts [START_REF] Panier | Double-strand break repair: 53BP1 comes into focus[END_REF][START_REF] Rass | 53BP1: Keeping It under Control, Even at a Distance from DNA Damage[END_REF]:

-The N-terminal part which plays a role in 3' overhangs control following DNA damage. This region is composed of 28 Serine-Threonine-Glutamine (STQ) sites that can be phosphorylated by ATM/ATR and are known to interact with RIF1 and PTIP proteins to control resection and promote the NHEJ repair pathway.

-A central part which is the minimum focus forming region and therefore is involved in the recruitment of 53BP1 to double-strand break sites. This region includes the Oligomerization Domain (OD), the Glycine-Arginine Rich (GAR) domain, two Tudor domains, the Ubiquitin-Dependent Recruitment (UDR) motif and the Nuclear

53BP1 and RNA

As described in the section I-1, numerous studies have shown that RBPs and RNAs play a role in the regulation of the response to DNA damage. Furthermore, some proteins in the DNA damage response, such as Ku, have been characterised as having the ability to bind RNA. As mentioned above, since 53BP1 contains a putative RNAbinding domain with GAR motifs (RGRGRRGR) (Thandapani et al., 2013) (Fig. 17), it therefore raises the question of whether 53BP1 also possesses an RNA-binding activity. Over the past few years, several studies have attempted to provide answers to this question.

In 2005, Fiona Pryde and colleagues have observed that the formation of 53BP1 foci at DNA damage sites is mediated by an RNA component. After induction of doublestrand breaks in murine (NIH3T3) and human (HeLa) cells, by exposure to ionizing radiation, RNAse A treatment, that cleaves single-strand RNA, decreased the recruitment of 53BP1 at DNA break sites (Pryde et al., 2005).

These results were also obtained in another study in 2012 where Francia et al, showed that after irradiation in human cells (HeLa), the RNAse A treatment leads to a decrease in 53BP1 foci (Francia et al., 2012). Moreover, the inhibition of the ribonucleases Dicer and Drosha, that are responsible for the production of small double-strand RNAs in cells, also decreased 53BP1 recruitment to DNA damage. The team therefore concluded that at DNA damage sites RNAs are produced and cleaved by Dicer and Drosha and they play a role in the activation of DNA repair signalling.

The production of DNA damage response RNA (DDRNA) at DNA damage sites was subsequently confirmed (Michelini et al., 2017(Michelini et al., , 2018)). After DNA break, MRN is recruited, triggering the transcription by RNA polymerase II (RNA Pol II) on each side of the break. The RNA Pol II then synthesizes damage-induced long non-coding RNAs (dilncRNAs) [START_REF] Pessina | Functional transcription promoters at DNA double-strand breaks mediate RNA-driven phase separation of damage-response factors[END_REF]. These RNAs contribute to the formation of doublestrand RNAs, the DDRNA precursors. These precursors are subsequently cleaved by Dicer and Drosha to generate DDRNAs involved in DDR signalling, by recruiting factors such as 53BP1. The mechanism of obtaining DDRNA precursors is still poorly described (Audoynaud et al., 2021) (Fig. 18).

Furthermore, 53BP1 is negatively regulated by the RNA-binding protein Tudorinteracting repair regulator (TIRR). Under normal conditions, TIRR binds to 53BP1 and inhibits its activity. By binding to the Tudor domain of 53BP1, TIRR masks the methylated histone-binding surface, preventing 53BP1 from being recruited and therefore from acting. TIRR also inhibits the formation of the 53BP1-p53 complex and the loss of TIRR leads to an abnormal increase in the transcriptional response mediated by p53 [START_REF] Botuyan | Mechanism of 53BP1 activity regulation by RNAbinding TIRR and a designer protein[END_REF][START_REF] Drané | TIRR regulates 53BP1 by masking its histone methyl-lysine binding function[END_REF][START_REF] Parnandi | TIRR inhibits the 53BP1-p53 complex to alter cell-fate programs[END_REF]. Upon deletion of the GAR domain does not impact 53BP1 foci. After RNAse A treatment, the recruitment of 53BP1 deleted from the GAR domain decreases only at early stages (20min after irradiation) and not at late stages (2h after irradiation). This suggests that the GAR domain helps the formation of 53BP1 foci but is not essential for its recruitment. This has been confirmed in a 2013 study where Lottersberger et al., studied the phenotype associated with 53BP1 mutated in the GAR domain (three Arginines substituted in three Lysines). They showed that the GAR mutant had no impact on the various functions of 53BP1, e.g. in DNA repair signalling, in class switch recombination and in blocking the resection at dysfunctional telomeres [START_REF] Lottersberger | Role of 53BP1 oligomerization in regulating double-strand break repair[END_REF]. The GAR domain is therefore not essential in most functions of 53BP1. However, mutations in the GAR domain have not been studied in the context of DNA replication, so it cannot be excluded that this domain is required in this context.

Even though no direct binding between 53BP1 and RNA has yet been demonstrated, recent proteomic analyses have suggested 53BP1 as a potential RBP. Indeed, using the OOPS assay, it was shown that a 53BP1 peptide could associate with RNA [START_REF] Queiroz | Comprehensive identification of RNAprotein interactions in any organism using orthogonal organic phase separation (OOPS)[END_REF]Trendel et al., 2019). However, these observations do not demonstrate a direct interaction between the 53BP1 protein and RNA.

In conclusion, the studies described above show that the role of 53BP1 in its DNA repair function is dependent on an RNA component and that 53BP1 could be characterised as an RBP. These findings raise two questions: Does 53BP1 have an RNA-binding activity and is 53PB1's function in DNA replication also mediated by RNA? The aim of this thesis was thus to address these questions.

OBJECTIVES

As described in the introduction, RNA-binding proteins are important in maintaining genome stability throughout the cell cycle by participating in DNA damage repair and response to replication stress. Additionally, an increasing number of proteins involved in the NHEJ pathway, a process of repairing double-strand break, have been suggested

to exhibit a potential RNA-binding activity (Audoynaud et al., 2021). However, the exact role of this possible activity in the functions of these proteins remains unclear.

The laboratory has explored these candidate RBPs with particular emphasis on the 53BP1 protein for several reasons.

Firstly, 53BP1 possesses an RNA-binding domain, the GAR domain rich in Arginine and Glycine where the Arginines are methylated by PRMT1 and 5 [START_REF] Boisvert | The GAR motif of 53BP1 is arginine methylated by PRMT1 and is necessary for 53BP1 DNA binding activity[END_REF][START_REF] Hwang | PRMT5 promotes DNA repair through methylation of 53BP1 and is regulated by Srcmediated phosphorylation[END_REF][START_REF] Panier | Double-strand break repair: 53BP1 comes into focus[END_REF]Thandapani et al., 2013).

Furthermore, in the context of DNA damage, the functions of 53BP1 are mediated by an RNA component. Indeed, treating cells with RNAse A significantly reduces the formation of 53BP1 foci following ionising irradiation. However, these observations do not demonstrate a direct interaction between 53BP1 and RNA. In addition, deletion or mutation of three Arginines in Lysines in the GAR domain does not affect the recruitment of 53BP1 to the lesion sites, nor does it impact other functions of 53BP1 such as class switch recombination (Francia et al., 2012;[START_REF] Lottersberger | Role of 53BP1 oligomerization in regulating double-strand break repair[END_REF]Michelini et al., 2017;Pryde et al., 2005). Nevertheless, the alteration of the GAR domain has not been tested in the functions of 53BP1 related to the response to replication stress, therefore a potential role of this domain in these functions cannot be ruled out. Indeed, in the S-phase 53BP1 is recruited to the stalled replication fork within 5min following the replication stress. This recruitment aims to protect the nascent DNA from degradation by nucleases such as MRE11. Therefore, 53BP1

contributes to the resolution of stalled or regressed forks (Dungrawala et al., 2015;Her et al., 2018;[START_REF] Liu | Two replication fork remodeling pathways generate nuclease substrates for distinct fork protection factors[END_REF].

To detect and characterise the interaction between a protein and RNA, the laboratory used the cross-linking immunoprecipitation (CLIP) and the complex capture (2C)

techniques (Asencio et al., 2018;[START_REF] Ule | CLIP: A method for identifying protein-RNA interaction sites in living cells[END_REF]. These methods are based on UV-C crosslink which induces a covalent bond between RBPs and their associated RNAs. In CLIP, the protein is immunoprecipitated and then the bound RNAs are radioactively labelled, whereas in 2C the RNAs, and thus the bound proteins, are retained on a silica column followed by a western blot to analyse the RBPs. These two approaches have enabled the laboratory to previously identify the direct RNA-binding activity of 53BP1

(endogenous and transfected) in both cancerous and epithelial cells. Furthermore in vitro data have demonstrated that the GAR-Tudor region of 53BP1 possesses this RNA-binding ability (Leriche, Bonnet et al., in revision). By identifying the nature of this RNA, after extracting the nucleic acids bound to 53BP1 following the CLIP procedure, the laboratory found that 53BP1 binds to a specific substrate composed of The prior identification by the laboratory of the interaction between 53BP1 and RNA serves as the foundation for this thesis project, which seeks to better characterise this interaction and its implications in DNA replication. Three objectives are thus defined (Fig. 19):

1-Determine whether the recruitment of 53BP1 to the replication fork is dependent on the RNA primer present in the Okazaki fragments.

2-Understand the role of this RNA-binding in the normal mechanism of DNA replication, without induction of replication stress.

3-Characterise how the interaction between 53BP1 and the Okazaki fragments occurs, by identifying the protein domains implicated and an associated mutant.

RESULTS

RESULTS

I -Article: 53BP1 interacts with the RNA primer from

Okazaki fragments to support their processing during unperturbed DNA replication

The first objective of my thesis is to understand the role of the interaction between 53BP1 and RNA during DNA replication. To monitor this binding, in vitro assays were conducted. Subsequently transitioning to in cellulo model, we employed approaches based on the covalent bond between protein and RNA in living cells induced by UV-C: the cross-linking immunoprecipitation (CLIP) and complex capture (2C) techniques.

The choice of UV-induced crosslink offers the advantage of irreversible binding, enabling stringent wash conditions. In addition, CLIP and 2C are complementary methods. While the CLIP is protein-centric through immunoprecipitation, the 2C is RNA-centric using a silica column. Moreover, CLIP can be used to extract the nucleic acids bound to the protein and thus determine their nature (chimeras, hybrids, singlestrand, double-strand).

Once the interaction between 53BP1 and RNA-DNA primers, similar to the Okazaki fragments, had been identified, we used microscopy approaches to characterise the specific interactions to S-phase and at nascent DNA. Firstly, the quantitative imagebased cytometry (QIBC) technique was performed to determine the abundance of a specific protein in chromatin according to the cell cycle. To further elucidate the interaction at nascent DNA, the in situ protein interaction with nascent DNA replication forks (SIRF) technique was employed. This method is based upon proximity ligation assay (PLA) procedure and enables the visualisation of the association between 53BP1 and nascent DNA.

Through the application of all these techniques, we have demonstrated that 53BP1 interacts with the RNA primer of Okazaki fragments, with the purpose of facilitating their maturation during unstressed DNA replication.

Report

53BP1 interacts with the RNA primer from Okazaki fragments to support their processing during unperturbed DNA replication

INTRODUCTION

Recent observations highlight the importance of RNA-binding proteins (RBPs) in genome maintenance. RBPs, through their direct and generally specific binding to cis-acting elements in mRNAs, usually act as post-transcriptional regulators of gene expression. 1 Importantly, in addition to their proven role in DNA-damage-induced post-transcriptional regulation of DNA damage response (DDR) gene expression, RBPs, and the RNAs to which they bind, may play a more direct role in genome integrity. 2-4 Several proteins involved in the DDR have been suggested to be RBPs. However, a potential RNA-binding activity of these proteins remains to be evaluated. The p53-binding protein 1 (53BP1) is a key protein that mediates the signaling of DNA double-strand breaks (DSBs) in G1 and S/G2 phases of the cell cycle. In G1, 53BP1 acts as a molecular scaffold that recruits additional DSB-responsive proteins to damaged chromatin to limit DNA end resection and promote repair by non-homologous end joining (NHEJ). 5 The accumulation of 53BP1 at DSBs is RNA dependent, 6-8 and RNA immunoprecipitation (RIP) assays have shown that 53BP1 associates with RNAs. 7,8 These observations, however, do not demonstrate a direct interaction between 53BP1 and RNA, even if a recent proteomic analysis has identified 53BP1 as a candidate protein interacting with RNA. 9 In addition to its role in the DDR, 53BP1 plays an important function in DNA replication. Indeed 53BP1 has been shown to be rapidly recruited to stalled replication forks following a replicative stress induced by hydroxyurea (HU) treatment. 10 Lack of 53BP1 decreases cell survival and enhances chromosomal aberration after replication arrest. 11 53BP1 protects replication forks during the cellular response to replication stress in primary B cells, 12 and the S. cerevisiae 53BP1 ortholog (RAD9) protects stalled replication forks from degradation in Mec1 (ATR)-defective cells. 13 The mechanism of 53BP1 recruitment to replication forks may not be completely similar to the one existing for 53BP1 recruitment to DSBs since depletion of RNF8, RNF168, or 53BP1, but not MDC1, displays a similar replication defect (i.e., delayed fork progression and reversed fork accumulation). 14 It is therefore not clear which interaction(s) 53BP1 establishes to mediate its functions in DNA replication.

During initiation of DNA replication, several ribonucleotides are polymerized by the combined action of an RNA polymerase (DNA primase) and an error-prone DNA polymerase (POLa). DNA POLs are unable to initiate DNA synthesis de novo and require an initiating step to generate an RNA primer. This ''priming'' role is fulfilled by specialized DNA-dependent RNA POLs, called DNA primases containing a small catalytic subunit (PRIM1) and a large accessory subunit (PRIM2), capable of synthesizing short RNA chains (7-12 ribonucleotides). Primases are also essential throughout the process of lagging strand replication, where they initiate synthesis of the discontinuously synthesized Okazaki fragments. 15 On the lagging strand, the DNA POLd synthesizes approximately 100 deoxyribonucleotides downstream of this RNA-DNA primers to form the Okazaki fragments. 16 When POLd displaces the downstream fragment, it creates a short 5 0 flap single-stranded nucleic acid that is cleaved by the flap structure-specific endonuclease 1 (FEN1) to allow subsequent ligation of the two Okazaki fragments by DNA ligase I (LIG1). In some instances, a long flap is created that is covered with replication protein A (RPA), thereby blocking the access to FEN1. RPA can then recruit the DNA nuclease/helicase DNA2, which cleaves the long flap into a short flap that can be subsequently cleaved by FEN1. 17-20 Whether RBPs, reported to be present at replication forks, 21 directly bind RNA primers of Okazaki fragments during unperturbed DNA replication has not been established.

Using UV-C-induced crosslinking IP (CLIP 22 ) and complex capture (2C 23 ) experiments in living cells, we provide evidence that 53BP1 directly interacts with RNA. By analyzing the nature of the nucleic acids bound to 53BP1, we found entities of about 20-200 nt, composed of ribonucleotides at the 5 0 end followed by deoxyribonucleotides at the 3 0 end, that constitute Okazaki fragments. Consistent with the hypothesis that 53BP1 directly interacts with Okazaki fragments, we found that ribonuclease (RNase) treatment decreases 53BP1 recruitment to the fork (as determined by in situ analysis of protein interactions at DNA replication forks [SIRF] 24 ). In addition, while depletion of PRIM1 leads to reduced 53BP1 assembly, loss of FEN1 enhances the 53BP1 association with nascent DNA. Finally, depletion of 53BP1 leads to reduced RPA and DNA2 loading at nascent DNA. Altogether, these results indicate that the interaction between 53BP1 and Okazaki fragments is a key determinant of their processing and, more generally, of the functions of 53BP1 in DNA replication.

RESULTS

53BP1 is an RBP 53BP1 contains a putative intrinsically disordered GAR (glycinearginine-rich) RNA-binding domain (Figure S1A). 25 To investigate a potential RNA-binding activity, we first used an electrophoretic mobility shift assay (EMSA). The migration of the structured SC1 RNA (Figure S1B) (known to interact with the GAR domain of the FMRP protein; Figure S1C) was shifted by a recombinant GAR-Tudor 53BP1 protein (Figures S1D andS1E). Surface plasmon resonance (SPR) experiments indicated that the GAR-Tudor fragment interacts with the SC1 RNA with an affinity of about 8 nM, while a GAR-Tudor fragment mutated in the GAR domain (3R to 3K) completely lost its affinity with the SC1 RNA (Figure S1F). The ability of 53BP1 to directly bind to RNAs in vivo was analyzed following irradiation of living cells with high doses of UV-C (254 nm) in a CLIP experiment. 22 UV crosslinking requires direct contact (''zero'' distance) between protein and RNA and does not promote protein-protein crosslinking. 26,27 We irradiated hu-man HEK293T living cells with UV-C and immediately prepared cell lysates. Following 53BP1 IP and stringent washes, the IP was radiolabeled with T4 polynucleotide kinase (PNK), which incorporates 32 Pa tt h e5 0 end of nucleic acids (Figures 1Aa n d 1B). In UV-exposed cells, we observed a sharp radioactive band at the size of 53BP1 (Figure 1B), indicating that 53BP1 directly interacts with RNAs. A small interfering RNA (siRNA) pool directed against 53BP1 was used to ascertain that the band migrating at the size of 53BP1 corresponds to 53BP1 and that the lower band (labeled with an asterisk [*]) corresponds to a non-specific signal. We obtained very similar results in other cell lines, including HeLa cervical cancer cells, A2058 melanoma cells, RPE1 nontransformed human epithelial cells, and U2OS osteosarcoma cells (Figure S2A). To ascertain that the above CLIP results were not linked to an artifact related to the use of the 53BP1 antibody, we expressed GFP-53BP1 fusion proteins in HEK293T cells and performed the CLIP experiment with a GFP antibody instead of the 53BP1 antibody. A radioactive band was observed at the predicted size of the GFP-53BP1 protein or of the GFP-53BP1-Cter protein containing the carboxy-terminal half of 53BP1 that is still able to be recruited to ionizing radiation-induced damaged DNA (Figures S2B andS2C). Altogether, these data show that an RNA-binding activity can be found in the carboxy-terminal half of 53BP1 containing the GAR-Tudor domain.

Even if UV irradiation is mainly used to monitor RNA-protein interactions, 22 it has also been used to establish crosslinks between proteins and single-stranded DNA (ssDNA) (e.g., Steen et al. 28 ). We therefore used an orthogonal approach (2C) that uses the property of silica matrices to strongly and specifically retain nucleic acids and crosslinked nucleic acid-protein complexes based on charges (Figures 1A,1C,and 1D). 23 53BP1 was retained on the silica matrix in a UV-and RNA-dependent manner (Figure 1C). The well-characterized RBP HuR and the tubulin were used as positive and negative controls, respectively. Consistent with the EMSA data, the GAR-Tudor region is sufficient to interact with RNAs in living cells (Figure 1D). Altogether, these data show that 53BP1 directly interacts with RNA in vitro and in vivo.

53BP1 binds RNA-DNA chimeras

To analyze the nature of the nucleic acids bound to 53BP1, we extracted nucleic acids from the 32 P-labeled crosslinked nucleic acid-GFP-53BP1 complexes, loaded them on a denaturing TBE-urea gel, and revealed the radioactive signal by autoradiography. We found that nucleic acids bound to 53BP1 migrate as a smear of about 20-150 nt (Figure 2A). The smear collapsed below 25 nt following treatment with RNase A (which, in our experimental conditions, cleaves both ssRNA and double-stranded RNA, but not DNA or RNA engaged in an RNA:DNA duplex; Figure S3), showing that these nucleic acids possess ribonucleotides at their 5 0 ends. Strikingly, the size of the smear was also shortened following treatment with deoxyribonuclease I (DNase I) (which specifically cleaves DNA but not RNA; Figure S3), indicating that the nucleic acids interacting with 53BP1 are also composed of deoxyribonucleotides. The remaining DNase I-resistant shorter smear was composed of RNA at its 5 0 end since it disappeared following treatment with both RNase A and DNase I (Figure 2A). Similar results were found with the endogenous 53BP1 (Figure S4A). DNase I had no effect on nucleic acids bound to the RBP RBMX (Figure 2B), ruling out the possibility that DNase I was contaminated with RNases. We concluded that the cellular nucleic acids interacting with 53BP1 are RNA-DNA chimeras composed of 5 0 -end RNAs of different sizes (less than 25 nt) followed by deoxyribonucleotides (up to 100 nt) at their 3 0 ends. Of note, the GAR-Tudor fragment that is sufficient to bind RNA (Figures 1D andS1) cannot bind RNA-DNA chimeras since DNase I treatment did not lead (in contrast to RNase treat-ment) to a reduction in the size of the smear (Figure S4B), which is, therefore, only composed of ribonucleotides. This indicates that the binding of 53BP1 to Okazaki fragments through the GAR domain requires additional 53BP1 protein domains.

53BP1 interacts with Okazaki fragments

Based on these findings, we hypothesized that Okazaki fragments (the only known RNA-DNA chimeras in human cells) might To test this hypothesis, we depleted the human PRIM1 subunit that synthesizes RNA primers of replication and employed quantitative image-based cytometry (QIBC 29 ) to quantify the chromatin abundance of 53BP1 at the single-cell level. The depletion of PRIM1 with two independent siRNAs led to a decrease of 53BP1 association with chromatin (Figures 3A andS5A) in the different phases of the cell cycle (Figure S5B) without altering the cell-cycle profile (Figures S5C andS5D). Consistent with previous results, 30 PRIM1 loss led to DNA replication stress, as seen from the pronounced induction of ssDNA and chromatin loading of RPA2 during the S phase (Figures S5E andS5F). Interestingly, while PRIM1 depletion decreased the global chromatin association of 53BP1 (Figure S6A), it had a modest effect of disturbing spontaneously arising 53BP1 foci assembled on chromatin due to dedicated DDR signaling (Figure S6B). Together, these results suggest that in addition to the well-established histone-modification-based chromatin loading of 53BP1 at DSBs, 5 RNA primers synthesized by PRIM1 could facilitate the recruitment of 53BP1 on chromatin during DNA replication.

While the QIBC approach allows the determination of the chromatin abundance of proteins, it does not allow the determination of the abundance of proteins at replicating DNA. We therefore used a SIRF procedure that allows for the quantitative assessment of the proximity of proteins with ongoing, stalled, and previously active replication forks using a proximity ligation assay (PLA) approach. 24 As expected, 53BP1 could be found associated with nascent DNA (Figure S7). Treatment of cells with RNase A (Figure 3B) led to a decreased proximity between 53BP1, but not the proliferating cell nuclear antigen (PCNA), a component of the replisome, and nascent DNA, without changes in EdU incorporation (Figures S8A andS8B). The same result was observed upon depletion of PRIM1 (Figures 3Ca n dS8C). In contrast, depletion of FEN1, which leads to an accumulation of unligated Okazaki fragments, 20 led to an increased proximity between 53BP1 and nascent DNA (Figures 3Da n dS8D Okazaki fragments through depletion of PRIM1/FEN1 or RNA degradation leads to reduced proximity between 53BP1 and nascent DNA.

Depletion of 53BP1 perturbs the lagging strand processing Okazaki fragment maturation during lagging strand replication involves several layers of nuclease-driven pathways for proper RNA-DNA primer removal. Two pathways exist: the short flap pathway mediated by FEN1 and the long flap pathway mediated by DNA2/RPA. Based on our results that 53BP1 associates with Okazaki fragments during active DNA synthesis, we next sought to determine whether 53BP1 binding to RNA-DNA primers plays a role in supporting Okazaki fragment maturation. We found that 53BP1 depletion leads to an increased accumulation of mono-and poly-(ADP-ribose) (MAR and PAR) in chromatin (Figures 4A andS9) that is more pronounced in S and G2 (Figure S9B) and which constitutes a sensor of unligated Okazaki fragments. 31 Short incubation with PARG inhibitor to preserve nascent PAR further enhanced the levels of ADP ribosylation in 53BP1depleted cells (Figure 4A). We also found that depleting 53BP1 leads to reduced RPA and DNA2 loading at chromatin (Figures 4B,4C, 4D, and S10A-S10D), as well as accumulation of ssDNA (Figure S10E). Collectively, these results indicate that 53BP1 binds to long flaps of Okazaki fragments, fosters proper RNA-DNA primer removal, and prevents genotoxic DNA lesions that might arise from the lagging DNA strands in the absence of external stress inducer.

DISCUSSION

Here, we show that 53BP1 directly interacts with RNA in vitro and in vivo in human cells, revealing an additional activity of 53BP1 as a bona fide RBP (Figure 1). Strikingly, the nucleic acids interacting with 53BP1 are RNA-DNA chimeric structures composed of 5 0 -end ribonucleotides followed by deoxyribonucleotides at their 3 0 ends (Figure 2) that constitute Okazaki fragments (Figure 3). 53BP1, distinct from its canonical mode of chromatin binding, is anchored at the replication fork via its RNA-binding activity, thereby constituting a direct binder of Okazaki fragments during unperturbed DNA replication. Of note, it has been recently established that, during replication stress in fission yeast, the NHEJ factor Ku, also known as an RBP,4 binds to RNA:DNA hybrids originating from Okazaki fragments and establishes a barrier to replication fork degradation. 32 Similar to its role in DNA DSB repair pathway choice, 5 53BP1 may contribute to the choice between the short and long flap pathways during Okazaki fragment maturation. In yeast, it has been show that a delay in cleavage by FEN1 can induce a long flap. 18 Our data on the association of RPA and DNA2 with nascent DNA upon 53BP1 depletion (Figure 4) indicate that the interaction between 53BP1 and the long flap may contribute to prevent the action of FEN1, thus supporting the long flap pathway and further recruitment of RPA/DNA2. Maturation through the long flap pathway was shown to limit mutation rates, especially in transcribed regions, because of the replacement of the DNA primer synthesized by the error-prone POLa with a DNA strand synthesized by the error-free POLd. 33,34 In this scenario, 53BP1 may contribute to maintain genome integrity during DNA replication. This role of 53BP1 in the vicinity of replication fork might be of prime importance in human health. Indeed, defects in the maturation of Okazaki fragments lead to increased genetic instability, which causes certain cancers. 35-37 Moreover, several studies have indicated that TP53BP1 mutation might be associated with cancer risk. 38,39 Hence, the evaluation of 53BP1 mutations should be extended, beyond their effects on DSB signaling, to replication defects. In addition, it will be interesting to analyze the role of the RNA-binding activity of 53BP1 in the context of the sensitivity to PAR POL inhibitors (PARPis) of tumor cells, which are deficient in hereditary breast cancer (BRCA) genes (BRCA1 and BRCA2). PARPi, which has anticancer activity in BRCA-deficient cancers, perturbs the processing of Okazaki Cell Reports 42, 113412, November 28, 2023 5 Report fragments, a process that is modulated by 53BP1 loss. 40-42 The Okazaki fragment binding activity of 53BP1 may specifically be involved in the sensitivity of BRCA1-deficient cells to PARPi and may be linked to DNA replication gaps.

Limitations of the study

Our findings show that while the GAR-Tudor domain of 53BP1 is sufficient to bind RNA, it is not able to bind RNA-DNA chimeras in living cells. This indicates that binding of 53BP1 to RNA-DNA chimeras requires an additional RNA-binding domain or an additional 53BP1 domain that recruits the protein in the vicinity of Okazaki fragments. The GAR-Tudor domain alone could also adopt a conformation that affects its binding to RNA-DNA chimeras. Ideally, structural biology approaches will facilitate the defining of the minimal 53BP1 domain able to interact with RNA-DNA chimeras and an associated mutant. Moreover, even if our data indicate that 53BP1 interacts with Okazaki fragments on the lagging strand, the binding of 53BP1 to any RNA-DNA primers including the one present on the leading strand cannot be completely ruled out. Furthermore, our study does not provide visual insight with A CD B Quantitative image-based cytometry (QIBC) QIBC was performed as previously described. 44 Briefly, images were acquired with a ScanR inverted microscope high-content screening station (Olympus, IX81) equipped with wide-field optics, air objective, fast excitation and emission filter-wheel devices for DAPI, FITC, Cy3, and Cy5 wavelengths, an MT20 illumination system, and a digital monochrome Hamamatsu ORCA-Flash 4.0LT CCD camera. Images were acquired in an automated fashion with the ScanR acquisition software (Olympus, 3.2.1). 81-100 images were acquired containing at least 5,000 cells per condition. Acquisition times for the different channels were adjusted for non-saturated conditions in a 12-bit dynamic range, and identical settings were applied to all the samples within one experiment.

Images were processed and analyzed with ScanR analysis software. First, a dynamic background correction was applied to all images. The DAPI (Sigma-Aldrich) signal was then used to generate an intensity-threshold-based mask to identify individual nuclei as the main objects. This mask was then applied to analyze pixel intensities in different channels for each individual nucleus. For analysis of 53BP1 foci, additional masks were generated by segmentation of individual 53BP1 spots with spot-detector modules included in the software. Each focus was defined as a subobject, and this mask was used to quantify pixel intensities in foci. After this segmentation of objects and subobjects, the desired parameters for the different nuclei or foci were quantified with single parameters (mean and total intensities, area, foci count, and foci intensities). These values were then exported and analyzed with TIBCO Spotfire Software, version 11.1, to quantify absolute, median, and average values in cell populations and to generate all color-coded scatterplots. Within one experiment, similar cell numbers were compared for the different conditions (at least 4,000-5,000 cells), and for visualization, low x axis jittering was applied (random displacement of objects along the x axis) to make overlapping markers visible.

In situ protein interaction with nascent DNA replication forks (SIRF) U2OS were seeded on Millicell EZ SLIDE 8-well glass (Millipore, #PEZGS0816) 48h before experiment and then incubated with 25 mM EdU for 10 min. After treatment, cells were washed with cold PBS, pre-extracted with cold CSK (10 mM PIPES, 100 mM NaCl, 300 mM sucrose, 3 mM MgCl 2 , protease inhibitor cocktail), permeabilized with cold CSK-T buffer (CSK buffer supplemented with 2% Triton X-100) during 5 min at 4 C, treated or not with 1 mg/mL of RNAse A for 5 min at 37 C and then fixed with 4% PFA in PBS for 20 min at room temperature. After fixation, slides were placed in a humid chamber, and incubated with a click-it reaction (10 mM biotin azide, 100 mM sodium ascorbate, 2 mM CuSO 4 ,1mM Alexa Fluor 488 Azide, in PBS in this order) at room temperature for 1 h. After the click reaction, slides were washed with PBS and blocked with blocking buffer (10% goat serum and 0.1% Triton X-100 in PBS) for 1 h at room temperature. Primary antibodies (anti-53BP1, anti-biotin, anti-PCNA) were diluted in blocking buffer and incubated at 4 C overnight. Then a PLA procedure is performed using Duolink In situ Detection Reagents FarRed. Slides were imaged using Leica 3D upright deconvolution microscope with a CoolSNAP HQ camera, at the PICT-IBiSA Imaging Facility in Orsay, and analyzed using a semi-automatic macro on Fiji software.

Cloning GFP-RBMX and GFP-53BP1 plasmids correspond to RBMX or 53BP1 sequence inserted in pcDNA 6.2 (Vivid Colors pcDNA6.2/N-EmGFP-GW/TOPO Mammalian Expression Vector, Thermofisher). The GFP-53BP1-Cter corresponds to the 1235 to 1972 encoding sequence of 53BP1. The Hs53BP1-1288-1659 (HA-NLSx3-GAR-Tudor) encoding sequence was cloned by site-directed recombination using the Gateway technology (Life Technologies). The ORF was amplified by PCR using specific primers: G1_forward; Hs53BP1_F4; Hs53BP1_R4. The resulting PCR products were recombined during BP reaction with pDONO207 to generate Entry clones used for further LR reactions with pGGWA. 45 The triple mutation (Arg1396Lys, Arg1398Lys, Arg1401Lys) was generated by ''rolling-circle'' PCR using specific primers as previously described 46 : Hs53BP1-MutGAR_F and Hs53BP1-MutGAR_R.

53BP1 fragments purification

GST-Hs53BP1-1288-1659 fragments (GAR-Tudor or GAR(3K)-Tudor) purifications were performed as described. 47 Fractions eluted from Superdex 75 10/300 GL (Cytiva) were directly subjected to TEV cleavage at the ratio of 1 TEV for 10 GST-Hs53BP1 fragment (w/w). After 2 h incubation at 20 C, the mixture was incubated with Glutathione Sepharose 4B (Cytiva) and Ni-NTA Superflow (Qiagen). The Hs53BP1 fragment without its GST tag was recovered in the flow through and concentrated using Vivaspin concentrator (Sartorius).

Electrophoretic mobility shift assay (EMSA)

Appropriate concentrations of each protein and trace amount (25 nM) of ATTO700-labeled oligonucleotides were incubated with binding buffer (10 mM Tris-HCl, 1 mM EDTA, 100 mM KCl, 0.1 mM DTT, 5% glycerol, 0.01 mg/mL BSA, pH 7.5) in a 20 mL final reaction volume at room temperature for 45 min. The incubated samples were loaded on an 8% polyacrylamide (37.5:1 acrylamide/bisacrylamide) non-denaturing gel. Gels were imaged using an Odyssay scanner.

Surface plasmon resonance (SPR)

The interaction between nucleic acids (ligands) and the proteins (analytes) were measured by the surface plasmon resonance (SPR) technique using a ProteOn XPR36TM (Bio-Rad Laboratories). NLC Sensor chips precoated with NeutrAvidin (Bio-Rad Laboratories) were conditioned with 30-mL injections of 50 mM NaOH and 1.0 M NaCl at a flow rate of 30 mL/min in both ligand and analyte flow directions. Immobilization was performed in the running buffer (10 mM Tris-HCl, 200 mM KCl at pH 7.5, Tris-HCl, 0.01% v/v Tween).

The temperature of the chip surface was maintained at 25 C. The all biotinylated oligonucleotides were immobilized one by one onto a single channel of the six channel NLC sensor chip. Each oligonucleotide construct was immobilized onto an individual ligand channel by flowing the constructs at a concentration of 10-40 nM for 60 to 240 s injection. Binding to the chip surface resulted in approximately 60-100 response units (RU). The chip was rotated to the analyte flow direction and stabilized by flowing 60 mL analyte EMSA buffer three or more times, as required. Analyte (purified protein or buffer) was applied at various concentrations through the analyte channels at 100 mL/min for an association phase of 150 s, followed by a dissociation phase (running buffer only) of 600 s. The surface was regenerated with 18 mL of 0.5% SDS, at a flow rate of 30 mL/min followed by flowing 60 mL running buffer in the analyte direction at 100 mL/min. The resultant sensorgrams for each analyte interaction were analyzed with ProteOn software (Bio-Rad Laboratories). Kinetic parameters for first the dissociation constant (kd) and then the association constant (ka) were estimated by fitting the kinetic data according to the 1:1 Langmuir model and values of KD were then calculated (using the ratio kd/ka).

DNA damage induction before IF Micro-irradiation: the confocal microscope TCS SP5 Leica allowed DNA damage induction in cultured living U2OS cells using laser at 405 nm.

QIBC of native BrdU

To detect ssDNA in parental strands, the cells were labeled with 10 mM BrdU for 24 h and then released into a fresh growth medium for 1h prior to fixing cells. Cells were fixed, permeabilized, and processed for immunofluorescence staining as described in the IF and QIBC protocol. To detect BrdU in the native state, fixed cells were incubated with anti-BrdU antibody for 90 min followed by Alexa 594 secondary antibody at room temperature. To assess bulk BrdU incorporation levels as well as expose further BrdU epitopes, fixed cells were treated with DNase prior to immunostaining (data not shown degradation by nucleases such as MRE11 (Dungrawala et al., 2015;Her et al., 2018;[START_REF] Liu | Two replication fork remodeling pathways generate nuclease substrates for distinct fork protection factors[END_REF]. CLIP and 2C experiments performed in part I (Leriche, Bonnet et al., in revision) were carried out without induced stress, suggesting a new role for 53BP1 during unperturbed DNA replication, in addition to its known one at stalled replication forks.

We first aimed to determine the nature of the RNA that 53BP1 was binding to. We demonstrated that 53BP1 binds to RNA-DNA primers sensitive to RNAse A treatment, indicating that the RNA was single-strand. We next wanted to ascertain whether 53BP1 could bind RNA:DNA hybrids, present at the beginning of the elongation of Okazaki fragments. Of note those RNA:DNA hybrids can also be found at sites of transcription-replication conflicts and form R-loops. We used RNAse H treatment, which cleaves RNA in RNA:DNA hybrids, and we observed that the nucleic acids bound to 53BP1 is also sensitive to RNAse H, indicating that 53BP1 binds RNA:DNA hybrids (Fig. 1A). In vitro assays further revealed that the GAR-Tudor region of 53BP1 had the ability to bind the RNAse H-substrate RNA:DNA hybrids and that the 3K mutation of the GAR domain impaired the binding (Fig. 1B). Next, we wanted to ascertain whether the recruitment of 53BP1 to the fork was dependent on RNA. For this purpose, we used the SIRF technique that allows studying protein-protein interactions, close to less than 40nm, at nascent DNA (Roy et al., 2018) (Fig. 1C). The thymidine analogue

EdU is incorporated into nascent DNA in U2OS cells and then bound by biotin using a click-it reaction. A proximity ligation assay (PLA) is performed to analyse the association between 53BP1 and nascent DNA. The recruitment of 53BP1 to the fork decreased after treatment with RNAse A (Leriche, Bonnet et al., in revision) and also with RNAse H, without impacting the replisome factor PCNA (Fig. 1D).

Next, our findings show that while the GAR-Tudor domain of 53BP1 is sufficient to bind RNA, it is not able to bind RNA-DNA primers in living cells (Leriche, Bonnet et al., in revision). This contrasts with our in vitro data, where we found that the GAR-Tudor domain alone was sufficient to bind RNA-DNA primers (Fig. 1C). This indicates that the GAR-Tudor domain alone could adopt a structural conformation that affects its binding to RNA-DNA primers, since Tudor domain is known to interact with methylated Arginines [START_REF] Charier | The tudor tandem of 53BP1: A new structural motif involved in DNA and RG-rich peptide binding[END_REF][START_REF] Côté | Tudor domains bind symmetrical dimethylated arginines[END_REF]. Another hypothesis is that the binding of 53BP1 to RNA-DNA primer could require an additional domain that recruits the protein near the Okazaki fragments. Indeed, using the CLIP technique, I found that the C-terminal region of 53BP1 is the minimal binding region for Okazaki fragments, suggesting that other domains are involved in the interaction with RNA-DNA primers and/or in 53BP1 recruitment to the fork (Fig. 5A).

I then deleted the GAR domain and Tudor domain on the 53BP1 protein and performed CLIP experiments to visualize the nucleic acids bound to these constructs.

The same profile was obtained for the full length and the GAR-Tudor deleted versions, indicating that the GAR-Tudor region does not appear to be essential for binding to

Okazaki fragments (Fig. 5B).

DISCUSSION AND PERSPECTIVES

During my thesis, we characterised 53BP1 as a novel RNA-binding protein that interacts with a specific substrate composed of a short 5' RNA fragment directly followed by a longer 3' DNA fragment. Taken with additional experiments to study the association of proteins with nascent DNA during the S-phase, we concluded that 53BP1 directly interacts with the RNA primer of Okazaki fragments, where the RNA hybridises with the template DNA at the beginning of elongation, or as a flap structure during maturation of the lagging strand. However, the residues/domains of 53BP1 involved in this RNA-binding activity are not yet clearly defined. We particularly focused on the GAR domain, a known RNA-binding domain. The deletion or mutation of the GAR domain does not impact various function of 53BP1 such as foci formation, class switch recombination and anti-resection activity [START_REF] Lottersberger | Role of 53BP1 oligomerization in regulating double-strand break repair[END_REF]Pryde et al., 2005), while it appears to affect the replication functions of 53BP1, suggesting a dissociation of 53BP1 roles mediated by its domains. We thus assume that the GAR domain is important for 53BP1 to bind RNA-DNA primers at the replication fork, however our in vitro and in cellulo data show that the GAR domain is likely not the only player and other regions are directly or indirectly involved in the RNA-binding activity. Interestingly, a region located in the N-terminal part of 53BP1 may interact with RNA [START_REF] Queiroz | Comprehensive identification of RNAprotein interactions in any organism using orthogonal organic phase separation (OOPS)[END_REF] thus the impact of mutations in the Nterminal part on the RNA-binding activity could also be explored. These observations demonstrate the structural complexity of 53BP1 with domains possessing different functions that may or may not be synergistic. Additional validations are required, and a more in-depth structural biology study would help to define the RNA-binding residues and therefore to identify a mutant of 53BP1 that loses the interaction with RNA.

The factors involved in the signalling and regulation of this interaction remain unknown. We found that 53BP1 is recruited to the fork in a PRMTs-dependent manner. It is therefore possible that methylation of the GAR domain by PRMTs is involved in the signalling. Additionally, a negative regulator of 53BP1 is the RNAbinding protein TIRR. TIRR interacts with the Tudor domains of 53BP1 to prevent its recruitment to chromatin and also to regulate the 53BP1-p53 interaction. It would be interesting to determine whether TIRR also regulates the 53BP1-RNA interaction, for example by masking RNA-binding residues near the Tudor domains.

By identifying this 53BP1 interaction, we have also discovered a new role in DNA replication. Under normal conditions, 53BP1 interacts with the RNA-DNA primers of Okazaki fragments throughout their synthesis to assist the maturation. Okazaki fragments are mainly processed by the short flap pathway through the action of FEN1-LIG1. However in some case, when FEN1-clevage is delayed, Okazaki fragments are matured through the long flap pathway mediated by RPA/DNA2. We found that 53BP1 acts upstream of RPA/DNA2. In addition, a genetic study in 2011 proposed that Rad9, the 53BP1 homolog in s. cerevisiae, was recruited to long flaps in the absence of DNA2, associating with histone marks to induce cell cycle arrest [START_REF] Budd | Inviability of a DNA2 deletion mutant is due to the DNA damage checkpoint[END_REF].

Our data show that 53BP1 binds long flaps even in the presence of DNA2 and appears to promote the long flap pathway, consistent with the hypothesis that 53BP1 only binds a subset of Okazaki fragments. Through this role, 53BP1 could help to correct possible errors made during DNA replication by the error-prone Polα that produces the DNA primer following the RNA primer generated by the primases, and thus maintain genetic stability. Indeed, maturation through the long flap pathway would limit mutations due to transcriptional activity (Balakrishnan and Bambara, 2011;Balakrishnan et al., 2010). Therefore, it would be interesting to measure the mutation rate in the presence and absence of WT/mutated 53BP1. Additionally, sequencing the Okazaki fragments bound to 53BP1 could provide valuable information about the genomic localisation of this interaction, and thus answer to the question of whether 53BP1 binds to flap predominantly in transcriptionally active regions. Moreover, since we have observed that 53BP1 binds RNA:DNA hybrids at the fork, we could suppose that 53BP1 is also involved in the resolution of RNA:DNA hybrids due to transcriptionreplication conflicts. Indeed, 53BP1 could be recruited to active chromatin and thus be available to play two distinct roles: inducing the long flap on the lagging strand and preventing the formation of R-loops.

Furthermore, sequencing data would also help to better understand the 53BP1-RNA interaction, especially to determine whether this association is mediated by a specific RNA sequence, since the GAR domain is known to recognise G-rich regions. Several sequencing techniques for Okazaki fragments are possible (Ok-seq [START_REF] Petryk | Replication landscape of the human genome[END_REF]; Gloe-seq [START_REF] Sriramachandran | Genome-wide Nucleotide-Resolution Mapping of DNA Replication Patterns, Single-Strand Breaks, and Lesions by GLOE-Seq[END_REF]; TrAEL-seq [START_REF] Kara | Genome-wide analysis of DNA replication and DNA double-strand breaks using TrAEL-seq[END_REF]), but they are mostly developed in yeast models or they require a large quantity of cells making them challenging to perform. Attempts from the lab to perform Ok-seq on nucleic acids covalently bound to 53BP1 (CLIP) were so far unsuccessful.

It is important to highlight that these discoveries were made in unstressed conditions, a context in which 53BP1 is less studied. However, it would be interesting to extend our findings to stressed contexts where 53BP1 is active and thereby determine whether the RNA-binding regulates other functions of 53BP1. As the function of 53BP1 in DNA repair in G1-phase is mediated by RNA, it would be insightful to define whether this mediation is direct or indirect. To assess this, CLIP could be performed on G1-phase cells to elucidate whether 53BP1 exhibit an RNA-binding activity throughout the cell cycle or whether it is specific to the S-phase.

Moreover, the study of 53BP1 in BRCA1-deficient contexts could provide a better understanding of the sensitivity/resistance of certain cancers to clinically used drugs, such as PARP inhibitors. In the context of BRCA1 deficiency, there is a defect in HR and also a defect in processing the lagging strand, which promotes the development of cancers. These BRCA1-deficient cancers are sensitive to DNA-damaging agents, such as PARP1 inhibitors. In such scenarios, the loss of 53BP1 results in resistance to these drugs, and additionally rescues the defect in maturation of Okazaki fragments [START_REF] Chaudhuri | Replication fork stability confers chemoresistance in BRCAdeficient cells[END_REF]Cong et al., 2021;[START_REF] Paes Dias | Loss of nuclear DNA ligase III reverts PARP inhibitor resistance in BRCA1/53BP1 double-deficient cells by exposing ssDNA gaps[END_REF][START_REF] Panzarino | Replication gaps underlie BRCA deficiency and therapy response[END_REF]. Furthermore, it has been shown that aberrant expression or loss of function of 53BP1 contributes to the onset and development of tumors [START_REF] Mirza-Aghazadeh-Attari | 53BP1: A key player of DNA damage response with critical functions in cancer[END_REF]Rafnar et al., 2011;[START_REF] Timofeeva | Influence of common genetic variation on lung cancer risk: Meta-analysis of 14 900 cases and 29 485 controls[END_REF]. By using published databases on 53BP1 mutations, it would therefore be interesting to establish a potential correlation between its RNA-binding activity and predisposition to certain pathologies. Indeed, we can assume that this correlation between 53BP1 defects and cancer risk is not only associated with the DNA repair function of 53BP1 but also with its function in DNA replication, through its RNA interaction. Understanding the role of this binding could therefore be crucial in understanding the mechanisms of tumour progression and potentially lead to better therapeutic treatments, particularly in the context of cancers resistant to PARP inhibitors.

  sur l'ARN, tel que le « complex capture » (2C) et des procédures centrées sur les protéines, tel que le « cross-linking immunoprecipitation » (CLIP). Dans le cas du CLIP et du 2C, comme pour de nombreuses techniques d'identification des RBP, la première étape est une étape de liaison covalente entre les protéines et les ARN pour figer les interactions dans les cellules vivantes. Pour induire ce crosslink, l'exposition aux UV-C est souvent utilisée. Les UV-C nécessitent un contact direct entre la protéine et les acides nucléiques et ne crosslinkent pas les interactions protéine-protéine. Couplés à ces méthodes expérimentales, de nombreux outils in silico permettent également la prédiction des sites d'interactions sur l'ARN ou sur les RBP. Ces techniques computationnelles ne cessent d'émerger et récemment, AlphaFold et RoseTTAFold ont révolutionné le domaine de la biologie structurale.

  Le domaine GAR est une répétition de motif Glycines et Arginines, dont les arginines sont méthylées par les enzymes PRMT1 et PRMT5, qui est connu pour lier l'ARN et est retrouvé dans des RBP canoniques. Cette découverte soulève alors la question : est-ce que 53BP1 a une capacité de liaison à l'ARN et est-ce que cette protéine peut être caractérisée comme une RBP ? Plusieurs études se sont intéressées à cette potentielle activité et ont démontré que les fonctions de 53BP1 dans les dommages de l'ADN sont médiées par une composante ARN. En effet, les traitements RNAse A, qui clive l'ARN simple-brin, dans les cellules empêchent le recrutement de 53BP1 aux sites de cassures. Cependant, ces observations ne démontrent pas d'une interaction directe entre 53BP1 et l'ARN. De plus, l'inhibition de Dicer et Drosha, responsables de la production de petits ARN double-brin, diminue également le recrutement de 53BP1, indiquant alors que de l'ARN est produit au niveau des sites de lésion et est clivé par Dicer/Drosha. Quelques années après ces découvertes, une équipe a démontré qu'au niveau des cassures, l'ARN polymérase II est recrutée et produit de longs ARN non codants, de part et d'autre de la cassure, qui seront par la suite clivés par Dicer/Drosha. Cependant, le mécanisme de passage entre le long ARN non codant et un ARN double-brin reconnu par Dicer/Drosha n'est pas encore bien compris. La transcription d'ARN par l'ARN polymérase II permet également d'activer 53BP1 en dissociant le complexe 53BP1-TIRR. TIRR est une RBP qui régule négativement 53BP1 en interagissant avec ses domaines Tudor, masquant ainsi la surface de liaison aux histones et empêchant alors le recrutement de 53BP1 à la chromatine. Néanmoins, une liaison directe entre 53BP1 et l'ARN n'est pas encore démontrée et par ailleurs, le rôle du domaine GAR n'est pas clairement défini. La délétion ou la mutation de ce domaine n'affectent pas les fonctions de 53BP1, dans la signalisation du DDR ou dans la recombinaison de classe. Cependant, l'altération du domaine GAR n'a pas été étudiée dans un contexte de réplication de l'ADN, donc un potentiel rôle de ce domaine dans ces fonctions n'est pas à exclure. Enfin, même si 53BP1 n'a pas encore été caractérisée comme étant une RBP, deux études protéomiques récentes l'ont suggérée comme un réel candidat. OBJECTIFS DE LA THESE Le laboratoire s'est donc intéressé à la caractérisation d'une interaction directe entre 53BP1 et l'ARN. Pour détecter une activité de liaison à l'ARN, des techniques in cellulo de CLIP et 2C, basées sur un lien covalent entre protéine et ARN induit par les UV-C, sont utilisées. Ces techniques sont complémentaires puisque dans le CLIP, la protéine est immunoprécipitée puis les ARN liés sont marqués radioactivement à l'extrémité 5' alors que dans le 2C, les ARN sont retenus sur une colonne de silice suivie d'une analyse en western blot pour analyser les RBP liées aux ARN. Ces deux approches ont permis de détecter une activité de liaison à l'ARN de 53BP1 (endogène et transfectée) dans plusieurs lignées cellulaires, cancéreuses et épithéliales. Par ailleurs, des données in vitro ont montré que le fragment GAR-Tudor de 53BP1 possédait cette capacité de liaison à l'ARN. Par la suite, pour déterminer la nature de l'ARN lié par 53BP1, la membrane, contenant les complexes ARN-protéines après la procédure de CLIP, est découpée pour extraire les acides nucléiques, qui sont alors déposés sur un gel dénaturant. Il a été observé que 53BP1 interagit directement avec des chimères ARN-ADN, avec une courte partie ARN en 5' et une longue partie ADN en 3', caractéristiques des amorces ARN-ADN générées par PRIM1-Polα au niveau des fragments d'Okazaki. L'objectif de cette thèse est donc de déterminer si le recrutement de 53BP1 à la fourche dépend de l'amorce ARN des fragments d'Okazaki, de comprendre le rôle de cette interaction dans le mécanisme de réplication de l'ADN et enfin, de mieux caractériser l'interaction entre 53BP1 et les fragments d'Okazaki en définissant les domaines protéiques impliquées et en identifiant un mutant associé. RÉSULTATS Nous avons démontré par des expériences de SIRF, permettant l'étude des interactions entre protéines et ADN naissant, que le recrutement de 53BP1 est inhibé si les cellules sont traitées préalablement avec la RNAse A, qui clive l'ARN simple-brin, et la RNAse H, qui clive l'ARN contenu dans des hybrides ARN:ADN. Par ailleurs, la déplétion de PRIM1, qui génère l'amorce ARN des fragments d'Okazaki, induit une diminution significative de 53BP1 à la fourche. À l'inverse la perte de FEN1, qui clive l'amorce ARN lors de la maturation des fragments d'Okazaki et accumule des flaps non-clivés, augmente l'association de 53BP1 à l'ADN naissant. Ces données indiquent donc que 53BP1 interagit avec l'amorce ARN des fragments d'Okazaki, sous la forme hybridée à l'ADN, au début de l'élongation, ainsi que sous la forme simple-brin, pendant la maturation, et que cette interaction est essentielle pour le recrutement de 53BP1 à la fourche. Nous avons alors supposé que le rôle de 53BP1 était d'aider la maturation des fragments d'Okazaki. Si tel est le cas, la déplétion de 53BP1 devrait induire un défaut dans le traitement du brin retard. Ces défauts sont visibles par la présence de chaînes PAR car ils résultent en la formation d'un espace d'ADN simple-brin, entre deux fragments d'Okazaki non ligaturés, qui est reconnu par PARP conduisant alors à une cascade de PARylation. En utilisant une approche quantitative de cytométrie, le QIBC, permettant de déterminer l'abondance d'une protéine à la chromatine selon la phase du cycle cellulaire, nous avons observé que la déplétion de 53BP1 accumulait les chaînes PAR en phase S, traduisant ainsi par un défaut de maturation du brin retard. De plus, la perte de 53BP1 diminue l'association à l'ADN naissant de RPA et DNA2, qui sont les acteurs de la voie du long flap. Cela indique donc que 53BP1 agit en amont de ces protéines et pourrait promouvoir la voie du long flap pour maturer les fragments d'Okazaki. Ces résultats ayant été obtenus dans un contexte non stressé de la fourche, il serait intéressant de déterminer si le rôle connu de 53BP1 dans la gestion du stress réplicatif est aussi médié par l'amorce ARN. Nous avons ensuite tenté d'identifier un mutant de 53BP1 qui perd la liaison à l'ARN et donc potentiellement qui altère ses fonctions dans la réplication de l'ADN. En première intention, nous nous sommes concentrés sur le domaine GAR qui un domaine putatif de liaison à l'ARN. Ce domaine contient une répétition d'arginines et de glycines dont les arginines sont méthylées par PRMT1 et 5. Nous avons observé que le recrutement de 53BP1, à la fourche de réplication, est dépendant de cette signalisation de méthylation médiée par les enzymes PRMT. En outre, en utilisant la technique de CLIP et 2C, nous avons démontré que la région GAR-Tudor de 53BP1 n'est pas suffisante pour lier les amorces ARN-ADN, et que la région C-terminal est la région minimale de liaison aux fragments d'Okazaki. Ces données in cellulo sont en contradiction avec les données in vitro où le fragment GAR-Tudor suffisait pour lier des amorces ARN-ADN. Cela indique que cette région seule peut adopter une structure in cellulo qui affecte la liaison aux fragments d'Okazaki, ou que d'autres domaines protéiques de 53BP1 sont nécessaires pour son recrutement à la fourche de réplication. La délétion de la région GAR-Tudor sur la protéine 53BP1 ne semble pas affecter la liaison aux fragments d'Okazaki par CLIP. Cependant, en réalisant des expériences de QIBC, la délétion du domaine GAR seul semble suffire à diminuer l'association de 53BP1 à la chromatine. L'implication du domaine GAR dans l'interaction 53BP1fragments d'Okazaki n'est donc pas encore claire et une étude de biologie structurale plus approfondie aiderait à la caractérisation de l'interaction. DISCUSSION ET PERSPECTIVES Ces données ont donc permis de définir un nouveau rôle de 53BP1 et de l'identifier comme une RBP. Similaire à son rôle connu dans le choix des voies de réparation, 53BP1 pourrait être impliquée dans le choix des voies de maturation des fragments d'Okazaki. La voie que 53BP1 semble favoriser est la voie du long flap, qui paraît être une voie moins mutagène et plutôt choisie dans des zones transcriptionnellement actives. Par ailleurs, 53BP1 lie des hybrides ARN:ADN qui pourraient être la conséquence des conflits réplication-transcription présents au niveau de la chromatine active (R-loop). 53BP1 serait alors recrutée dans ces régions et ainsi jouer deux rôles distincts : induire un long flap sur le brin retard et prévenir de la formation des R-loop. Il serait alors intéressant de séquencer les fragments d'Okazaki liés à 53BP1 pour définir la localisation de cette interaction. De plus, cette nouvelle liaison que 53BP1 établie, dans la réplication de l'ADN, pourrait être étudiée dans un contexte cancéreux puisque la délétion de 53BP1 est associée à des pathologies humaines ou à des résistances thérapeutiques. En effet, dans des cancers déficients pour BRCA, la perte de 53BP1 est connue pour induire une résistance aux inhibiteurs de PARP. Comprendre l'interaction 53BP1-ARN, dans ces situations, pourrait donc s'avérer important pour mieux appréhender les mécanismes de progression tumorale, déterminer des prédispositions pour certaines maladies et ainsi conduire à de meilleurs traitements thérapeutiques. Par ailleurs, l'interaction 53BP1-ARN est la première interaction décrite entre une RBP et les fragments d'Okazaki. Il serait donc intéressant de caractériser d'autres RBP liant ces amorces ARN dans la réplication de l'ADN, par des approches à grande échelle d'identification de protéines à la fourche, tel que l'iPOND, couplées à des approches d'identification de RBP, tel que l'OOPS. Cette étude permettrait potentiellement de découvrir de nouvelles interactions protéines-ARN impliquées dans la réplication de l'ADN.

1. 2 .

 2 End-joining processes ............................................................................................................ 1.3. Double-strand break repair choice .................................................................................. 1.4. Replication stress response.................................................................................................. 2. Structure ............................................................................................................................................... 3. 53BP1 and RNA .................................................................................................................................. OBJECTIVES ............................................................................................................................................. RESULTS ................................................................................................................................................... I -Article: 53BP1 interacts with the RNA primer from Okazaki fragments to support their processing during unperturbed DNA replication ....................................................................................... II -Additional results .......................................................................................................................................... 1. Investigation of the role of 53BP1-Okazaki fragments interaction ............................ 2. Characterisation of a 53BP1 mutant with altered function in replication .............. DISCUSSION AND PERSPECTIVES ............................................................................................. REFERENCES ........................................................................................................................................ ABSTRACT ............................................................................................................................................

  properties of the biotin ligase (BirA) from the bacterium Escherichia coli. The RNA of interest is tagged with an aptamer called boxB, enabling the recruitment of BirA which can biotinylate proteins in close proximity of the RNA of interest. The biotinylated proteins are then captured by streptavidin-coated beads and analysed by western blot or mass spectrometry. More recently, new techniques have emerged with the discovery of CRISPR systems. In dCas13-based approaches, such as the CRISPR-based RNA-united interacting system (CRUIS), a guide RNA recruits catalytically dead Cas13

  human cells and the fidelity of this process rely on three different polymerases: Polα, Polδ and Polε. There are notable differences in fidelity among these three polymerases. Unlike Polδ and Polε, Polα lacks 3'-exonuclease activity, and

  damage, ATM phosphorylates 53BP1, recruiting RIF1 which assists in the dissociation of the 53BP1-TIRR complex to release the inhibition of 53BP1 and enables 53BP1 to play its role in DNA damage response. This dissociation is made possible by RNAs transcribed at DSBs by RNA Pol II and is dependent on Dicer and Drosha[START_REF] Ketley | DNA double-strand break-derived RNA drives TIRR/53BP1 complex dissociation[END_REF]. Moreover, using in vitro EMSA assays to visualize the affinity between a fragment of 53BP1 containing only the Tudor region and various substrates (ssRNA, ssDNA and RNA beacon), Ketley et al. did not observe a binding of the Tudor domains to any of the tested substrates. Therefore, the Tudor domains do not seem to have an RNA-binding activity. But what about the GAR domain of 53BP1, the characterised RNA-binding domain? The role of the GAR domain of 53BP1 is still not well understood and a direct binding to RNA has not yet been demonstrated. In 2005 Pryde et al, interestingly demonstrated that the

a 5 '

 5 RNA part and a 3' DNA part, similar to Okazaki fragments. The Okazaki fragments are discontinuously produced on the lagging strand during S-phase and the RNA-DNA primers are essential for their synthesis. During the replication, ribonucleotides are produced by the action of the primase PRIM1 and, following this RNA primer, the DNA polymerase α can initiate DNA synthesis. The RNA primer (referred to as a flap in its single-strand form) is then processed and cleaved by the endonuclease FEN1, before the ligation of Okazaki fragments by the DNA ligase 1. Up to now, although certain RBPs are present at the replication fork, it has never been demonstrated that these RBPs directly interact with the RNA primers of the Okazaki fragments.

d

  53BP1 is an RNA-binding protein that directly interacts with RNA-DNA chimeric structures d Perturbation of Okazaki fragment processing changes the proximity of 53BP1 with nascent DNA d Depletion of 53BP1 perturbs the maturation of Okazaki fragments Authors Melissa Leriche, Clara Bonnet, Jagannath Jana, ..., Kumar Somyajit, Patricia Uguen, Ste ´phan Vagner Correspondence stephan.vagner@curie.fr In brief Leriche et al. show that 53BP1, distinct from its canonical mode of recruitment to chromatin, is an RNA-binding protein that interacts with RNA-DNA chimeric structures constituting Okazaki fragments. 53BP1 acts as a binder of Okazaki fragments and plays a role during unstressed DNA replication. Leriche et al., 2023, Cell Reports 42, 113412 November 28, 2023 ª 2023 The Authors. https://doi.org/10.1016/j.celrep.2023.113412

Figure 1 .

 1 Figure 1. 53BP1 directly interacts with RNA (A) Schematic representation of the crosslinking immunoprecipitation (CLIP) and complex capture (2C) procedures. WB, western blot; IP, immunoprecipitation; PNK, polynucleotide kinase. (B) Top: autoradiography of 53BP1-nucleic acid complexes in UV-C-treated (+) or -untreated (À) HEK293T cells depleted of 53BP1 (si53BP1) or left untreated (siControl [siCtrl]). Bottom: IP of 53BP1 and WB of 53BP1 and GAPDH (used as a normalization control); same conditions as on the top. The asterisk indicates a non-specific band (*), and the arrow indicates the position of 53BP1-nucleic acid complexes (n = 3 biological replicates). (C) WB on 2C experiments performed in A2058 cells (n = 3 biological replicates). (D) WB on 2C experiments performed in GAR-Tudor-transfected HEK293T cells (n = 3 biological replicates).

Figure 2 .

 2 Figure 2. 53BP1 binds an RNA-DNA chimera CLIP (left) and nucleic acid extraction (right) from GFP-53BP1-(A) or GFP-RBMX (B)-transfected HEK293T cells (n = 3 biological replicates).

Figure 3 .

 3 Figure 3. 53BP1 interacts with the RNA primer of Okazaki fragments (A) QIBC of chromatin-loaded 53BP1 mean intensity. U2OS cells were treated with indicated siRNAs for 48 h (also see Figure S6). The horizontal lines are median values. p values were determined by one-way ANOVA with Tukey's test, ****p < 0.0001 (n > 10,000 cells from 3 biological replicates). (B) Representative images and/or analysis of 53BP1-EdU and PCNA-EdU SIRF in U2OS cells permeabilized and treated with RNase A (RNase A) or left untreated (Mock); PLA signal (red), Alexa Fluor 488-EdU staining (green), and DAPI staining (blue). The significance for 53BP1-EdU and PCNA-EdU PLA values (shown as a scatterplot) was derived from the Mann-Whitney statistical test. Bars represent the mean ± SD. ****p < 0.0001 (n = 3 biological replicates). (C and D) Representative scatterplot of U2OS cells depleted of PRIM1 (siPRIM1) (C) or FEN1 (siFEN1) (D) or left untreated (siCtrl) for 48 h. The significance was derived from the Mann-Whitney statistical test. Bars represent the mean ± SD. ****p < 0.0001 (n = 3 biological replicates).

Figure 4 .

 4 Figure 4. Depletion of 53BP1 perturbs lagging strand processing (A) QIBC of chromatin-loaded protein mono-ADP-ribosylation (MARylation) or poly-ADP-ribosylation (PARylation). U2OS cells were treated with either control or 53BP1 siRNAs for 72 h and incubated with DMSO or PARGi (10 mM) for the last 30 min before pre-extraction and cell fixation. Nuclear DNA was counterstained by DAPI. The horizontal lines are medians. Right: quantification of MAR/PAR signals from the experiment on the left. In boxplots, center lines are medians, the boxes indicate the 25th and 75th centiles, and the whiskers indicate Tukey values. p values were determined by one-way ANOVA with Tukey's test. ****p < 0.0001 (n > 10,000 cells from 2 biological replicates). (B) QIBC of chromatin-loaded RPA2 mean intensity. U2OS cells were treated with either control or 53BP1 siRNAs for 72 h. The horizontal lines are medians (n > 5,000 cells from 2 biological replicates). (C and D) Representative scatterplot of SIRF experiments of RPA (C) or DNA2 (D) in U2OS cells depleted of 53BP1 (si53BP1) or left untreated (siCtrl) for 48 h. The significance was derived from the Mann-Whitney statistical test. Bars represent the mean ± SD. ****p < 0.0001 (n = 3 biological replicates).

Figure S2 .

 S2 Figure S2. Related to Figure 1. 53BP1 directly interacts with RNA (A) Autoradiography of 53BP1-nucleic acid complexes in UV-C treated (+) or untreated (-) cervix cancer cells (HeLa), melanoma cells (A2058), osteosarcoma cells (U2OS) or epithelial cells (RPE1) depleted of 53BP1 (si53BP1) or left untreated (siCtrl). The asterisk indicates a non-specific band (*), the arrow indicates the position of 53BP1-nucleic acid complexes. (B) CLIP experiments performed in HEK29T cells transfected with the plasmids expressing the GFP-53BP1 proteins or the C-terminal part of 53BP1 (1235 -1972 amino acids) or GFP alone. 10% of the IP was analysed by western blot to confirm immunoprecipitation (lower panel). (C) Immunofluorescence was performed in U2OS cells expressing the GFP-53BP1 or GFP-53BP1-Cter (1235 -1972 amino acids) proteins. The cells were irradiated with laser micro irradiation at 405 nm (stripes) and stained with anti-γH2AX (red) or GFP (green) antibodies.

2 .

 2 Figure S3. Related to Figure 2. Enzymatic activities of RNAse A and DNAse I In vitro assays of RNAse A and DNAse I activities on synthetic 20 nt-long oligonucleotides. The synthetic oligonucleotides were 32 P-5'end labelled on both strands.

Figure S4 . 3 .

 S43 Figure S4. Related to Figure 2. Endogenous 53BP1 binds RNA-DNA chimeras while the GAR-Tudor fragment does not (A) Nucleic acid extraction after CLIP procedure of endogenous 53BP1 in A2058 cells. (B) Nucleic acid extraction after CLIP procedure of HA-NLS x3 -GAR-Tudor fragment in transfected HEK293T cells.

TOP1 LIST OF FIGURES AND TABLES Figure 1 | Number of identified RNA-binding proteins in different organisms

  ....................

	RBMX SYF2	RNA-binding motif protein X-linked SYF2 pre-mRNA Splicing Factor
	RBP TERRA	RNA-binding protein long non-coding Telomeric Repeat-containing RNA
	RBPome TERT	RNA-Bound Proteome Telomerase Reverse Transcriptase
	RecQ TIRR	RecQ DNA helicase (RecQ1 ; RecQL4) Tudor-Interacting Repair Regulator
	RFC TLS Figure 2 |	Replication Factor C Translesion DNA Synthesis
	RGG	Arginine/Glycine-rich repeats
	RIC	RNA Interactome Capture
	RIF1	Guanine quadruplexes Replication Timing Regulatory Factor 1
	hnRNP RIP	heterogenous nuclear Ribonucleoprotein (hnRNP A1; C; F; G; H; U) RNA Immunoprecipitation
	HR R/K patch	Homologous Recombination Arginine-or Lysine-rich patch
	HTLF RNA	Helicase-like Transcription Factor Ribonucleic Acid
	HU RNA Pol II	Hydroxyurea RNA Polymerase II
	HuR RNAse	Human antigen R Ribonuclease
	IDR RNF	Intrinsically Disordered Region Ring Finger Protein (RNF8; RNF168)
	IdU RPA	5-iodo-2′-deoxyuridine Replication protein A
	IR RRM	Ionizing Radiation RNA Recognition Motif
	IRES rRNA	Internal Ribosome Entry Site ribosomal RNA
	KASRIC RT-qPCR	Kethoxal Assisted Single-strand RIC Reverse Transcription-quantitative PCR
	KH SAMHD1	Lysine Homology SAM domain-and HD domain-containing protein 1
	LIG s. Cerevisiae	DNA Ligase (LIG1; LIG3; LIG4) saccharomyces Cerevisiae
	LINP1 SDS-PAGE	lncRNA in NHEJ pathway 1 Sodium dodecyl sulfate polyacrylamide-gel electrophoresis
	lncRNA serIC	long non-coding RNA serial RIC
	MCM2-7 SFPQ	Minichromosome maintenance 2-7 Splicing Factor Proline-and Glutamine-rich
	MDM SIRF	Murine Double Minute (MDM2; MDMX or 4) in situ protein Interaction with nascent DNA Replication Forks
	METTL SMARCAL1	Methyltransferase-like Enzyme SWI/SNF-related matrix-associated actin-dependent regulator of
	MMR	Mismatch Repair chromatin subfamily A-like protein 1
	M-phase SMN	Mitosis phase Survival Motor Neuron protein
	MRE11 S-phase	Meiotic Recombination protein 11 Synthesis phase
	MRN SR	MRE11-RAD50-NBS1 Serine/Arginine repeats
	mRNA SRSF1	messenger RNA Serine/Arginine-rich Splicing Factor 1
	Ms2-BioTRAP SSB	MS2 in vivo Biotin-tagged RAP Single-strand break
	NBS1 ssDNA	Nijmegen Breakage Syndrome 1 single-strand DNA
	NER ssRNA	Nucleotide Excision Repair single-strand RNA
	STQ sites	Serine/Threonine/Glutamine sites

Canonical roles of RNA-binding proteins.............................................................................. Figure 3 | RNA-binding proteins help to maintain genome stability

  ..........................................

	Figure 4 |

Complex capture (2C) procedure

  ..............................................................................................

	Figure 5 |

Cross-linking immunoprecipitation sequencing (CLIP-seq) procedure

  ............ Origin licensing ................................................................................................................................... Origin firing ...........................................................................................................................................Figure 8 | DNA elongation ....................................................................................................................................

	Figure 6 | Figure 7 | Figure 9 |

Short and long flap pathways to mature Okazaki fragments

  ....................................

	Figure 10 |

Termination of the replication

  ................................................................................................

	Figure 11 |

Main sources of replication stress

  .........................................................................................

	Figure 12 |

Recovery of stressed replication forks

  ................................................................................Figure 13 | Fork reversal mechanism ...........................................................................................................

	Figure 14 |

Double-strand break repair mechanisms

  .........................................................................

	Figure 15 |

Roles of 53BP1 during the different phases of the cell cycle

  ..................................

	Figure 16 |

Double-strand break repair pathways

  ................................................................................

	Figure 17 |

Structure of the 53BP1 protein

  ...............................................................................................

	Figure 18 |

Production of RNA at DNA damage sites

  ............................................................................

	Figure 19 |

Objectives of the thesis

  .................................................................................................................

Table 1 | Examples of canonical and uncoventional RNA-binding domains

 1 ...........................

Table 2 | Examples of methods to monitor RNA-protein interactions

 2 .......................................

Table 3 | Major characteristics of eukaryotic DNA polymerases

 3 ..................................................

	INTRODUCTION

Table 1 |

 1 Examples

of canonical and unconventional RNA-binding domains. Adapted from

[START_REF] Corley | How RNA-Binding Proteins Interact with RNA: Molecules and Mechanisms[END_REF]

  target RNA are not always well understood, but various crystallography and structural biology techniques have provided some insights. Several methods are available to characterise RNA-protein interactions in vivo, both from a protein-centric or RNAcentric perspectives. In other words there are techniques to identify the RNAs bound to a specific RBP or to determine the RNA-bound proteome (RBPome), i.e. the entire set of proteins interacting with RNA at a given time[START_REF] Van Ende | Single and combined methods to specifically or bulk-purify RNA-protein complexes[END_REF][START_REF] Ramanathan | Methods to study RNA-protein interactions[END_REF] (Table

Table 2 |

 2 Examples of methods to monitor RNA-protein interactions. Adapted from[START_REF] Ramanathan | Methods to study RNA-protein interactions[END_REF][START_REF] Van Ende | Single and combined methods to specifically or bulk-purify RNA-protein complexes[END_REF]

	Methods	Advantages	Disadvantages	References

Table 3 | Major characteristics of eukaryotic DNA polymerases.

 3

		Polα	Polδ	Polε
	Activities	Polymerase	Polymerase 3'-exonuclease	Polymerase 3'-exonuclease

  ). Briefly, individual coverslips were incubated with 10 units of RNase-Free DNase (M6101, Promega) in 1X Reaction Buffer (400 mM Tris-HCl [pH 8.0 at 25 C], 100 mM MgSO 4 , 10 mM CaCl 2 ) for 40 min at room temperature. After that, coverslips were washed once with Stop Buffer (20 mM EGTA [pH 8.0 at 25 C]) in PBS and further washed two-three times with PBS. Coverslips were then equilibrated in Antibody diluent media for 15 min at room temperature before processing for immunofluorescence staining as described earlier. The predictive docked structure of SC1 RNA (grey in colour) with GAR-TUDOR RGG motif (PRGRGRRGRP, red in colour) superimposed with X-ray crystal structure of the FMRP RGG peptide (GDGRRRGGGGRGQG, green in colour)-SC1 RNA complex. The RGG motif of GAR-TUDOR and FMRP protein showed comparable binding mode with SC1 RNA in the duplex-

	Figure S1. 53BP1 binds RNA in vitro
	(A) 53BP1 domains map.
	(B) Structure of SC1 RNA.
	Report (C) quadruplex junction.
	(D) Gel purification of GAR-Tudor and GAR (3K)-Tudor recombinant proteins.
	(E) Electrophoretic Mobility Shift Assay (EMSA) performed using purified recombinant 53BP1
	containing the Tudor and the GAR domains incubated with a 5'-end fluorescently labelled synthetic
	36 nt-long SC1 RNA.
	(F) Surface plasmon resonance (SPR) experiments with the indicated recombinant proteins and a
	5'-end biotinylated SC1 RNA.
	QUANTIFICATION AND STATISTICAL ANALYSIS
	Statistical analyses were performed on GraphPad Prism (Version 8.0) and were carried out using either a one-way ANOVA with Tu-
	key's test or a Mann-Whitney U test, as indicated in the figure legends.

Cell

Reports 42, 113412, November 28, 2023 13

Cell Reports 42, 113412, November 28, 2023Report

Cell Reports 42, 113412, November 28, 2023Report

Cell Reports 42, 113412, November 28, 2023Report

Cell Reports 42, 113412, November 28, 2023Report

CellReports 42, 113412, November 28, 2023 9Report

Cell Reports 42, 113412, November 28, 2023Report

ACKNOWLEDGMENTS

ACKNOWLEDGMENTS

We thank Laetitia Besse from the Multimodal Imaging Center Imaging Facility of the Institut Curie (CNRS UAR2016/INSERM US43/Institut Curie/ Universite ´Paris-Saclay) for support with imaging and analyses. We thank the Institut Curie Next Generation Sequencing (Sylvain Baulande) platform for high-throughput sequencing. We also thank Aura Carreira and Martin Dutertre for helpful comments on the manuscript. This work was supported by grants from Institut Curie, Gustave Roussy, INSERM, CNRS, and Equipe labellise ´e Ligue Nationale Contre le Cancer (LNCC) (to S.V.); the Lundbeck Foundation Fellowship (R345-2020-1770) and the Danish Cancer Society (R325-A18913) (to K.S.); and the Association for Research against Cancer (Fondation ARC) (ARCPJA32020070002430), AT Europe Association, a Radiobiology program CEA grant, and INSERM house funding (SGCSR unit) (to P.B.). M.L. was supported by a pre-doctoral fellowship from Ligue Nationale Contre le Cancer (LNCC) and C.B. and S. Mennour by pre-doctoral fellowships from the Ministe `re de l'Enseignement Supe ´r i e u re td el aR echerche (MESR).

electron microscopy of the effect of 53BP1 depletion on Okazaki fragments.

STAR+METHODS

Detailed methods are provided in the online version of this paper and include the following:

AUTHOR CONTRIBUTIONS

M.L., C.B., and P.U. performed the CLIP, SIRF, and 2C experiments. J.J. performed the gel shift and SPR experiments. G.C. and K.S. performed the QIBC experiments. P.B., D.B., and X.V. performed recombinant protein production and associated molecular cloning. S. Mennour and S. Martineau performed all other molecular cloning and laser micro-irradiation experiments. S.V., P.U., K.S., M.L., and C.B. conceived the experiments and analyzed data. S.V. coordinated studies and wrote the manuscript with the help of P.U., C.B., K.S., and S.L. and with input from all other authors. S.V., K.S., and P.B. acquired funding.

DECLARATION OF INTERESTS

The authors declare no competing interests.

STAR+METHODS KEY RESOURCES

EXPERIMENTAL MODEL AND SUBJECT DETAILS

Cells were grown in 5% CO 2 humidified incubator at 37 C. U2OS, RPE1, HeLa and HEK293T cells were grown in DMEM supplemented with 10% FBS and 2 mM L-glutamine. A2058 cells were grown in MEM supplemented with 10% FBS and 2 mM L-glutamine.

METHOD DETAILS

siRNA and plasmid transfections siRNA and plasmid transfections were performed, respectively, using Lipofectamine RNAiMAX (Life Technologies, #13778150) or JetPEI (Polyplus Transfection, #101000053) according to manufacturer's instructions.

CrossLinking and immunoprecipitation (CLIP)

Cells were irradiated with UV-C light at 254nm, lysed in RIPA buffer (50 mM Tris-HCl pH 7.4, 100 mM NaCl, 1% Nonidet P-40, 0.1% SDS, 0.5% Sodium deoxycholate), supplemented with RNAse OUT (1mL/1mL, Invitrogen, #10777019) and protease inhibitor cocktail, and immunoprecipitated with 50 mL of Dynabead Protein G (Thermofisher, #10009D) using 5 mgo fr a b b i ta n t i -5 3 B P 1o r5mgo fm o u s e anti-GFP antibodies for each condition. Immunocomplexes were washed twice with RIPA-S buffer (50 mM Tris-HCl pH 7.4, 1 M NaCl, 1 mM EDTA, 1% Nonidet P-40, 0.1% SDS, 0.5% Sodium deoxycholate) and once with PNK buffer (20 mM Tris-HCl pH 7.4, 10 mM MgCl 2 , 0.2% Tween 20). Then, crosslinked nucleic acids were radiolabeled using ATP, [g-32P] (PerkinElmer, #BLU502A250UC) and T4 polynucleotide kinase (PNK, Thermofisher, #EK0032), washed twice with RIPA-S buffer and once with PNK buffer. Protein-RNA complexes are eluted and separated by SDS-PAGE. Then the gel was dried and complexes were visualized by exposure to a phosphoimager screen. In the case of nucleic acids extraction, RNA-proteins complexes were transferred onto nitrocellulose membrane 0.45 mm and complexes were visualized by exposure to a phosphoimager screen. The nitrocellulose membrane was cut out at the suspected molecular weight of the radiolabeled protein of interest. Nitrocellulose pieces were treated with proteinase K (1 mg/mL, ThermoScientific, #EO0491) in 200 mL of PK buffer (100 mM Tris-HCl pH 7.4, 50 mM NaCl, 10 mM EDTA), once without urea and then once with urea 7 M. Nucleic acids were purified with phenol/chloroform/isoamylic alcohol 25:24:1 (Sigma) and then precipitated overnight by adding Glycoblue (Thermofischer, #AM9515), sodium acetate and 100% ethanol. After centrifugation and washes (80% ethanol), nucleic acids pellet was resuspended in water and divided into three conditions. One condition without treatment, one with RNAse A treatment (1 mL at 10 mg/mL, ThermoScientific, #EN0531) and one with DNAse treatment (1 mLa t2 U / mL, Invitrogen, Beyond the 53BP1-RNA association on the lagging strand, our study provides a range of experiments to identify new RBPs at the replication fork and thus better understand the DNA replication process. Indeed, other proteins have been suggested

as candidate RBPs, such as Ku and MRE11, and by using the described techniques in our study, it would be possible to validate whether or not they can interact with the Okazaki fragments. For instance, MRE11 also possesses a GAR domain that is methylated by PRMT1, and Arginines mutations disrupt its exonuclease activity, which is necessary for MRE11 functions in response to replication stress or DNA damage (Boisvert et al., 2005a[START_REF] Boisvert | Methylation of MRE11 regulates its nuclear compartmentalization[END_REF]. Ku is an identified RBP [START_REF] Baltz | The mRNA-bound proteome and its global occupancy profile on protein-coding transcripts[END_REF][START_REF] Queiroz | Comprehensive identification of RNAprotein interactions in any organism using orthogonal organic phase separation (OOPS)[END_REF] and is known to play a role in the fork reversal mechanism. A recent study conducted in yeast also determined that the functions of Ku during replication stress are linked to RNA:DNA hybrids originating from Okazaki fragments.

In a fork reversal context, the last Okazaki fragment on the lagging strand hybridises with the nascent DNA of the leading strand, forming the regressed arm. The RNA primer contributes to establish the Ku barrier to protect against fork degradation (Audoynaud et al., 2023). Large-scale techniques such as iPOND (Dungrawala et al., 2015), coupled with RNA-binding activity techniques such as OOPS [START_REF] Queiroz | Comprehensive identification of RNAprotein interactions in any organism using orthogonal organic phase separation (OOPS)[END_REF], would potentially allow the identification and characterisation of new RNAprotein interactions at the replication forks.

RÉSUMÉ

Au cours de cette décennie, des centaines de protéines de liaison à l'ARN (RBP) ont été identifiées. Les interactions ARN-protéines sont essentielles pour de nombreux processus cellulaires, de la régulation posttranscriptionnelle de l'expression des gènes au maintien de l'intégrité du génome. La réplication de l'ADN est un mécanisme critique pour la stabilité du génome et des défauts peuvent entraîner des pathologies. Par ailleurs, des ribonucléotides sont produits pendant la réplication de l'ADN et plusieurs RBP sont présentes aux fourches de réplication.

ABSTRACT

Over the last decade, hundreds of RNA-binding proteins (RBPs) have been identified. The RNA-protein interactions are essential for many cellular processes, from post-transcriptional regulation of gene expression to maintenance of genome integrity. DNA replication is a critical mechanism in genome stability and defects can lead to diseases. Furthermore, ribonucleotides are produced during DNA replication and several RBPs are found at replication forks. However, their interaction with the RNA primer, present in the DNA template, remains unexplored. Using in vitro assays and UV-Cinduced crosslink methods in living human cells, the laboratory identified 53BP1 (p53-binding protein 1), involved in the response to DNA damage and replication stress, as an RBP that directly interacts with RNA-DNA species, similar to the RNA-DNA primers synthesized on the lagging strand to form the Okazaki fragments (OF). The aim of my thesis was to better characterise the interaction between 53BP1 and OF and to understand its role during DNA replication.

We found that the C-terminal part of 53BP1 is the minimum RNA-binding region. Furthermore, using a variety of approaches to study the association of proteins with nascent DNA during S-phase, we showed that the recruitment of 53BP1 to replication forks was decreased upon treatment with ribonuclease A or upon depletion of PRIM1, the catalytic subunit of primase. In contrast, depleting the endonuclease FEN1, which results in the accumulation of uncleaved RNA primers, led to an increased presence of 53BP1 at the fork. In addition, we found that 53BP1 depletion induced an accumulation of S-phase poly(ADP-ribose), which constitutes a sensor of unligated OF, and led to reduced DNA2/RPA loading at nascent DNA, which are factors of OF maturation. Altogether, these results show that 53BP1 interacts with OF to help in the processing of lagging strand, highlighting the role of a novel RNA-protein interaction at DNA replication forks. KEYWORDS 53BP1; Okazaki Fragments; DNA Replication; RNA-binding protein