ADAR-mediated RNA editing of DNA:RNA hybrids is required for DNA double strand break repair

ARTICLE ADAR-mediated RNA editing of DNA:RNA hybrids is required for DNA double strand break repair Sonia Jimeno1,2,6✉, Rosario Prados-Carvajal1,2,6, María Jesús Fernández-Ávila 2, Sonia Silva 1,2, Domenico Alessandro Silvestris3, Martín Endara-Coll4, Guillermo Rodríguez-Real1,2, Judit Domingo-Prim 4,5, Fernando Mejías-Navarro 1,2, Amador Romero-Franco1,2, Silvia Jimeno-González 1,2, Sonia Barroso 1,2, Valeriana Cesarini3, Andrés Aguilera 1,2, Angela Gallo3, Neus Visa 4& Pablo Huertas 1,2✉ The maintenance of genomic stability requires the coordination of multiple cellular tasks upon the appearance of DNA lesions. RNA editing, the post-transcriptional sequence alteration of RNA, has a profound effect on cell homeostasis, but its implication in the response to DNA damage was not previously explored. Here we show that, in response to DNA breaks, an overall change of the Adenosine-to-Inosine RNA editing is observed, a phenomenon we call the RNA Editing DAmage Response (REDAR). REDAR relies on the checkpoint kinase ATR and the recombination factor CtIP. Moreover, depletion of the RNA editing enzyme ADAR2 renders cells hypersensitive to genotoxic agents, increases genomic instability and hampers homologous recombination by impairing DNA resection. Such a role of ADAR2 in DNA repair goes beyond the recoding of specific transcripts, but depends on ADAR2 editing DNA:RNA hybrids to ease their dissolution. https://doi.org/10.1038/s41467-021-25790-2 OPEN 1Departamento de Genética, Universidad de Sevilla, Sevilla 41080, Spain. 2Centro Andaluz de Biología Molecular y Medicina Regenerativa-CABIMER, Universidad de Sevilla-CSIC-Universidad Pablo de Olavide, Sevilla 41092, Spain. 3RNA Editing Lab, Oncohaematology Department, IRCCS Ospedale Pediatrico “Bambino Gesù”, Viale San Paolo 15, 00146 Rome, Italy. 4Department of Molecular Biosciences, The Wenner-Gren Institute, Stockholm University, 10691 Stockholm, Sweden. 5 Present address: Moirai Biodesign SL, Parc Científic de Barcelona, 08028 Barcelona, Spain. 6 These authors contributed equally: Sonia Jimeno, Rosario Prados-Carvajal. ✉email: [email protected];[email protected] NATURE COMMUNICATIONS | (2021) 12:5512 | https://doi.org/10.1038/s41467-021-25790-2| www.nature.com/naturecommu nications 1 1234567890():,; Cells are continuously challenged by DNA damage. Among all kinds of insults that a DNA molecule has to deal with, double-strand breaks (DSBs) are the most dangerous. Indeed, just one unrepaired DSB is enough to either kill or terminally arrest cells. For these reasons when DSBs are formed, a complex cellular response—the DNA damage response (DDR)— is triggered in order to ensure the proper repair of such a threat to genomic integrity1. There are several pathways that can be used in order to repair a DSB and the choice between them is highly regulated. A eukaryotic cell can repair a DSB either by the simple re-ligation of the DNA ends (a process known as Non-Homologous End-Joining, NHEJ)2or by a homology-driven repair event. There are different routes among the repair pathways that use homologous regions for repair, all of which are grouped in a process called homologous recombination (HR)3. All HR events share a first biochemical step called DNA resection, which is the key to decide the pathway that will be eventually used to repair the DSB4,5. This process consists of the nucleolytic degradation of the DNA ends of the break that produces tails of 3' ended single-stranded DNA (ssDNA), that are rapidly protected by the RPA protein complex. In recent years, the importance of RNA and RNA-related factors in DNA repair has become clear6–9. Indeed, many RNA- related proteins have been shown to be targets of the DNA damage-induced post-translational modifications10–12. Also, direct roles of specific RNA-related factors in DNA repair have been recently reported (for a review see9). Moreover, the RNA molecule itself seems to impact DNA repair. Several labs have shown the formation of DNA:RNA hybrids around DSBs in different eukaryotes, either dependent on previous transcription13,14 or upon de novo transcription of the broken chromatin15,16. The relevance of such RNA molecules is still under debate, with both pro- and anti-repair effects ascribed to them9. An important co-transcriptional RNA modification that, so far, has not been extensively studied in its putative relationship with DNA repair and the response to DNA damage is RNA editing. This process alters RNA sequences by the action of specific deaminases that convert one base into another. Every mammalian transcript can be subjected to RNA editing17–19. RNA editing can be classified into several categories20, including adenosine-to-inosine (A-to-I) deamination, which is accomplished by a family of RNA-specificadenosinedeaminases known as ADARs18,19. This family is formed by ADAR1, ADAR2 (also known as ADARB1), and ADAR3; however, only ADAR1 and ADAR2 have been shown to present catalytic activity. A-to-I deamination is the most abundant form of RNA- editing in mammals and defects in this process are associated with human diseases, such as disorders of the central nervous system21 or pediatric astrocytomas22. Only limited information has been published regarding the connection of A-to-I editing and DNA damage, albeit at least the mRNA of NEIL1, has been shown to be re-coded by ADAR1 to alter its enzymatic properties23. Moreover, A-to-I editing has been proposed to be involved in the pathogenesis of cancer24,25. Here, we show that the general pattern of ADAR2-mediated A- to-I editing changes upon DSB formation. Such changes depend on the DDR, specifically the ATR kinase and the resection protein CtIP. As a consequence, ADAR2 is required for the maintenance of genomic integrity and, specifically for DNA end resection and HR. Strikingly, mRNAs from either resection-related or recombination-related genes are not affected by ADAR2. Instead, ADAR2 role in resection is related to its ability to edit DNA:RNA hybrids. Not only do such structures increase when ADAR2 is depleted, but this protein physically and functionally interacts with the BRCA1-SETX complex for this role. Results RNA editing changes after DNA damage. As previously mentioned, crosstalk between RNA metabolism and DNA repair has been extensively documented9, but a connection between DNA repair and RNA editing has not been extensively analyzed. Thus, we wanted to study whether the appearance of DNA damage had any effect on RNA editing. In order to explore this possibility, we used a previously published reporter system (RNAG) that measures levels of RNA editing using the accumulation of the fluorescent proteins GFP and RFP26. This system bears both the RFP and GFP ORFs in a single transcript, with a stop codon between them (Fig. 1A). So, cells bearing such reporter express RFP constitutively, but GFP is only produced if an RNA editing event changes the A of the stop codon to an I (Fig. 1A)26. Therefore, the number of red cells that are also green indicates the efficiency of RNA editing. As a control to discard other effects non-related to the editing on this system, we used the RNWG control reporter, in which the stop codon is pre-edited, so all cells bearing the construct fluoresce, indeed, in red and green26.In U2OS cells stably transfected with the reporter, we observed that DNA damage induced by ionizing radiation increased GFP expression by 50% specifically in the RNAG reporter and not in the RNWG control, in agreement with a DNA damage stimulation of RNA A-to-I editing in this system (Fig. 1B). Similar results were obtained when using the DNA damage-inducing drug camptothecin (Fig. 1C), where we could observe a dosedependent effect on RNA editing stimulation. One possibility is that DNA damage induces the accumulation of the A-to-I editing machinery, namely ADAR1 and ADAR2 enzymes, thus increasing this process. However, neither of these proteins was upregulated, but were slightly downregulated, upon exposure to IR (Supplementary Fig. 1A). Then, in order to confirm this was a canonical induction of RNA editing, we depleted the A-to-I editing machinery. To choose which member of the ADAR family to downregulate, we revisited the data we obtained in a previous genome-wide screening for factors that unbalance the choice between DSB repair pathways27. Interestingly, both ADAR1 and ADAR2, but not the catalytically inactive ADAR3, skewed DSB repair towards end-joining (Supplementary Figure 1B), and this was not due to changes in the cell cycle (Supplementary Fig. 1C). However, the effect was more prominent and clearer upon ADAR2 depletion. Indeed, downregulation of ADAR2 severely compromised both the basal and the DNA damage-induced expression of the GFP in the RNAG (Fig. 1D; for ADAR2 depletion efficiency see Supplementary Fig. 2A), but, as expected, not in the RNWG control reporter (Supplementary Fig. 2B). To better understand this phenomenon, we decided to look for DDR factors that affected the DNA damage-induced RNA editing. Recently, we have found that CtIP, a core DNA end resection factor that is also required for ATR activation, plays additional roles in DNA damage-induced RNA splicing28. Interestingly, we could see that CtIP downregulation specifically eliminated the DNA damage-dependent induction of RNA editing without affecting the basal levels (Fig. 1D). Again, CtIP depletion did not alter GFP levels in the control RNWG system (Supplementary Fig. 2B). We could also complement this effect with the expression of siRNA-resistant flag-tagged CtIP in CtIP depleted cells, to the same extent as the control cells transfected with a non-targeting siRNA, even though overexpression of FLAG-CtIP on its own reduced the intensity of this DNA damage-induced phenotype (Supplementary Fig. 2C). The general response to DNA damage is mainly controlled by the activation of two related apical kinases, ATM and ATR1. Thus, we also tested if any of them was required for the induction of RNA editing upon irradiation. Interestingly, ATM inhibition did not affect DNA damage-induced RNA editing, while ATR ARTICLE NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-021-25790-2 2NATURE COMMUNICATIONS | (2021) 12:5512 |https://doi.org/10.1038/s41467-021-25790-2| www.nature.com/naturecommunications inhibition decreased the DNA damage-induced editing increase (Fig. 1E). This agrees with the notion that ATR and CtIP act on the same branch of the DNA damage checkpoint in response to DSBs29. The lack of response with the ATM inhibitor could be explained by a compensation by another member of the PIKK family, most likely DNA-PK. Along those lines, the ATR effect could also be affected by this phenomenon. Thus, we repeated the experiment with ATM, ATR, and DNA-PK inhibitors in different combinations (Fig. 1F). As shown, inhibition of ATR suppressed the DNA damage-induction of RNA editing, regardless of the presence of the inhibitors of ATM or DNA-PK. Interestingly, chemical inhibition of DNA-PK showed a limited increase in the basal levels of RNA editing, but importantly the exposure to DNA damage still provoked a hyperactivation of the process. Notably, AB C RNA Editing GAU NLS GFP RFP GIU NLS GFP RFP * RNWG RNAG 0.0 0.5 1.0 1.5 2.0 % Red cells positive for GFP normalized to untreated cells -IR +IR RNWG RNAG 0.5 1.0 1.5 2.0 0,001 mM 0,005 mM 0,05 mM 0,1 mM 0 mM % Red cells positive for GFP normalized to DMSO siNT siCtIP siADAR2 0 5 10 15 20 25 % Red cells positive for GFP *** *** * -IR +IR * D - IR +IR 0 5 10 15 % Red cells positive for GFP ** *** * * DMSO ATRi ATMi E DMSO ATRi ATMi DNA-PKi ATRi + ATMi ATRi + DNA-PKi ATMi + DNA-PKi 0 5 10 15 20 25 * * ** % Red cells positive for GFP F -IR +IR Fig. 1 DNA damage increases RNA editing. A Scheme of the RNAG editing system. A bi-cistronic mRNA containing the RFP and GFP sequences is produced. The presence of a stop codon impedes the expression of the GFP ORF, except when the adenine is edited to inosine. The presence of a secondary structure containing such stop codon allows its recognition and deamination of the adenine by ADAR proteins. BDNA damage-induced RNA editing. The plot shows the percentage of cells bearing the RNAG reporter or the constitutively edited RNWG system that express both the RFP (red cells) and GFP (green cells). Cells were either irradiated (+IR; 10 Gy; black bars) or mock-treated (–IR; white bars) and incubated for 12 h. The percentage of green cells over 10.000 red cells were analyzed on BD FACSAriaTM using FACSDiva v5.0.3 software. For each reporter, the ratio of green and red cells was normalized with the untreated conditions. Statistical significance was determined with a two-tailed paired Student’st-test. CSame as B, but cells were treated with the indicated concentration of camptothecin (CPT). DCells bearing the RNAG reporter were transfected with the indicated siRNAs and irradiated or not, and the percentage of red cells that were also green is plotted. Statistical significance was determined with a two-way ANOVA. ESame as D, but cells were pretreated for 2 h with 10 μM of inhibitors of ATM (ATMi), ATR (ATRi), or DMSO as control, previous to the irradiation. Cells were collected to check for editing levels 10 h after irradiation. The inhibitors were kept for the duration of the experiment. Statistical significance was determined with a two-way ANOVA. FSame as E, but, cells were also treated with DNA–PK inhibitor (DNA–PKi), as well as the double combinations of the ATM, ATR, and DNA–PK inhibitors, as indicated. Statistical significance was determined with a two-way ANOVA. The average and the standard deviation of the medians of four (panels B and F) or three (panels C–E) independent experiments are shown. Each individual replica is marked with a colored symbol. One, two, or three asterisks represent p< 0.05, p< 0.01, or p< 0.001, respectively. Actual p-values can be found in the Source data file. Only biological relevant comparisons are shown. NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-021-25790-2 ARTICLE NATURE COMMUNICATIONS | (2021) 12:5512 |https://doi.org/10.1038/s41467-021-25790-2| www.nature.com/naturecommunications 3 concomitant inhibition of both DNA-PK and ATM abolished the induction of RNA editing caused by irradiation. Thus, it seems that those two kinases could have an overlapping role in this phenomenon. ADAR2 depletion causes genomic instability and DNA damage sensitivity. To understand the consequences of reduced A-to-I RNA editing for genomic stability, we decided to globally reduce such RNA modifications. Based on our previous data with ADAR2 (Fig. 1D and Supplementary Fig. 1B), we decided to use the downregulation of this protein as a tool to reduce A-to-I RNA editing. Strikingly, and in agreement with a role in maintaining genomic stability, the depletion of ADAR2 impaired DSB repair, measured as the presence of γH2AX foci 24 h after irradiation. Spontaneous DNA damage accumulated in the absence of any exogenous genotoxic agent in ADAR2-depleted cells (Fig. 2A). Confirming a DSB repair impairment, the disappearance of γH2AX foci upon exposure to ionizing radiation was delayed (Fig. 2B). Indeed, repair levels of DSBs at 6 and 24 h after irradiation in ADAR2-depleted cells were similar to those observed after downregulation of the critical repair factor CtIP (Fig. 2B). Interestingly, and in agreement with an increased burden of spontaneous DNA damage, in the absence of ADAR2, we observed a significant increase of BRCA1 foci in cells unchallenged with any genotoxic agent (Fig. 2C). A similar effect was observed upon ADAR1 downregulation (Fig. 2D). Furthermore, micronuclei accumulated at high levels in ADAR2-depleted cells, regardless of the exposure to an external source of DNA damage (Fig. 2E). Finally, and confirming the role of ADAR2 in DNA repair and the maintenance of genomic stability, its depletion rendered cells hypersensitive to DSBs-inducing agents such as ionizing radiation or camptothecin (Fig. 2F, G). ADAR2 depletion affects DNA repair pathway choice. Next, we decided to test a possible requirement of A-to-I RNA editing for DSB repair. As mentioned, ADAR2 depletion skewed the balance between HR and NHEJ towards the latter (see ref. 27and Supplementary Fig. 1B), suggesting that recombination might be compromised. Indeed, both RAD51-dependent gene conversion (GC) and RAD51-independent Single Strand Annealing (SSA), two types of homology-dependent repair (HDR), were reduced in cells downregulated for ADAR2 (Fig. 3A, B). In stark contrast, there was no impact on NHEJ efficiency (Fig. 3C), arguing that ADAR2 was particularly required for HR. The cell cycle is a major regulator of DSB repair pathway choice, as HR is limited in G1. However, the observed HR defect was not caused by an accumulation of G1 cells (Supplementary Fig. 1C). Thus, we conclude that ADAR2 facilitates repair by HR. DNA resection requires ADAR2 editing activity. Due to its effect in recombination, we hypothesized that ADAR2 might have a role in the common, early steps of the homology-dependent repair pathways, namely in DNA end resection. To test this idea, we first studied RPA foci formation upon ionizing radiation in ADAR2-depleted cells. RPA is an ssDNA binding complex that accumulates at DNA breaks as a direct consequence of DNA end resection4,5,30. Thus, the percentage of RPA foci-positive cells is the standard readout of resection in mammalian cells. Depletion of ADAR2 in U2OS cells caused a significant defect in resection, though less pronounced than that observed with the downregulation of the key resection factor CtIP (Fig. 4A and Supplementary Fig. 3A). For representative images of the experiment see Supplementary Fig. 3A. The same resection impairment was also observed upon depletion of ADAR2 in the HeLa cells (Supplementary Fig. 3B). Strikingly, similar results were observed upon depletion of ADAR1, but not the catalytically dead member of the ADAR family, ADAR3, thus suggesting that active RNA editing is required for DNA end resection (Fig. 4A and Supplementary Fig. 3A). To confirm that the observed phenotype in DNA resection was truly due to the reduction of ADAR2 levels and not to an indirect off-target effect, we studied RPA foci formation in cells bearing siRNA-resistant, GFP-tagged variants of ADAR2. Indeed, the resection impairment caused by depletion of ADAR2 was rescued by wild-type GFP-ADAR2 (Fig. 4B and Supplementary Fig. 3C). Importantly, this rescue was not observed with the expression of a catalytically dead version of the protein (GFP-ADAR2E-A) (Fig. 4B), arguing that ADAR2 deaminase activity was required for processive resection. Moreover, we could reproduce the resection defect in U118 cells31, a glioblastoma cell line that is defective for ADAR2 expression, when compared with the same cells complemented with a wild-type copy of the gene, but not a catalytically dead mutant (Fig. 4C). To validate these observations, we analyzed recruitment of RPA to DSBs by other means. U2OS cells depleted for ADAR2, or CtIP as a control, were laser micro-irradiated and immunostained with antibodies against RPA and γH2AX to identify the irradiated areas. The percentage of γH2AX-positive stripes that were costained by RPA was determined (Fig. 4D). In agreement with our previous results, depletion of ADAR2, or CtIP, significantly diminished the presence of RPA at the irradiated areas. Finally, in order to analyze in more detail the resection defect related to ADAR2 depletion and to investigate whether only resection initiation was impaired or if resection processivity was also compromised, we used SMART, a high-resolution technique that measures resection in individual DNA fibres32,33. As seen in Fig. 4E, the length of ssDNA fibres formed during the resection process was reduced upon ADAR2 depletion. Again, similar results were observed upon ADAR1 downregulation (Supplementary Fig. 3D), reinforcing the connection between A-to-I editing and DNA end resection. Altogether, these results confirm that RNA editing by ADAR proteins facilitates DNA end resection at DSBs. The role of ADAR2 on resection does not rely on the recoding of mRNAs that encode resection factors Next, we wondered how this RNA editing activity might be needed for DNA end processing. We studied the recruitment DSB repair factors, such as 53BP1, BRCA1, and CtIP, to DNA damage foci shortly after DNA damage induction upon ADAR2 depletion. Notably, neither of them was affected in cells exposed to ionizing radiation or laser micro-irradiation (Supplementary Fig. 4A–C). Then, we wondered if ADAR2 was specifically editing mRNAs that code for resection factors. To analyze this possibility, we exposed the ADAR2-defective cell line U118 to ionizing radiation or mock treatment, isolated total RNA, and sequenced it. U118 cells complemented with either wild-type ADAR2 or a catalytically dead mutant were also processed in parallel. The levels of ADAR2 mRNA in the different samples are shown in Supplementary Fig. 4D. As expected, U118 cells complemented with wild-type ADAR2 showed a higher efficiency in editing the coding codons of known ADAR2 targets, expressed as the weighted average over all known recoding sites, known as the REI34. Instead, non-complemented U118 cells showed little recoding editing, regardless of whether or not exposed to ionizing radiation (Fig. 4F). Equally expected, the expression of a catalytically dead enzyme also showed almost no recoding of mRNAs (Fig. 4F), despite the fact that such variant was expressed almost 3 times more than the wild-type ADAR2 (Supplementary Fig. 4D). Thus, only the expression of catalytically active enzyme led to the expected ADAR2-dependent recoding due to editing of specific codons (Fig. 4F). Interestingly, although some specific ARTICLE NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-021-25790-2 4NATURE COMMUNICATIONS | (2021) 12:5512 |https://doi.org/10.1038/s41467-021-25790-2| www.nature.com/naturecommunications coding codons were edited more efficiently upon irradiation than in mock-treated cells, we observed a general decrease in the recoding editing efficiency of known ADAR2 substrates upon exposure to DNA damage (Fig. 4F and Supplementary Data 1). Moreover, we could not observe any change in the editing of mRNA from genes that code for recombination or resection factors either in cells exposed to DNA damage or in undamaged cells (Supplementary Table S1). Thus, we conclude that the role of ADAR2 in resection and recombination does not rely on changes in the sequence or expression of specific mRNAs of bona fide DNA repair factors. To integrate our data, in which we observed a general ADAR2-mediated increase in RNA editing in the reporter A 0 24 0 20 40 60 80 siNT siADAR2 Time after IR (h) * *** γH2AX siADAR2siNT DAPI 0 h 24h siADAR2siNT % of γH2AX positive cells- 16 24 0 20 40 60 80 100 % of γH2AX positive cells - siNT siADAR2 siCtIP B Time after IR (h) 0 20 40 60 ** siADAR2 siNT % BRCA1 foci-positive cells C 0 10 20 30 40 siNT siADAR2 % of micronucleus-positive cells * * -IR +IR E siNT siADAR2 00.0050.010 0 50 100 CPT ** * 024 0 20 40 60 80 100 Gy * % Survival % Survival siNT siADAR2 FG 0 20 40 60 * siADAR1 siNT % BRCA1 foci-positive cells D Fig. 2 ADAR2 depletion causes genetic instability and DNA repair defects. A Percentage of cells positive for γH2AX foci upon spontaneous accumulation (0 h) or 24 h post irradiation with 10 Gy in U2OS cells transfected either a siRNA against ADAR2 or with control siNT. Quantification is shown on the left. A representative image is shown on the right. Scale bars represent 10 µm. BRepair kinetics is shown as the disappearance of γH2AX foci 1, 6, and 24 h post irradiation with 10 Gy in U2OS cells transfected either a siRNA against ADAR2, CtIP or with control siNT. CPercentage of spontaneous BRCA1 foci-positive cells in cells transfected with either a siRNA against ADAR2 or with control siNT. The average and SD of three independent experiments is shown. DSame as Cbut in cells depleted for ADAR1. EPercentage of cells positive for micronucleus without exposure to DNA damage (-IR) or 24 h post irradiation with 10 Gy (+IR) in U2OS cells transfected either with a siRNA against ADAR2 or with control siNT. Other details as in (A). FClonogenic assays of U2OS cells depleted with a siRNA against ADAR2 or with control siNT after treatment with different doses of IR. Other details as in (A). GSame as Fbut cells treated with camptothecin (CPT; μM; right). In all panels, the average and SD of three independent experiments are shown and statistical significance was determined with a two-tailed paired Student’st-test. Each individual replica is marked with a colored symbol. One, two, or three asterisks represent p< 0.05, p< 0.01, or p< 0.001, respectively. Actual p-values can be found in the Source data file. Only biological relevant comparisons are shown. NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-021-25790-2 ARTICLE NATURE COMMUNICATIONS | (2021) 12:5512 |https://doi.org/10.1038/s41467-021-25790-2| www.nature.com/naturecommunications 5 (Fig. 1) accompanied by a general reduction of its activity on known targets (Fig. 4F), we hypothesized that upon DNA damage ADAR2 is mobilized to induce RNA editing at new sites. Such redistribution could mean that at least a fraction of the protein would localize at sites of DNA-DSBs. To test this idea, we analyzed the presence of GFP-tagged ADAR2 at damaged chromatin at different time points by laser micro-irradiation (Fig. 4G). In agreement with our idea, a fraction of GFP-ADAR2 was readily recruited to sites of DNA damage as early as 5 minutes after laser micro-irradiation, something not observed in cells expressing only GFP as a control. In fact, 50% of cells showed colocalization between γH2AX and ADAR2 stripes. Interestingly, such recruitment seemed transient, as it was no longer observed 30 min after irradiation. This recruitment pattern agrees with the idea that ADAR2, or at least a fraction of the protein, changes its substrates and is channeled towards RNAs at the sites of broken chromatin to play a role in the early steps of DSB repair. ADAR2 facilitates resection over DNA:RNA hybrids.We hypothesized that the ADAR2-depletion related phenotype in DNA resection could be caused by a direct effect of ADAR2 on an RNA molecule located in the vicinity of the break that would act as a physical barrier for DNA end processing. The presence of DNA:RNA hybrids close to DSBs has been documented from yeast to mammals, both as pre-existing R-loops, R-loops formed as a consequence of breaks in transcribed regions or as DNA:RNA hybrids resulting from de novo transcription of resected DNA ends9,13,16,35–37. The actual effect of such DNA:RNA hybrids in resection is controversial, with both pro- and anti-resection effects described9,16,38. Importantly, ADAR2 has been proposed to recognize and edit DNA:RNA hybrids in vitro39. In order to define whether ADAR2 involvement in DNA end resection depended on the presence of DNA:RNA hybrids, we repeated the resection assay in the presence or absence of ectopically overexpressed RNaseH1, an enzyme that degrades the RNA moiety of such structures40. ADAR2 depletion and RNaseH1 overexpression efficiency are documented in Supplementary Fig. 5A. Strikingly, the overexpression of RNaseH1 in U2OS reverted ADAR2 resection phenotype as measured by RPA foci accumulation (Fig. 5A and Supplementary Fig. 5B). Indeed, the mere overexpression of RNaseH1 facilitated RPA foci formation even in cells transfected with a control siRNA, arguing that DNA:RNA hybrids act generally as physical barriers for the resection process, and that ADAR2 helps overcome such roadblocks. To confirm this finding, we studied the recruitment of RPA to damaged chromatin in HeLa cells depleted of ADAR2 and overexpressing RNaseH1. Again, such overexpression rescued the resection phenotype of ADAR2 depletion (Fig. 5B). The same was observed, albeit only partially, when laser microirradiation experiments were performed (Fig. 5C). In none of those cases, the increase in RPA-positive cells was due to changes AB 0.0 0.5 1.0 1.5 DR-GFP HR efficiency normalized to control ** ** siNT siCtIP siADAR2 0.0 0.5 1.0 1.5 2.0 EJ5-GFP siNT siCtIP siADAR2 ** 0.0 0.5 1.0 1.5 SA-GFP ** siNT siCtIP siADAR2 SSA efficiency normalized to control NHEJ efficiency normalized to control GFP iGFP GFP iGFP GFP iGFP I-SceI target site 2.7 kb 2.7 kb 5’GFP 3’GFP 5’GFP 3’GFP GFP I-SceI target site GFP Puro GFP GFP GFP NHEJ I-SceI target site C Fig. 3 ADAR2 depletion affects homologous recombination. A Effect of ADAR2 depletion in the DR-GFP reporter. A scheme of the reporter is shown on the top. Induction of a DSB using I-SceI meganuclease renders GFP-positive cells when the donor repeat (iGFP) is used in a gene conversion event. The efficiency of classical recombination (HR) was calculated as the percentage of GFP-positive cells in response to I-SceI expression upon downregulation of the indicated genes and normalized with the control. The average and standard deviation of at least three independent experiments are shown. BSame as Abut using the Single Strand Annealing (SSA) reporter SA-GFP (top). In this case, the induction of a DSB located between two repeats in direct orientation will render GFP-positive cells only when intramolecular SSA takes place. CSame as Abut using the non-homologous end-joining (NHEJ) reporter EJ5-GFP (Top). In this case, two I-SceI-induced DSBs could be repaired by conservative or mutagenic NHEJ granting the accumulation of functional GFP. In all panels, the average and SD of three independent experiments are shown and statistical significance was determined with a two-tailed paired Student’s t-test. Each individual replica is marked with a colored symbol. One, two, or three asterisks represent p< 0.05, p< 0.01, or p< 0.001, respectively. Actual p-values can be found in the Source data file. Only biological relevant comparisons are shown. ARTICLE NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-021-25790-2 6NATURE COMMUNICATIONS | (2021) 12:5512 |https://doi.org/10.1038/s41467-021-25790-2| www.nature.com/naturecommunications in cell cycle profile when the RNaseH1 was overexpressed (Supplementary Fig. 5C). Then, to assess whether ADAR2 helped remove DNA:RNA hybrids, we analyzed the accumulation of such structures by immunofluorescence using the DNA:RNA hybrid-specific antibody S9.641. As shown in Fig. 5D, depletion of ADAR2 increases the nuclear signal with that antibody. To rule out the contribution of other nucleic acid structures to the increase in the S9.6 signal, we overexpressed RNaseH1 and observed a significant reduction in the staining (Fig. 5D). Of relevance, ADAR2 has been previously found to be a part of the so-called “DNA:RNA hybrid interactome”42. Therefore, we decided to test if ADAR2 could interact directly with DNA:RNA hybrids. Indeed, ADAR2 was specifically immunoprecipitated using the S9.6 antibody, in a similar fashion as Senataxin (SETX), a helicase described to dissolve such hybrids and that has been shown to be recruited to DSBs at transcribed regions13 (Fig. 5E). This immunoprecipitation was specific, as did not occur when a control antibody was used (Fig. 5E). Taken together, our results suggest the ADAR2 effect in resection was caused by DNA:RNA hybrid accumulation. 0 20 40 60 80 siADAR2 siNT siCtIP % RPA foci positive cells 0 20 40 60 80 siNT siADAR2 * GFP-ADAR2 GFP-ADAR2-E/A % RPA foci positive cells 0 20 40 60 80 100 % RPA stripes positive cells *** ** A GFP B 0 20 40 60 80 *** *** U118 cells % RPA foci positive cells WT-ADAR2 ADAR2-E/A empty C siNTsiCtIP γH2AX RPA D siADAR2 * * * *** 0 50 100 150 200 Resection tracts length (Micons) E siADAR2 siNT siADAR2 siNT siCtIP empty ADAR2 ADAR2-E/A 0.0 0.2 0.4 0.6 0.8 1.0 Recoding Index untreated irradiated F * siADAR1 siADAR3 GγH2AX GFP DAPI GFPGFP-ADAR2GFPGFP-ADAR2GFPGFP-ADAR2 5 min 15 min 30 min NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-021-25790-2 ARTICLE NATURE COMMUNICATIONS | (2021) 12:5512 |https://doi.org/10.1038/s41467-021-25790-2| www.nature.com/naturecommunications 7 A prediction of this model is that the decrease in HR caused by ADAR2 downregulation should be suppressed by RNaseH1 overexpression. In fact, recombination was almost completely restored in ADAR2-depleted cells when such enzyme was ectopically expressed (Fig. 5F). Such effect was not due to a general increase of recombination mediated by RNaseH1 overexpression, as it was not observed either in CtIP depleted or control cells. Increase of DNA:RNA hybrids generally impairs DNA end resection. Due to our observation that DNA:RNA hybrids hampered resection in the absence of ADAR proteins, we wondered if other enzymes involved in the removal of hybrids also affected DNA end resection, therefore this represented a more general phenomenon. We decided to test SETX, due to its aforementioned relationship with DSBs. So, we checked if its loss also affected RPA foci formation. Indeed, depletion of Senataxin also produced a DNA resection defect measured by RPA foci accumulation (Fig. 6A) and reduced the length of resected DNA (Fig. 6B). As seen for ADAR2 depletion, SETX downregulation also led to an increased burden of spontaneous DNA damage, measured as BRCA1 foci in unchallenged cells (Fig. 6C). ADAR proteins physically interact with Senataxin and BRCA1. Our data suggested a possible role of ADAR2 in R-loop resolution, which would be key for facilitating DNA end resection and HR. Hence, we wondered if ADAR2 might interact with known players in the homeostasis of R-loops that are also connected with DSB repair. Many different enzymes have been associated with the removal of R-loops. Among them, we decided to focus on the BRCA1–SETX complex. These two proteins have been shown to cooperate in the elimination of R-loops in the 3' end of many transcribed genes43. Moreover, both proteins play roles in DNA end resection, as it has been previously established for BRCA133 and in this study for Senataxin (Fig. 6A–C). Thus, we tested whether ADAR proteins might physically interact with BRCA1 and Senataxin. Using antibodies against BRCA1, we confirmed the previously shown interaction of this protein with Senataxin43 and also established an interaction with ADAR1 and ADAR2 (Fig. 6D). An additional, independent co-immunoprecipitation experiment between ADAR2 and BRCA1 can be found in Supplementary Fig. 6A. These interactions were not stimulated by the presence of DNA damage, at least measured 1 h after irradiation (Fig. 6D). Also, by co-immunoprecipitations with different antibodies we could observe the reciprocal interaction of ADAR proteins with both BRCA1 and SETX (Fig. 6E–F). The possibility that ADAR immunoprecipitation could be bringing down any proteins accumulating at sites of DNA DSBs was excluded, since we could not observe an interaction of ADAR2 with the DNA- end binding protein Ku80, suggesting that those interactions were, indeed, specific. It is worth pointing out that these coimmunoprecipitations were performed in the presence of benzonase, thus they are not bridged by DNA. Moreover, immunoprecipitation using an antibody against SETX also allows the coprecipitation of ADAR2 (Fig. 6G). Additional, independent coimmunoprecipitation experiments between ADAR1, ADAR2, and SETX can be found in Supplementary Fig. 6B. Therefore, we could confirm that these proteins interact in a DNA damageindependent fashion, most likely to facilitate the removal of DNA:RNA hybrids globally. Discussion mRNA post-transcriptional modifications are now being revealed as potent regulators of cellular metabolism that allow flexibility and adaptability to a changing environment44. Such chemical modifications of the mRNA can recode coding sequences, create or destroy splicing sites, affect RNA stability and structure, or directly interact with specific readers in order to recruit specific machinery affecting translation and/or mRNA localization. Therefore, those modifications affect virtually every aspect of the biological response of the cells. In fact, it has been recently shown that other base modifications of RNA happen at DNA:RNA hybrids and contribute to genomic stability45. Thus, it was reasonable to expect that A-to-I RNA editing would contribute to the global response to DNA damage. Indeed, our results support this notion, uncovering a bona fide role of A-to-I deamination by ADAR1 and ADAR2 in the maintenance of genomic integrity upon exposure to genotoxic agents. ADAR2-mediated RNA editing seems, indeed, completely upturned upon treatment with Fig. 4 ADAR depletion impairs in DNA resection. A DNA resection proficiency after 10 Gy of irradiation in U2OS cells measured as the percentage of RPA foci-positive cells in cells transfected either with siRNAs against ADAR1, ADAR2, ADAR3, CtIP or with control siNT. The average and SD of three independent experiments are shown. Each individual replica is marked with a colored symbol. Significance was determined by two-tailed Student’st-test comparing each condition to siNT cells. *P< 0.05. Actual p-values can be found in the Source data file. Representative images of the experiments are shown on Supplementary Fig. 3A. BDNA resection proficiency measured as the percentage of RPA foci-positive cells in U2OS cells expressing either GFPADAR2 wild type or a catalytically dead version of the protein (ADAR2 E/A) transfected either with a siRNA against the 3'UTR of ADAR2 (black boxes) or a control siNT-UTR (white boxes). Each individual replica is marked with a colored symbol. Significance was determined by two-tailed Student’st-test comparing each condition to siNT cells. *P< 0.05. Actual p-values can be found in the Source data file. Other details as in (A). CRPA foci formation upon in U118 cells treated with 10 Gy of radiation in cells expressing either GFP, GFP-ADAR2 wild type or a catalytically dead version of the protein. Each individual replica is marked with a colored symbol. Significance was determined by two-tailed Student’st-test. ***P< 0.001. Actual p-values can be found in the Source data file. Other details as in (A). DDNA resection proficiency was measured as RPA stripes-positive cells upon laser microirradiation in cells transfected either an siRNA against ADAR2, CtIP or with control siNT. The average and SD of four independent experiments are shown. Each individual replica is marked with a colored symbol. Representative images of the experiments are shown on the right. Scale bars represent 10 µm. Significance was determined by two-tailed Student’st-test comparing each condition to siNT cells. **P< 0.01; ***P< 0.001. Actual p-values can be found in the Source data file. Other details as in (A). EResection length measured with the SMART assay using DNA fibers extracted from U2OS downregulated for ADAR2. A nontarget siRNA (siNT) was used as control. One out of three representative experiment with similar results is shown. Significance was determined by twotailed Student’st-test comparing. ***P< 0.001. Actual p-values can be found in the Source data file. Other details as in (A). FRNA sequencing of U118 cells complemented with a plasmid bearing wild type ADAR2, catalytically dead ADAR2 E/A or the empty vectors in untreated conditions (black bars) or upon exposure to 10 Gy of ionizing radiation (white bars) was used to analyze the changes in RNA sequence of codons known to be edited by ADAR2. The Recoding Editing Index (REI) was reported as percentage. The average of two independent experiments is shown. Each individual replica is marked with a colored symbol. GU2OS cells bearing GFP-ADAR2 or GFP, as a control, were micro-irradiated using a laser as described in the methods section. Cells were fixed at the indicated time points and the presence of γH2AX (red) or ADAR2 (green) at lasers stripes was analysed. Representative images out of three independent experiments are shown. Scale bars represent 10 µm. ARTICLE NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-021-25790-2 8NATURE COMMUNICATIONS | (2021) 12:5512 |https://doi.org/10.1038/s41467-021-25790-2| www.nature.com/naturecommunications DNA damaging agents. Many of the codons in mRNAs that are usually targeted by ADAR2 decrease their editing, whereas specific codons on natural mRNAs and in other RNA species such as the RNAG reporter increase their editing. Based on these findings, we propose an RNA-editing DDR (REDAR) that is essential for DNA repair, and specifically for HR, contributing to the maintenance of genomic integrity. Specifically, we postulate that upon the triggering of the DDR, REDAR is activated by ATR and CtIP. Then, ADAR2, and maybe ADAR1, is mobilized from its usual targets to new ones, most likely including DNA:RNA hybrids, in order to help with their dissolution. In this regard, ADAR2 action might have two nonmutually-exclusive outcomes; on the one hand, it can rapidly and transiently relocate to promote RNA editing at the sites of DSBs, aiding in the removal of DNA:RNA hybrids and facilitating DNA end resection; and on the other hand, it can increase the editing of a small fraction of yet-undisclosed mRNAs, whose role in the DDR should be clarified in further studies. Interestingly, it has been previously shown that ADAR action can alter the biochemical properties of some DNA repair enzymes23. We envision that the transcriptional inhibition induced by the DDR46,47 will cause a reduction of the normal co-transcriptional RNA editing48 and will free ADAR to act on alternative substrates such as the aforementioned DNA:RNA hybrids. We favor the idea that the editing itself would happen on the RNA moiety of the structure. However, by performing an in silico study we observed that A B 0 10 20 30 40 50 ** % RPA positive HeLa cells DAPIJH2AXRPA siADAR2 siNTsiADAR2 siNT - RNAse H1 + RNAse H1 C siNT siADAR2 0 20 40 60 80 100 % RPA positive cells - RNase H1 + RNase H1 0 20 40 60 80 100 %H2AX positive U2OS cells with RPA foci * * * - RNase H1 + RNase H1 siADAR2 siNT siADAR2 siNT - RNase H1 + RNase H1 S9.6 RNaseH1 DAPI +S9.6 +RNaseH1 - siNT siADAR2 +-+ Relative S9.6 Intensity **** **** RNaseH1 -+-+ siNT siADAR2 0.0 0.5 1.0 1.5 2.0 2.5 1 1 1.15 0.96 D ADAR2 SETX Input S9.6 IgG IP E siNT siADAR2 siCtIP 0.0 0.5 1.0 HR efficiency -RNase H1 +RNase H1 * * F Size ladder 235 kDa 180 kDa 75 kDa NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-021-25790-2 ARTICLE NATURE COMMUNICATIONS | (2021) 12:5512 |https://doi.org/10.1038/s41467-021-25790-2| www.nature.com/naturecommunications 9