Rescuing bacterial genome replication: essential functions to repair a double-strand break and restart DNA synthesis

Rescuing bacterial genome replication: essential functions to repair a double-strand break and restart DNA synthesis | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Article Rescuing bacterial genome replication: essential functions to repair a double-strand break and restart DNA synthesis Charles Winterhalter, Stepan Fenyk, Heath Murray This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-6364374/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 26 Nov, 2025 Read the published version in Nature Communications → Version 1 posted You are reading this latest preprint version Abstract Many antibiotics indirectly generate reactive oxygen species (ROS) that can damage bacterial genomes. Oxidised nucleobases become genotoxic when they are targeted for repair through excision, generating a single-strand discontinuity that can be converted to a double-strand break (DSB) by an oncoming replication fork. Because the genomic location of nucleobase oxidation is stochastic, investigating the fate of DNA replication machinery (replisome) at single-strand discontinuities has been limited. Here we have addressed this issue by expressing Cas9 nickases in Bacillus subtilis to create site specific single-strand discontinuities in a bacterial chromosome. We find that nicks in either leading or lagging strand arrest bacterial replication fork progression and generate a DSB that requires repair using homologous recombination to allow replication restart. These discoveries provoke reassessment of the fundamental mechanism of bacterial homologous recombination and provide insights to the development of alternative antimicrobials by identifying a specific pathway that can potentiate ROS-dependent bacterial killing. Biological sciences/Microbiology/Cellular microbiology Biological sciences/Molecular biology/DNA damage and repair Biological sciences/Genetics/Genomic instability Biological sciences/Molecular biology/DNA replication/Replisome Biological sciences/Biological techniques/Genomic analysis/Chromatin immunoprecipitation DNA replication nick helicase fork homologous repair recombination PriA restart Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Highlights Single-strand discontinuities inactivate the bacterial replisome Nicks on the leading or lagging strand template differentially affect fate of the helicase Double-strand breaks are repaired via homologous recombination and elicit the replication restart pathway Resection often generates a DNA strand with a 5′-end that must be processed to ensure productive helicase reloading MAIN Emergence of antibiotic resistant bacterial pathogens represents an urgent threat to human health and food production 1 . Understanding how bacteria respond to antibiotics is necessary to generate effective treatments. While antibiotics generally target specific essential bacterial systems (e.g. cell wall synthesis, translation, transcription) to inhibit bacterial proliferation, there is growing evidence that diverse antibiotics share the potential to disrupt bacterial physiology, thereby generating a second challenge to pathogens 2 . Work in several bacterial species indicates that a range of antibiotics ultimately generate reactive oxygen species (ROS) which can damage biomolecules including DNA 3 , 4 . Importantly, antibiotics that generate ROS elicit the bacterial DNA damage response, indicating the presence of double-strand breaks (DSB) under these conditions 4 – 7 . The potency of ROS-mediated DNA damage has been suggested to arise from conflicts between genome repair and replication systems 4 – 6 , 8 . Oxidised nucleobases located within the genome must be excised as part of the requisite repair pathway, leading to transient single-strand discontinuities in the DNA template 9 . If a bacterial replication fork arrives at a single-strand discontinuity, the encounter generates a DSB 10 . Elegant experiments using bacteriophage λ demonstrated that an engineered nick in either strand of the viral genome generated replication-dependent DSBs (Fig. 1 A) 11 . While active DNA replication is necessary to generate DSBs at a nick, the fate of the bacterial DNA replication machinery (replisome) at single-strand discontinuities has not been determined. The replisome is a multicomponent complex of proteins and enzymes that unwinds the chromosome and coordinates synthesis of new DNA polymers on the leading and lagging strands 12 , 13 . The most stable component of the replisome and most pivotal for its processivity is helicase 14 , 15 . Cellular replicative helicases are toroid enzymes that unwind DNA by encircling and translocating along a single client nucleic acid strand, acting as a wedge to disrupt base pairing 16 . Studies of bacterial replisomes in vitro and in vivo have revealed that the machinery is intrinsically dynamic, with most factors observed to bind and unbind interaction partners multiple times per replication cycle 17 , 18 . This behaviour enables the replisome to overcome a range of potential roadblocks and remain associated with the template, competent to continue DNA synthesis 19 . However, recent studies in eukaryotic systems have indicated that single-strand discontinuities generate DSBs and arrest DNA replication by impairing helicase progression 20 , 21 . Determining how bacteria react and respond when a replication fork encounters a single-strand discontinuity is crucial to understanding how ROS inhibits bacterial proliferation. Here we have investigated this question by expressing Cas9 nickases (nCas9) in the model bacterium Bacillus subtilis . The results show that a nick located in either template strand blocks downstream DNA synthesis, and that resumption of DNA replication is dependent upon a core set of homologous recombination proteins and the PriA-dependent replication restart system necessary to reload the replicative helicase. Unexpectedly, interrogation of the DNA repair pathway suggests that processing of the DSB often leaves the 5′-terminated DNA strand intact, which can hybridise to the homologous chromosome during recombination and perturb replication restart. We ascribe the enigmatic RecG helicase as the factor that remodels these recombination products containing a 5′-terminated DNA strand, thereby generating the replication fork structure recognised by PriA for productive helicase loading. These findings have critical implications for the fundamental mechanism of homologous recombination in bacteria and the aim of potentiating ROS-dependent antibiotic toxicity. RESULTS Single-strand discontinuities in a bacterial chromosome arrest DNA synthesis Oxidised nucleobases generated by ROS can be located randomly throughout a genome, making it a challenge to directly analyse conflicts with DNA replication during base excision repair. To mimic the activity of oxidised base repair, an experimental system capable of generating a site-specific single-strand discontinuity was developed in B. subtilis using Cas9 nuclease variants and single-guide RNAs (sgRNA) 20 – 23 . Cas9 carrying single amino acid substitutions (Cas9 D 10 A or Cas9 H840A ) created nickases (nCas9), while the double mutant (Cas9 D10A/H840A ) created a catalytically inactive enzyme (dCas9) 24 . Each cas9 allele was integrated into the B. subtilis chromosome under the control of a xylose-inducible promoter (Fig. 1 B) 25 . The location of sgRNA hybridisation is indicated by coordinates between the origin (0°) and terminus (~ 180°) on either the right (+) or left (-) replicore of the chromosome. To avoid complications arising from potential activation of prophage following DNA damage in the laboratory strain of B. subtilis , a derivative lacking these genetic elements (Δ6) was used throughout this study 26 . Cell viability was quantified using spot-titre assays, where strains are grown to saturation in liquid medium without xylose, before samples were serially diluted and plated onto solid medium with or without xylose to enumerate the number of colony forming units (CFUs). It was observed that the Cas9 D 10 A nickase elicited a stronger phenotype than Cas9 H840A upon induction with xylose (Fig. S1 A-B) 24 . To achieve comparable nicking efficiency at different sites in the chromosome, the cas9 D 10 A allele was used throughout the remainder of this study, in conjunction with distinct sgRNAs targeted to the template of either the leading or the lagging strand (in non-essential regions of the genome). Using sgRNAs to target leading and lagging strands at loci on both left and right chromosome arms, it was observed that expression of nCas9 resulted in severe growth inhibition (Figs. 1 C, S1C). In contrast, expression of dCas9 targeted to the same locations did not affect the number of CFUs (Figs. 1 C, S1D), indicating that DNA nicking was necessary for the observed growth defect. Next chromosome replication was investigated using whole genome marker frequency analysis (MFA). Expression of Cas9 variants was induced for one hour in exponentially growing cultures. Genomic DNA (gDNA) was harvested for next-generation sequencing, reads were mapped onto the reference genome sequence, and data is displayed with the chromosome origin ( oriC ) at the center of the graphs (Fig. S2 ). It was observed that nicks in either the leading or lagging strand resulted in a drop in sequencing coverage from the nick site to the terminus region, whereas expression of dCas9 with the same sgRNAs did not appear to inhibit DNA replication (Fig. 1 D). Because cells expressing nCas9 are unable to synthetise DNA downstream of the nick, this likely explains the strong growth inhibition observed in spot-titre assays (Fig. 1 C). Previous results indicated that a replication fork encountering a single-strand discontinuity generates a DSB 11 . To determine whether DSBs were being formed in B. subtilis following nCas9 induction, localisation of the reporter mNeonGreen-RecA (mNG-RecA) was observed in live cells using fluorescence microscopy. It has been shown that following artificial endonuclease cleavage to create a DSB, fluorescently labelled RecA recombinase assembles into bundles 27 – 29 . Prior to induction of nCas9, mNG-RecA is observed evenly distributed over the nucleoid (Fig. S3 A). Following expression of nCas9 for one hour during exponential growth, mNG-RecA bundles were detected in 58.5% of cells when the leading strand template was nicked and 48% of cells when the lagging strand template was nicked (Figs. 1 E-F, S3B). In contrast, expression of dCas9 did not significantly promote mNG-RecA bundling (Figs. 1 E-F, S3B). These results are consistent with the model that collision of a replication fork with a nick in either strand of the parental DNA duplex generates a DSB. Single-strand discontinuities in a bacterial chromosome inactivate the replisome MFA indicated that under conditions expressing nCas9, DNA synthesis does not proceed beyond the nick. We wondered whether the replisome becomes arrested at a single-strand discontinuity, or rather whether the replication fork collapses. If the latter event occurs, then the bacterial replication restart system would be required to reload helicase and promote replisome assembly 30 , 31 . In B. subtilis there is only one known restart system composed of the essential proteins PriA and the helicase loaders DnaD, DnaB, and DnaI (note that in B. subtilis the replicative helicase is DnaC) 32 – 36 . Chromatin immunoprecipitation (ChIP) followed by quantitative PCR (qPCR) was used to determine the enrichment of helicase and replication restart proteins surrounding a nick in either the leading or lagging strand template. Following induction of nCas9 for one hour in exponentially growing cells, samples were harvested for ChIP. Along with sites flanking the nick region, protein enrichment was also probed at the origin oriC as positive control and normalised to a site located on the opposite chromosome arm equidistant from oriC (+ 90, Fig. 2 A). ChIP showed that helicase is specifically enriched upstream of nicks located in either template strand (Figs. 2 B, S4). This upstream helicase enrichment was not detected when dCas9 was expressed, indicating that the Cas9-sgRNA nucleoprotein complex alone does not stall replication fork progression (Figs. 2 B, S4). Interestingly, significant enrichment of helicase was observed downstream of the nick site only when cleavage occurred in the template used for leading strand synthesis (Figs. 2 B, S4). Because the bacterial replicative helicase translocates along a single strand in the 5′◊3′ direction, ChIP results suggest that when the template for the lagging strand is nicked helicase runs-off the DNA substrate, whereas when the template for the leading strand is nicked helicase progresses downstream of the single-strand discontinuity (Fig. 2 G). Consistent with the ChIP enrichment observed in vivo (Fig. 2 A-B) and replisome inactivation (Fig. 1 D), reconstituted unwinding assays in vitro using bacterial helicase on model replication fork substrates have shown that if a flap is absent from the substrate (i.e. mimicking a nick), helicase transitions from encircling a single DNA strand onto double-stranded DNA (dsDNA) and slides along the duplex, no longer able to unwind 37 . The enrichment of helicase upstream of nick sites is consistent with either helicase stalling or helicase reloading via the replication restart pathway. To distinguish between these models, ChIP was used to probe for the presence of replication restart proteins surrounding the nicks. Following nCas9 induction, enrichment of PriA, DnaD, DnaB and DnaI was detected upstream (but not downstream) of the sgRNA target sequences, while these proteins were absent from this region when dCas9 was expressed (Fig. 2 C-F). These results suggest that nicks in either the leading or lagging strand template elicit the replication restart pathway for helicase reloading, whether helicase falls off the template (lagging strand template nick) or translocates downstream (leading strand template nick) in an inactive state (Fig. 2 G). A weakened nCas9 system permits DNA replication restart While replication restart proteins were enriched upstream of nick sites (Fig. 2 B-F), viability assays and MFA both indicated that downstream DNA synthesis was inhibited (Fig. 1 C-D). We speculated that persistent nCas9 cleavage activity competes with productive DNA replication restart. Therefore, to allow interrogation of processes occurring downstream of DNA cleavage and replisome collapse, a weaker nCas9 system was developed. An ssrA degradation tag 38 was fused to cas9 D 10 A to promote nCas9 proteolysis, and expression of both nCas9-ssrA and sgRNAs were placed under the control of xylose-inducible promoters (Fig. S5A). In this background, induction of nCas9 did not significantly inhibit cell viability or chromosome content (Fig. S5B-C) and MFA showed that DNA synthesis continued after a one-hour xylose induction, consistent with low tolerable levels of nickase activity (Fig. 2 H). This weakened nicking system was employed for all experiments described below. PriA is required to restart DNA replication at single-strand DNA discontinuities The master replication restart factor priA is essential for growth of B. subtilis. To ascertain whether PriA-dependent replication restart is required following chromosome nicking, a titratable PriA complementation system was developed (Fig. S6A). Here conditions were established to express PriA-ssrA at a low level, sufficient to maintain B. subtilis viability while limiting the amount of PriA activity within a cell (Fig. S6B-C). When nCas9-ssrA and sgRNA were induced with limiting PriA-ssrA, spot-titre analyses showed that cell growth was inhibited in the absence of the endogenous priA copy (Fig. 2 I-J). Yet in this background, higher expression of PriA-ssrA rescued viability (Fig. S6D). Following induction of nCas9 and sgRNA for one hour during exponential growth, MFA showed that limited levels of PriA-ssrA could not sustain chromosome synthesis downstream of the nick site (Fig. 2 K). Therefore, these data indicate that PriA is essential to restart DNA replication following replisome inactivation at single-strand DNA discontinuities. The DSB resulting from a single-strand discontinuity is repaired using a core set of homologous recombination proteins Cytological analysis indicated that a nick in the template of either the leading or lagging strand generated a DSB (Fig. 1 E). Strong evidence indicates that in bacteria such DSBs are repaired by homologous recombination, thereby recreating a replication fork to allow resumption of DNA synthesis 39 . To determine the factors required to repair DSBs following nCas9 nicking, a targeted reverse genetic analysis was employed, focused on genes previously implicated in homologous recombination and other DNA repair processes (Table S1 ). Using this approach, spot-titre analyses revealed that eight non-essential genes were required for normal growth upon nick induction: DNA end-processing genes addA / addB and homologous recombination genes recF / recO / recR / recA / recG / recU (Fig. 3 A and S7A). Functional complementation of each synthetic lethal deletion confirmed that these gene products are necessary to support growth in cells expressing nCas9 (Fig. S7B). Surprisingly, the screen revealed that several factors implicated in DNA repair are not essential under these conditions, such as those classed for end-processing (RecQ, RecS, RecJ) 7 , 40 , maintaining chromosome structure (“SMC-like” RecN, SbcCD, SbcEF) 41 , 42 , recombination mediation (RecD2, RecX) 43 – 45 and Holliday junction migration (RuvA, RuvB) 40 (Fig. S8). Focusing on the eight mutants that produced a clear growth defect, MFA was used to analyse DNA replication following induction of nCas9 and sgRNA in exponentially growing cells. The results showed that DNA end-processing ( addA / addB ) and recombination mutants ( recF / recO / recR / recA / recG ) are defective in chromosome replication downstream of the nick site, with the only exception being recU (Fig. 3 B; note subtle differences in replication profiles of parental strains containing deletions of homologous recombination genes, consistent with previous findings 46 , 47 ). These observations support the hypothesis that homologous recombination is necessary to repair the DSB and restart DNA replication. Based on the proposed activities of these proteins, the following model was developed (Fig. 3 C). AddAB helicase/nuclease processes the DSB to generate a single-stranded substrate with a 3′-end required for recombination 48 , 49 ; RecFOR displaces the single-stranded binding protein (SSB) from single-stranded DNA substrates produced by AddAB, thereby allowing RecA to bind 50 , 51 ; RecA is the recombinase that binds to single-stranded DNA (ssDNA) with a free 3′-end, identifies homologous DNA on the non-broken chromosome, and performs strand invasion and exchange 52 , 53 ; RecU resolves the Holliday junction generated by RecA 54 . The role of RecG in vivo is enigmatic 55 , 56 and discussed further below. The SSB C-terminal tail is required for DNA repair following replisome inactivation The requirement of the RecFOR system for repair of a seDSB strongly suggested that the essential single-strand binding protein (SSB) was also involved. To interrogate this hypothesis, recruitment of SSB was probed using ChIP following induction of nCas9 and sgRNA in exponentially growing cells. The results showed that SSB is enriched upstream of either leading or lagging strand nicks (Fig. 4 A). SSB enrichment ~ 3 kb upstream from the nick site was dependent upon AddAB but independent of the remaining factors required for homologous recombination (Fig. 4 B), indicating that AddAB is necessary to generate ssDNA for SSB. SSB forms a homotetramer that binds ssDNA non-specifically via its N-terminal domain. Each SSB monomer harbours a long disordered C-terminal domain that contains a protein interaction sequence at the terminus (the C-terminal tail or CTT) (Fig. S9A-B) 57 . While the B. subtilis SSB-CTT is dispensable for cell viability, it is required for efficient repair of DNA lesions created by either UV irradiation or Mitomycin C treatment 42 , 58 . To determine whether the SSB-CTT is required for repair of a DSB following replisome inactivation at a nick, an IPTG-inducible SSB complementation system was generated in a Δ ssbA strain, allowing expression of either the wild-type protein or a variant lacking the last six amino acids (SSB ΔCTT , Fig. S9C). Spot-titre analyses confirmed that SSB ΔCTT can complement a Δ ssbA mutant similar to wild-type SSB and that viability was dependent upon IPTG (Fig. S9D), and immunoblots showed that the inducible SSB proteins were being expressed at levels comparable to the endogenous protein (Fig. S9E). The SSB complementation system was then moved into strains expressing nCas9 and sgRNA (Fig. S9F). Spot titre analyses showed that wild-type SSB was able to support viability following induction of nCas9 and sgRNA, whereas the SSB ΔCTT variant could not (Fig. 4 C). MFA confirmed that chromosome replication from the nick site to the terminus is arrested when SSB lacks its C-terminal tail (Fig. 4 D). Therefore, the results indicate that the SSB-CTT is necessary for DNA repair following replisome inactivation at a nick. The SSB-CTT is required to recruit RecO to the site of DNA repair during homologous recombination The SSB-CTT acts as a protein interaction hub 42 that directly interacts with several factors found to be necessary for repair of a DSB, including RecO, RecG, and PriA (Figs. S9G, 2I-K, 3A-B). To investigate the essential role of the SSB-CTT, ChIP was employed to probe the recruitment of SSB variants, His-RecO, and His-RecA upstream of nick sites. ChIP showed that the SSB ΔCTT variant remained enriched upstream of nicks at similar levels to the wild-type protein (Fig. 4 E). However, enrichment in His-RecO and His-RecA was lost in cells harbouring SSB ΔCTT (Fig. 4 E), indicating that the SSB:RecO interaction depends on the SSB-CTT (Fig. S9H). In cells encoding wild-type SSB, enrichment of His-RecA was dependent upon both AddAB and RecFOR (Fig. 4 F), consistent with the model that RecFOR is necessary for RecA loading onto ssDNA substrates bound by SSB. Taken together, these results indicate that an essential role of the SSB-CTT is to recruit RecO to repair sites following replisome inactivation at single-strand DNA discontinuities. What is the essential role of RecG for restart of DNA replication? While the role of most factors required for resuming DNA replication following replisome inactivation at a single-strand discontinuity could be rationalised, the role of RecG was unclear. RecG has been implicated in a myriad of DNA transactions including reversal of replication forks, migration of Holliday junctions, regulation of RecA strand exchange, processing of flaps generated when DNA replication forks converge, and stabilisation of D-loops 56 , 59 – 62 . Examination of the MFA in a Δ recG mutant showed that there was an increase in sequence read depth upstream of a nick targeted to either the leading or lagging strand template (Fig. 5 A), suggesting that DNA synthesis increases in this region in the absence of RecG. This observation was reminiscent of results reported in Escherichia coli during DSB repair in a Δ recG mutant, where it was proposed that RecG remodels intermediate DNA structures formed during homologous recombination to ensure PriA is correctly orientated to load helicase (Fig. 5 B, in the absence of RecG helicase reloading occurs on the opposite strand normally used for DNA replication, thus directing inappropriate DNA synthesis from the terminus towards oriC ) 47 . This hypothesis diverges from the generally accepted model for homologous recombination, in which resection of a DSB involves degradation of the strand with a 5′-end, thereby generating a single-stranded 3′-end substrate for recombinases (e.g. RecA, Fig. S10). The revised model instead suggests that following DNA end-processing, the strand containing a 5′-end remains intact and can hybridise the displaced strand (D-loop) formed as a consequence of RecA strand invasion 47 . It was suggested that this intermediate DNA structure was the substrate for RecG, such that the enzyme would remodel the strands by unwinding the annealed 5′-end until the annealed 3′-end abutted the fork junction, thus generating the correct substrate for PriA to reload helicase (Fig. S11) 63 . Biochemical data using purified proteins has shown that RecG and PriA can perform these activities on model replication fork substrates 63 , 64 . Therefore, we sought to test this hypothesis in vivo using the nCas9 system. RecG is required to remodel replication forks formed when a DSB contains a 5′-end The genetic analysis above indicated that the AddAB helicase/nuclease was responsible of end processing of the DSB (Fig. 3 A). While both AddA and AddB subunits harbour nuclease activity, the two enzymes cleave distinct strands of the DNA duplex following unwinding by AddA helicase activity (AddA cleaves the strand terminating with a 3′-end, AddB cleaves the strand terminating with a 5′-end) 49 . Therefore, it was hypothesised that inactivating either AddA or AddB nuclease activity would bias DSB processing to favour the generation of either a 3′-tail (AddA nuclease mutant) or a 5′-tail (AddB nuclease mutant). To begin, ChIP was used to confirm that AddAB was acting directly on the DSB generated following replisome inactivation at a nick. AddA and AddB were independently fused to the fluorescent protein mNeonGreen and enrichment of each subunit was determined. The results showed that mNG-AddA and mNG-AddB are both enriched upstream of nicks, consistent with their direct role in end-processing under these conditions (Fig. 5 C). Next, the enzymatic activities of AddA and AddB were reduced through substitution of catalytic residues 49 . Spot titre analyses showed that the helicase defective AddA K 36 A (AddA ΔHEL ) was unable to sustain bacterial growth upon induction of nicks, whereas nuclease defective variants of either AddA D1172A (AddA ΔNUC ) or AddB D961A (AddB ΔNUC ) grew similar to a strain with wild-type AddAB (Figs. 5 D, S12A). Notably, the double nuclease variant AddA ΔNUC AddB ΔNUC also supported normal growth (Fig. 5 D), suggesting that under these conditions in live cells the DSB may not require resection. MFA confirmed that DNA synthesis was blocked downstream of a nick in the strain expressing AddA ΔHEL , while the strains expressing AddA ΔNUC , AddB ΔNUC or AddA ΔNUC AddB ΔNUC were comparable to wild-type (Figs. 5 E, S12B-C). Moreover, ChIP demonstrated that SSB is not enriched upstream of nicks in a strain expressing AddA ΔHEL , whereas SSB was enriched in AddA ΔNUC and AddB ΔNUC backgrounds (Fig. 5 F). Together these results indicate that AddAB helicase activity is directly required to process a DSB generated after replisome inactivation at a nick, and that nuclease activity can be abolished without compromising DNA replication restart. Finally, the Δ recG mutant was moved into strains expressing either AddA ΔNUC or AddB ΔNUC and nicking was induced (Fig. 6 A-B). Spot-titre assays showed that RecG remained essential in the strains containing AddB ΔNUC , where degradation of the strand terminating with a 5′-end is inhibited (Fig. 6 C). Strikingly, in the AddA ΔNUC strain where degradation of the strand terminating with a 3′-end is inhibited, the absence of RecG was well tolerated (Fig. 6 C-D). These results are consistent with the model that RecG is required to remodel recombination intermediates that contain a single-strand with a 5′-end annealed to a D-loop. Importantly, the critical role of RecG in strains with single-strand discontinuities (Fig. 3 A) implies that wild-type processing of a DSB often generates a single-strand with a 5′-end, which is inconsistent with most current models of bacterial homologous recombination. DISCUSSION In this study we have characterised the fate of the bacterial replisome in vivo after it encounters a single-strand discontinuity in the DNA template (Fig. 1 A). The results indicate that a nick in the template for either the leading or lagging strand arrests DNA synthesis, generates a DSB, and elicits replication restart following repair of the DNA break through homologous recombination (Figs. 1 – 3 ). These findings are consistent with the observation that a nick in a replicating bacteriophage chromosome generates DSBs 11 and with recent reports of the impact of nicks in eukaryotic replication systems 20 , 21 , 23 . Importantly, while replisomes are thought to be generally intolerant to many types of DNA damage 65 , it appears that the bacterial replication machinery cannot readily bypass single-strand interruptions as these lesions affect helicase progression (Figs. 2 , 7 (I)). Repair of the DSB to facilitate replication restart requires homologous recombination (Fig. 3 ). Genetic analysis combined with ChIP indicates that a single pathway is utilised under these experimental conditions (Fig. 7 ): AddAB unwinds the DNA; SSB binds the ssDNA; RecFOR counteracts SSB and allows RecA to bind ssDNA; RecA promotes strand invasion of the homologous chromosome and generates a D-loop, which often anneals to a single-strand with a 5′-end; RecG regresses the fork and simultaneously separates the annealed strand with a 5′-end; PriA binds to the remodelled fork where it interacts with the strand harbouring a 3′-end and reloads helicase to promote replisome assembly and DNA replication restart. The observed lack of redundancy between proteins involved in recombinational DNA repair suggests that the pathway identified here is the major route for fixing this type of collapsed replication fork in B. subtilis 41 , 66 . Building upon these results, we investigated specific protein activities to help elucidate their roles during recombinational repair of a DSB. Below we discuss these findings in what we propose is chronological order. The replicative helicase is inactivated after it encounters a nick In bacteria the replicative helicase encircles and tracks along the template for lagging strand synthesis. ChIP data suggests that there are distinct fates of the helicase depending on the location of a nick (Figs. 2 A-B, S4). As expected for a nick in the lagging strand template, helicase enrichment was only observed at the cleavage site and upstream, consistent with the model that helicase runs-off the template when it encounters the single-strand discontinuity (Figs. 2 G, 7 (I)). In contrast, when the nick was in the leading strand template, helicase was also enriched downstream of the cleavage site, suggesting that the enzyme continued translocation. We propose that upon encountering a nick in the leading strand template, helicase slides onto the dsDNA template in an inactive conformation that cannot perform further DNA unwinding (Figs. 2 G, 7 (I)). This model is consistent with in vitro experiments using Thermus aquaticus helicase, where after unwinding up to a nick on the leading strand template, the enzyme can continue to translocate with the two strands of DNA passing through the central channel with no resultant unwinding 37 . Whether the bacterial helicase is actively removed from dsDNA under these conditions, akin to what has been observed for the eukayotic CMG helicase in Xenopus extracts 20 , requires further investigation. AddAB helicase activity is necessary and sufficient for recombinational repair AddAB and related enzymes have been shown to bind blunt end dsDNA or dsDNA with short overhangs (4–20 nucleotides with either a 5′- or a 3′-end) 49 , 67 . When the template for leading strand synthesis is nicked, the DNA polymerase likely runs-off the end of the substrate to produce a blunt or nearly blunt end, suitable for recognition by AddAB. However, when the template for lagging strand synthesis is nicked, a substrate with a 3′-tail would likely be produced. Because the reverse genetic screen suggested that no additional nucleases are required for this recombinational repair pathway (Figs. 3 , S8), we speculate that the high local concentration of replication factors would facilitate priming near the end of the 3′-tail, thereby filling the ssDNA with an Okazaki fragment to create a suitable substrate for AddAB. The master recombination factor RecA requires ssDNA to promote homologous pairing. A long-standing question is how this ssDNA is generated in vivo . Most current models suggest that enzymes resect a DSB to generate a single-strand terminating with a 3′-end 7 , 31 . Here we tested this model by mutating the nuclease activities of AddA and AddB. The results showed that neither nuclease activity is required for enzyme function under the experimental conditions tested (Figs. 5 D-E, S12B-C). In contrast, AddAB helicase activity was required for recombinational repair, consistent with a previous study of the ancestral AdnAB resection machinery in Mycobacteria tuberculosis 68 . These results suggest that the key role of AddAB is not to digest the dsDNA ends, but rather to separate the strands and create the ssDNA substrate for SSB (Figs. 5 F, 7 (II-III)). The local concentration of SSB is high at the site of DNA replication 69 , 70 , potentially facilitating its rapid association with the ssDNA emerging from AddAB. In fact, in vitro the AddA NUC AddB NUC nuclease-dead enzyme does not unwind dsDNA in the absence of SSB 71 , suggesting that AddAB unwinding and SSB binding ssDNA may be coupled. We propose that AddAB nuclease activity represents a secondary method of generating a ssDNA substrate for SSB, following initial AddAB helicase activity required to separate the DNA duplex. Interaction of the SSB C-terminal tail with RecO is necessary for replication repair The SSB-CTT is an established protein interaction hub that has been shown to directly bind several of the essential replication and repair factors required at nicks, including RecO 72 – 75 , RecG 76 , 77 , and PriA 58 , 78 . However, the critical activities performed by the SSB-CTT in vivo have not yet been established 42 . Based on the results above, we conclude that the SSB-CTT interaction with RecO is necessary for replication repair. RecO is part of the RecFOR system required for RecA loading onto ssDNA in vitro 79 and RecA-GFP foci formation in vivo 80 . Because RecO activity occurs before the activities of RecG and PriA (i.e. RecA binding ssDNA is blocked, Fig. 4 F), this precludes further conclusions regarding the importance of these protein:protein interactions at this time. In vitro the RecO:RecR complex facilitates loading of RecA onto SSB-coated ssDNA 81 , 82 . Because RecO, RecR, and RecF were all required for replication repair, we propose that the essential role of RecF here is to recruit RecR and generate the RecO:RecR complex. This model is consistent with a hand-off mechanism for RecR observed in vitro 81 . RecF is a dimeric ATPase that adopts a clamp conformation around dsDNA 51 . Recent studies in E. coli have shown that recruitment of RecF to sites of repair is mediated by the DNA polymerase sliding clamp processivity factor DnaN 82 . As was the case for SSB 70 , the local concentration of DnaN is high near the sites of DNA replication collapse 69 , thereby potentially facilitating RecF recruitment. Importantly for this model, since both SSB and DnaN are enriched at a replication fork, we predict that there is a mechanism to regulate RecO:RecR complex formation until it is required, thus safeguarding the SSB-coated strand from promiscuous RecA loading (Figs. 4 B, 4 F, 7 (III-V)). RecG remodels a repaired replication fork by unwinding dsDNA with an exposed 5′-end The biological role of RecG has been a long-standing question 55 , 56 . In vitro RecG is a DNA translocase that unwinds a myriad of DNA substrates, including Holliday junctions, D-loops, and model replication forks 47 , 60 – 62 , 64 . Here MFA indicates that a ΔrecG mutant cannot efficiently restart DNA replication at a nick directed to either the leading or the lagging strand template (Figs. 3 B, 5 A-B), consistent with a role in fork remodelling 46 . Based on genetic experiments and supported by biochemical assays in vitro , a model has been presented for RecG binding to a replication fork and unwinding dsDNA on the lagging strand template to allow correct orientation of PriA for helicase loading 47 , 63 . Critically, this model requires that the single strand with a 5′-end from the DSB can hybridise with ssDNA of the D-loop on the homologous chromosome. How this predicted intermediate structure is constructed remains to be determined, but the implication is that the separated strands of the DSB are either the same length or the strand with a 5′-end is longer (Figs. 6 A, 6 D, 7 (VI-VII)). This model contrasts with most current views of bacterial DSB end-processing where a single strand with a 3′-end is produced for RecA (Fig. S10) 7 , 31 , 83 . Moreover, because RecG is essential in strains expressing Cas9 nickases, it suggests that during wild-type DSB end-processing the separated strands often contain a 5′-end that requires remodelling by RecG (Fig. 7 (VII)). These unanticipated results, together with those of the accompanying manuscript analysing resolution of collapsed eukaryotic replication forks 84 , necessitate further evaluation of DSB management following replisome collapse across all domains of life. Holliday junction migration and resolution are not necessary for replication restart Branch migration of a Holliday junction is generally considered a hallmark of homologous recombination, therefore it was notable that the canonical Holliday junction translocase RuvAB was not required for replication repair at nicks (Fig. S8D). In fact, the ΔrecU ΔruvAB knockout attenuated the penetrance of the ΔrecU mutant upon introduction of single-strand discontinuities, suggesting that either another enzyme can functionally complement RuvAB activities or that branch migration is not required for this repair pathway. While genetic analysis shows that RecU is required for wild-type growth following expression of Cas9 nickases, MFA indicates that RecU is not necessary for replication restart per se (Fig. 3 ). These results are consistent with the model that RecU resolves the Holliday junction formed during recombinational repair 85 . We hypothesise that the growth defect observed in the ΔrecU mutant may be caused by aberrant chromosome segregation 85 , as the recombined DNA molecules need to be separated prior to completing cell division. It is known that multiple systems can facilitate chromosome separation following replication (e.g. Topoisomerase IV, XerD) 86 , 87 , potentially explaining why the growth defect of the ΔrecU mutant under these experimental conditions was less severe compared to the other recombination mutants identified. Potentiating ROS-mediated antibiotic toxicity by perturbing DNA repair In this work, the nicks generated by Cas9 enzymes were used as proxies for single-strand discontinuities formed during removal of damaged DNA from a bacterial genome. We find that when a bacterial replication fork encounters a single strand discontinuity it becomes inactivated, likely contributing to the toxic effect of ROS-dependent damage induced by antimicrobials. By identifying the essential factors and activities required to repair the DSB formed after a replisome encounters a nick, we have provided a set of molecular targets whose disruption could potentiate the activity of antibiotics that produce ROS-dependent DNA damage. Methods Experimental model and growth The following bacterial organisms were used: E. coli and B. subtilis. Unless otherwise stated in method details, strains were grown at 37°C using nutrient agar (NA) or lysogeny broth (LB: 10 g/L tryptone, 10 g/L NaCl, 5 g/L yeast extract) for routine selection and maintenance of bacterial strains. Supplements were added as required: ampicillin (100 µg/ml), erythromycin (1 µg/ml) in conjunction with lincomycin (25 μg/ml), chloramphenicol (5 µg/ml), kanamycin (5 µg/ml), spectinomycin (50 µg/ml), zeocin (10 μg/ml), xylose (1% w/v) and IPTG (0.05 mM, 0.1 mM or 1 mM). Bacterial strains Bacterial strains used in this study are listed in Table S2. E. coli plasmid construction E. coli transformations for constructs harbouring priA were performed in CW198 via heat-shock following the Hanahan method 88 and propagated in LB with appropriate antibiotics at 37°C. DH5α [ F- Φ80lacZΔM15 Δ(lacZYA-argF) U169 recA1 endA1 hsdR17(rk-, mk+) phoA supE44 thi-1 gyrA96 relA1 λ- ] was used for other plasmids construction. Plasmids were purified using QIAprep spin miniprep kits and recombinant DNA confirmed by sequencing (Table S3). Descriptions, where necessary, are provided below . pCW367, pCW368, pCW478, pCW479, pCW510, pCW717, pCW734, pCW736 and pCW850 were generated by Quickchange mutagenesis using mutagenic primer pairs containing an overlap of 10-15 bp. PCR products were treated with DpnI for at least 3 hours to digest template DNA and 10 μl were used for E. coli transformation. pCW300, pCW301, pCW327, pCW460, pCW471, pCW472, pCW575, pCW734, pCW736 and pCW766 were generated by Gibson assembly using the NEB Hi-Fi cloning kit according to manufacturer instructions following purification of individual PCR fragments using QIAquick PCR purification kits. Linear recombinant DNA assembly For recombinant DNA assembly without E. coli propagation, pCW573, pCW574, pCW577, pCW578, pCW605, pCW640, pCW672, pCW702, pCW720, pCW732, pCW758, pCW759, pCW760, pCW777, pCW779, pCW781, pCW783, pCW785, pCW802, pCW803, pCW804, pCW809, pCW834, pCW848, pCW854, pCW855, pCW857 and pCW859 were assembled linearly using a multistep in vitro assembly process followed by nested PCR amplification prior to B. subtilis transformation. Briefly, individual PCR fragments corresponding to homology regions and insert DNA were amplified, purified and DNA ends ligated using the NEB Hi-Fi cloning kit. Raw Gibson assembly products were then amplified using nested primer pairs annealing towards the end of homology regions and PCR products were used for B. subtilis transformation. Following isolation of B. subtilis colonies resistant to the appropriate antibiotic markers, genomic DNA was extracted using the DNeasy blood and tissue kit, regions corresponding to recombinant DNA were amplified via PCR using oligonucleotides annealing outside homology fragments to ensure double crossover had occurred and results were confirmed via PCR product sequencing. Note that CW2045 and derivatives harbour the additional addA D682E mutation necessary to propagate addB D961A -addA D1172A in live cells, which is surface exposed and structurally unlikely to impact AddAB end-processing activities (Fig. S12E). B. subtilis strain construction Transformation of competent B. subtilis cells was performed using an optimized two-step starvation procedure as previously described 89 . Briefly, recipient strains were grown overnight at 37°C in transformation medium (Spizizen salts: 0.2% w/v ammonium sulphate, 1.4% w/v dipotassium phosphate, 0.6% w/v potassium phosphate, 0.1% w/v tri-sodium citrate, 0.02% w/v magnesium sulphate supplemented with 1 μg/ml ammonium iron(III) citrate, 6 mM MgSO 4 , 0.5% w/v glucose, 0.02 mg/ml tryptophan and 0.02% w/v casein peptone) supplemented with IPTG where required. Overnight cultures were diluted 1:20 into fresh transformation medium supplemented with IPTG where required and grown at 37°C for 2.5-3 hours with continual shaking. An equal volume of prewarmed starvation medium (Spizizen salts supplemented with 6 mM MgSO 4 and 0.5% w/v glucose) was added and the culture was incubated at 37°C for two hours with continual shaking. DNA was added to 350 μl cells and the mixture was incubated at 37°C for one hour with continual shaking. 20-200 μl of each transformation was plated onto selective media supplemented with IPTG where required and incubated at 37°C for 24-48 hours. Results were validated by propagating cells on relevant antibiotic markers and sequencing. Spot titre assays Cells were grown in LB overnight in the presence of 0.5% w/v glucose. Overnight cultures were diluted in fresh LB medium using 10-fold serial dilutions and 5 μL aliquots were spotted onto NA plates with or without xylose. NA plates were supplemented with IPTG when required as indicated on individual figure panels. All plates were incubated for 24 hours or up to 48 hours when indicated. Experiments were performed independently at least three times and representative data are shown. Immunoblot analyses Cultures were grown overnight in LB and diluted 1:100 in fresh medium the next morning. Diluted cells were grown to an absorbance at 600 nm (A 600 ) of 0.2-0.3, induced with 1% w/v xylose or 1 mM IPTG where required and further incubated for one hour. Cells were harvested using centrifugation, resuspended in 1x PBS supplemented with one tenth of a peptidase inhibitor tablet and sonicated (40 amp) twice for 12 seconds with 3 second pulses at 4°C. Samples were adjusted to 1X LDS buffer and 125 mM DTT, fixed for 5 minutes at 95°C and centrifuged for 2 minutes prior to loading the supernatant on polyacrylamide gels. Proteins were separated by electrophoresis using NuPAGE 4-12% Bis-Tris gradient gels run in MES SDS buffer and transferred to methanol-activated Hybond-P PVDF membranes using Wypall X60 cloths for blotting in a semi-dry apparatus (Bio-rad Trans-Blot Turbo, transfer buffer: 300 mM Tris-HCl, 300 mM glycine, 140 mM tricine, 0.05% w/v SDS, 2.5 mM EDTA). Membranes were washed twice using PBST (1X PBS supplemented with 0.1% v/v Tween-20) prior to blocking for 90 minutes at room temperature (blocking buffer: PBST supplemented with 7.5% w/v milk). Individual polypeptides were probed using the following primary antibodies diluted in blocking buffer following overnight incubation at 4°C: anti-Cas9 (1:1000), anti-FtsZ (1:5000), anti-PriA (1:1000), anti-SSB (1:1000) and anti-mNeonGreen (1:1000). Detection was performed with anti-rabbit (PriA, SSB), anti-sheep (FtsZ) or anti-mouse (Cas9, mNeonGreen) horseradish peroxidase-linked secondary antibodies in combination with the Clarity ECL substrate using an ImageQuant LAS 4000 mini digital imaging system. Detection of Cas9, FtsZ, PriA, SSB and mNeonGreen was within a linear range. Experiments were independently performed at least twice and representative data are shown. Whole genome sequencing Cells were grown in rich chemically defined medium (RCDM: Spizizen salts supplemented with 1 μg/ml ammonium iron(III) citrate, 0.1 mM CaCl 2 , 0.13 mM MnSO 4 , 6 mM MgSO 4 , 0.5% w/v glucose, 0.1% w/v glutamate, 0.02 mg/ml tryptophan and 0.02% w/v casein peptone) overnight and diluted 1:100 in fresh RCDM the next morning. Diluted cells were grown to an absorbance at 600 nm (A 600 ) of 0.2-0.3, induced with 1% w/v xylose and further incubated for one hour. Growth was arrested by adding 0.05% w/v sodium azide to cultures followed by vigorous mixing, cells were collected by centrifugation, the supernatant discarded, and pellets were flash frozen in liquid nitrogen before genomic DNA extraction using the DNeasy blood and tissue kit and a final elution volume of 50 µl in ultrapure water. Individual gDNA libraries were prepared using the DNA Prep tagmentation kit and pre-paired i5/i7 indices prior to pooling and DNA quantification with the 1X dsDNA high sensitivity kit using a Qubit instrument. Pooled libraries were diluted to 100 pM in resuspension buffer and 20 µl were loaded onto iSeq 100 i1 Reagent v2 sequencing chips for whole-genome sequencing using an iSeq100 sequencer. Following whole-genome sequencing and demultiplexing embedded within the iSeq100 indexed sequencing workflow, .FASTQ files were extracted and loaded onto the CLC Genomics Workbench for initial data processing. Paired .FASTQ files were mapped onto the B. subtilis Δ6 reference genome using the ‘Map Reads to Reference’ function and a .BAM file was created from mapped sequencing reads for each sequenced genome. An in-house script written in R using the Rsamtools library was employed to deconvolute .BAM files to validate even read distribution for sense/antisense strands prior to calculating density coverage and extracting cumulated densities per chromosome position. Fluorescence microscopy Strains were grown overnight at 37°C in RCDM and the following day, cultures were diluted 1:100 into fresh imaging medium (RCDM where glucose is replaced by 0.5% v/v glycerol) in the presence of 0.1 mM IPTG for induction of mNeonGreen-RecA expression where required. Cells were allowed to grow to an A 600 of 0.3, induced with 1% w/v xylose and further incubated for 90 minutes. DAPI (5 µg/ml) and Nile red (1 µg/ml) stains were used to visualise the nucleoid and cell membrane following a 10 minute incubation on a benchtop incubator prior to imaging. Cells were mounted on ~1.25% agar pads (in sterile ultrapure water) and a 0.13- to 0.17-mm glass coverslip was placed on top. Microscopy was performed on an inverted epifluorescence microscope (Nikon Ti) fitted with a Plan Apochromat Objective 100x/1.40 NA Oil Ph3. Light was transmitted from a CoolLED pE-300 white light source through a Sutter Instruments liquid light guide and images were collected using a Photometrics Prime camera. Chroma fluorescence filter sets were used with 100 ms GFP, 250 ms DAPI and 50 ms mCherry exposure times at 100% LED power. Digital images were acquired using NIS Elements and analysed via the Fiji software 90 . Quantification of mNeonGreen-RecA features was manually curated from the count of 100 individual cells. All experiments were independently performed at least twice and representative data are shown. Chromatin immunoprecipitation Chromatin immunoprecipitation (ChIP) was performed as previously described with minor modifications 32 . Strains were grown overnight at 37°C in RCDM and the following day, cultures were diluted 1:100 into fresh medium and allowed to grow to an A 600 of 0.2-0.3. Cells were induced with 1% w/v xylose and further incubated for one hour prior to ChIP. Samples were adjusted to 1X PBS and crosslinked with 1% v/v formaldehyde for 10 minutes on a roller at room temperature, then quenched with 0.1 M glycine. Cells were pelleted at 4°C, washed three times with ice-cold 1X PBS (pH 7.3), frozen in liquid nitrogen and stored at -80°C. Frozen cell pellets were resuspended in 500 µl of lysis buffer (50 mM NaCl, 10 mM Tris-HCl pH 8.0, 20% w/v sucrose, 10 mM EDTA, 100 µg/ml RNase A, one quarter of a peptidase inhibitor tablet, 4 mg/ml lysozyme) and incubated at 37°C for 30 min to degrade the cell wall. Protoplasts were supplemented with 500 µl of immunoprecipitation buffer (300 mM NaCl, 100 mM Tris-HCl pH 7.0, 2% v/v Triton X-100, one quarter of a peptidase inhibitor tablet) to lyse cells and the mixture was incubated at 37°C for a further 10 minutes before cooling on ice for 5 minutes. Lysis and immunoprecipitation buffer volumes were multiplied by the number of antibodies to probe per sample. DNA samples were sonicated (40 amp) three times for 2 minutes at 4°C to obtain an average fragment size of 500 to 1000 base pairs. Cell debris were removed by centrifugation at 4°C and the supernatant transferred to a fresh Eppendorf tube. To determine the relative amount of DNA immunoprecipitated compared to the total amount of DNA, 100 µl of supernatant was removed, treated with 0.5 mg/ml pronase at 37°C for four hours then stored on ice. To immunoprecipate protein-DNA complexes, 800 µl of the remaining supernatant was incubated with individual antibodies (2 µl anti-SSB, 2 µl anti-PriA, 2 µl anti-DnaD, 2 µl anti-DnaB, 2 µl anti-DnaI, 2 µl anti-DnaC, 2 µl anti-mNeonGreen or 4 µl anti-His) for 90 minutes at room temperature. 750 µg of protein G Dynabeads were equilibrated by washing with bead buffer (1X PBS, 0.01% v/v Tween 20), resuspended in 25 µl of bead buffer and incubated with the sample supernatant for 1 hour at room temperature. Immunoprecipitated complexes were collected by applying the mixture to a magnet and washed with the following buffers in the respective order: 0.5X immunoprecipitation buffer for 15 min, 0.5X immunoprecipitation buffer supplemented with 500 mM NaCl for 15 min, stringent wash buffer (250 mM LiCl, 10 mM Tris-HCl pH 8.0, 0.5% v/v Igepal, 0.5% w/v sodium deoxycholate, 10 mM EDTA) for 20 min. Finally, protein-DNA complexes were washed a further three times with TET buffer (10 mM Tris-HCl pH 8.0, 1 mM EDTA, 0.01% v/v Tween 20) and resuspended in 100 µl of TEN buffer (10 mM Tris-HCl pH 8.0, 1 mM EDTA, 300 mM NaCl). Formaldehyde crosslinks of both the immunoprecipitate and total DNA were reversed by incubation at 65°C for 18 hours in the presence of 1 mg/ml proteinase K. DNA was then removed from the magnetic beads, cleaned using QIAquick PCR purification columns and used for quantitative PCR analyses. Quantitative PCR Quantitative PCR (qPCR) was performed using the Luna qPCR master mix to measure relative amounts of DNA bound by SSB, PriA, DnaD, DnaB, DnaI, DnaC, mNeonGreen-AddA, mNeonGreen-AddB, His-RecO and His-RecA at specific genomic locations (e.g. near the nick site located at -90° on the chromosome or the origin oriC compared to a control site located at +90° on the chromosome). All PCR reactions were assembled using the QIAgility robotic workstation in 20 μl reaction volumes in a Rotor-Disc 100 and qPCRs were run on a Rotor-Gene Q instruments. Standard curves were obtained using the Rotor-Gene Q software to calculate the efficiency of each primer pair, which varied ∼5% between sets. Oligonucleotides were designed to amplify specific genomic regions using the Primer3Plus tool 91 , were typically 20-25 bases in length (Table S4) and amplified a ~100 bp PCR product. Individual fold enrichment ratios were obtained as follows: first, every Ct value was converted to 1/2 Ct and technical triplicates were averaged to generate a single enrichment value; second, genomic location specific enrichment was normalised by corresponding values obtained at +90° on the chromosome. Error bars indicate the standard error of the mean for three biological replicates. Protein structure prediction and representation Protein models for SSB:ssDNA and SSB:RecO were generated using AlphaFold 3 and polypeptides corresponding to individual protein entities as input 92 . Main and alternative models were manually examined and Model 0 was chosen to highlight key features using the Pymol Molecular Graphics 2.1 software 93 . Structural highlights of the AddAB complex were derived from the crystal structure PDB: 4CEH 94 . Declarations DATA AVAILABILITY Lead contact For additional information and requests for resources and reagents, please contact the lead contact: Charles Winterhalter ( [email protected] ). Materials availability Genetic materials generated in this study can be obtained directly from the lead contact. Data and code availability R code (DOI: 10.5281/zenodo.14793122) and protein structures for SSB:ssDNA and SSB:RecO complexes (DOI: 10.5281/zenodo.14793919) are available from Zenodo. ACKNOWLEDGEMENTS This work was supported by a Wellcome Trust Early-Career Award [226338/Z/22/Z] to CW and Wellcome Trust Discovery Award [225811/Z/22/Z] to HM. We would also like to express our gratitude to Frances Davison for technical assistance, James Grimshaw for microscope maintenance and Frederic Schramm for insightful discussions prior to submission. AUTHOR CONTRIBUTIONS CW and HM contributed to the conception/design of the work. CW and SF generated results presented in the manuscript. CW created figures. CW and HM wrote/edited the manuscript. ETHICS DECLARATION The authors declare no competing interests. DECLARATION OF GENERATIVE AI AND AI-ASSISTED TECHNOLOGIES During the preparation of this work, the authors used AlphaFold 3 to generate SSB:ssDNA and SSB:RecO protein models. After using this tool, the authors reviewed the models and edited their content as needed and take full responsibility for the content of the publication. 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Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-6364374","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Article","associatedPublications":[],"authors":[{"id":442873159,"identity":"6902bdba-1b7a-4870-a3aa-fcaf43904d2b","order_by":0,"name":"Charles Winterhalter","email":"data:image/png;base64,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","orcid":"https://orcid.org/0000-0001-9673-1227","institution":"Newcastle University","correspondingAuthor":true,"prefix":"","firstName":"Charles","middleName":"","lastName":"Winterhalter","suffix":""},{"id":442873160,"identity":"8b9bf71f-a899-41d8-a444-a02cd72c864f","order_by":1,"name":"Stepan Fenyk","email":"","orcid":"","institution":"Newcastle University","correspondingAuthor":false,"prefix":"","firstName":"Stepan","middleName":"","lastName":"Fenyk","suffix":""},{"id":442873161,"identity":"e437ae59-a126-46b4-b009-3a5e89ec89e9","order_by":2,"name":"Heath Murray","email":"","orcid":"https://orcid.org/0000-0002-6467-3656","institution":"Newcastle University","correspondingAuthor":false,"prefix":"","firstName":"Heath","middleName":"","lastName":"Murray","suffix":""}],"badges":[],"createdAt":"2025-04-02 22:25:42","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-6364374/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-6364374/v1","draftVersion":[],"editorialEvents":[{"content":"https://doi.org/10.1038/s41467-025-66550-w","type":"published","date":"2025-11-26T05:00:00+00:00"}],"editorialNote":"","failedWorkflow":false,"files":[{"id":82236917,"identity":"ca00f6de-5802-4327-90d5-238dfb425824","added_by":"auto","created_at":"2025-05-08 07:18:08","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":237926,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eSingle-strand discontinuities arrest DNA synthesis and generate DSBs.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A)\u003c/strong\u003eDiagram illustrating the impact of leading and lagging strand discontinuities on a replication fork, with the bacterial replicative helicase translocating along the template for the lagging strand.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(B)\u003c/strong\u003eGenetic system employed to create a site-specific single-strand discontinuity in the \u003cem\u003eB. subtilis\u003c/em\u003e chromosome. Black boxes correspond to antibiotic resistance markers, pink to Cas9 variants and green to sgRNA. Chromosome annotations refer to the replication origin (0), terminus (180) and sgRNA target sequence (yellow box).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(C)\u003c/strong\u003eSpot titre analysis of strains engineered to express (+xylose) either nCas9 or dCas9. Individual sgRNAs were designed to target both strands at loci located at either -90° (top panel) or +90° (bottom panel).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(D)\u003c/strong\u003eMFA using whole genome sequencing of strains engineered to express either nCas9 or dCas9. Individual sgRNAs were designed to target both strands at loci located at either -90° (left panel) or +90° (right panel). The frequency of sequencing reads was plotted against genome position.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(E) \u003c/strong\u003eRepresentative fluorescence microscopy images of the DSB reporter mNG-RecA in strains engineered to express (+xylose) either nCas9 or dCas9 (sgRNAs target -90°). Fluorescent RecA organises into bundles in the presence of DSBs. Green signal corresponds to mNG-RecA fluorescence, grey scale images show corresponding phase contrast images.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(F)\u003c/strong\u003eQuantification of fluorescence microscopy images of the DSB reporter mNG-RecA in strains engineered to express (+xylose) either nCas9 or dCas9 (sgRNA target -90°). Detection of spots and bundles corresponds to distinct foci or filament-like structures annotated in panel (E). Error bars indicate the standard error of the mean.\u003c/p\u003e","description":"","filename":"1.png","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/7b482b5b33102cdec9a18e88.png"},{"id":82235469,"identity":"b5e8be3d-65a4-4f58-8558-3720f2eb9be4","added_by":"auto","created_at":"2025-05-08 07:02:08","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":151754,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eReplication restart is necessary to resume DNA synthesis following replisome inactivation at a single-strand discontinuity.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A)\u003c/strong\u003eDiagram illustrating genomic locations probed via ChIP-qPCR at loci located at either -90° or +90°. The sgRNA targeted Cas9 protein variants to the locus at -90°. The enrichment signal at the +90° locus was used for fold change normalisation. Lagging strand and leading strand templates are represented in black and grey lines, respectively (numbers refer to polarity). Blue arrows indicate leading strand synthesis and red arrows indicate lagging strand synthesis.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(B-F)\u003c/strong\u003eChIP-qPCR analyses of helicase (DnaC) and replication restart proteins (PriA, DnaD, DnaB, DnaI) in strains engineered to express either nCas9 or dCas9. The sgRNAs targeted Cas9 proteins to a locus located at -90° and fold change was normalised to enrichment values obtained at the +90° locus. Error bars indicate the standard error of the mean.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(G)\u003c/strong\u003eDiagram illustrating the fate of helicase (red hexagon) upon encountering a single-strand discontinuity in either the leading or lagging strand template.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(H)\u003c/strong\u003eMFA using whole genome sequencing in strains engineered to express lower levels of sgRNAs and Cas9 proteins (weak nicking system). The frequency of sequencing reads was plotted against genome position.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(I)\u003c/strong\u003eGenetic system employed to express low levels of PriA in a background encoding the weak nicking system.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(J)\u003c/strong\u003eSpot titre analysis of strains with a low level of PriA engineered to express nCas9 (+xylose). sgRNAs were targeted to a locus at -90°.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(K)\u003c/strong\u003eMFA using whole genome sequencing in strains with a low level of PriA engineered to express (+xylose) nCas9. The frequency of sequencing reads was plotted against genome position.\u003c/p\u003e","description":"","filename":"2.png","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/ebc2bd99bf274a38ecae07e1.png"},{"id":82236508,"identity":"b94c5b72-f7ad-4ec8-a0da-b6a68caa9419","added_by":"auto","created_at":"2025-05-08 07:10:08","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":270565,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eThe repair of single-strand discontinuities requires homologous recombination.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A)\u003c/strong\u003eSpot titre analysis of homologous recombination mutants in strains engineered to express (+xylose) nCas9. sgRNAs were targeted to a locus at -90° and control strains correspond to parental strains.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(B)\u003c/strong\u003eMFA using whole genome sequencing of homologous recombination mutants in strains engineered to express nCas9. The frequency of sequencing reads was plotted against genome position.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(C)\u003c/strong\u003eDiagram illustrating proposed molecular events required for homologous recombination to repair the DSB created after a replication fork encounters a single-strand discontinuity in the leading strand template.\u003c/p\u003e","description":"","filename":"3.png","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/84119c0e6727403124aae123.png"},{"id":82235475,"identity":"b5e56913-be35-4225-963f-04bca4fd0060","added_by":"auto","created_at":"2025-05-08 07:02:08","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":707404,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eThe SSB-CTT is essential to recruit RecO following replisome inactivation at a single-strand discontinuity.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A)\u003c/strong\u003eChIP-qPCR analyses of SSB in strains engineered to express nCas9. The sgRNAs targeted nCas9 to a locus located at -90° and fold change was normalised to enrichment values obtained at the +90° locus. Error bars indicate the standard error of the mean.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(B)\u003c/strong\u003eChIP-qPCR analyses of SSB in strains with homologous recombination mutants engineered to express nCas9. qPCR primers amplified either a site located at -3 kb from the sgRNA target (-90°) or a control locus (+90°) used for normalisation. Error bars indicate the standard error of the mean.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(C)\u003c/strong\u003eSpot titre analysis of SSB variants in strains engineered to express (+xylose) nCas9. Top panel shows the genetic system employed in this experiment. sgRNAs were targeted to a locus at -90°.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(D)\u003c/strong\u003eMFA using whole genome sequencing in strains with SSB variants engineered to express nCas9. The frequency of sequencing reads was plotted against genome position.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(E)\u003c/strong\u003eChIP-qPCR analyses of SSB, His-RecO and His-RecA proteins in strains engineered to express nCas9. qPCR primers amplified either a site located at -3 kb from the sgRNA target (-90°) or a control locus (+90°) used for normalisation. Error bars indicate the standard error of the mean.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(F)\u003c/strong\u003eChIP-qPCR analyses of His-RecA in strains with homologous recombination mutants engineered to express nCas9. Top panel shows the genetic system employed in this experiment. qPCR primers amplified either a site located at -3 kb from the sgRNA target (-90°) or a control locus (+90°) used for normalisation. Error bars indicate the standard error of the mean.\u003c/p\u003e","description":"","filename":"4.png","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/e21539b0789849d45779e9ca.png"},{"id":82236511,"identity":"10f3b628-08fd-4814-b6ac-9cb45f9d75e6","added_by":"auto","created_at":"2025-05-08 07:10:08","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":843499,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eDNA end resection via AddAB is not required to recruit SSB following replisome inactivation at a single-strand discontinuity.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A) \u003c/strong\u003e\u0026nbsp;MFA using whole genome sequencing in a Δ\u003cem\u003erecG\u003c/em\u003e mutant strain engineered to express nCas9. The frequency of sequencing reads was plotted against genome position (same data as in Fig. 3B Δ\u003cem\u003erecG\u003c/em\u003e panel). A region flanking the sgRNA binding site is enlarged.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(B)\u003c/strong\u003e Diagram illustrating incorrect helicase reloading leading to reverse restart on a repaired replication fork in a Δ\u003cem\u003erecG\u003c/em\u003e mutant. Helicase is shown as a red hexagon and RecG is shown in green; protein movement is indicated by colour-corresponding arrows. Lagging strand and leading strand templates are represented in black and grey lines, respectively. Blue arrows indicate leading strand synthesis.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(C)\u003c/strong\u003e ChIP-qPCR analyses of mNG-AddA and mNG-AddB in strains engineered to express nCas9. Left panel shows the genetic system employed in this experiment. The sgRNAs targeted nCas9 to a locus located at -90° and fold change was normalised to enrichment values obtained at the +90° locus. Error bars indicate the standard error of the mean.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(D)\u003c/strong\u003e Spot titre analysis in strains with either AddA\u003csup\u003eΔHEL\u003c/sup\u003e, AddA\u003csup\u003eΔNUC\u003c/sup\u003e or AddB\u003csup\u003eΔNUC\u003c/sup\u003e engineered to express (+xylose) nCas9. sgRNAs were targeted to a locus at -90° and control strains correspond to parental strains.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(E)\u003c/strong\u003e MFA using whole genome sequencing of strains with either AddA\u003csup\u003eΔHEL\u003c/sup\u003e or AddA\u003csup\u003eΔNUC\u003c/sup\u003e/AddB\u003csup\u003eΔNUC\u003c/sup\u003e engineered to express nCas9. The frequency of sequencing reads was plotted against genome position. Top panel shows the genetic system employed in this experiment.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(F)\u003c/strong\u003e ChIP-qPCR analyses of SSB in strains with AddA and AddB variants engineered to express nCas9. Alleles refer to wild-type (WT), knockouts (Δ), and point mutations inactivating nuclease (ΔNUC) or helicase (ΔHEL) activities. qPCR primers amplified either a site located at -3 kb from the sgRNA target (-90°) or a control locus (+90°) used for normalisation. Error bars indicate the standard error of the mean.\u0026nbsp;\u003c/p\u003e","description":"","filename":"5.png","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/4292e03b77ad5140d42dcf67.png"},{"id":82236510,"identity":"76af89ad-a3b5-457a-b527-5fe891168f96","added_by":"auto","created_at":"2025-05-08 07:10:08","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":470873,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eRecG is dispensable when DSB processing degrades the strand with a 5′-end.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A)\u003c/strong\u003e Diagram illustrating differential resection of a DSB in the presence of either the AddA or AddB nuclease activity. Leading and lagging strand templates are indicated by grey and blue lines, respectively (numbers indicate polarity). Dotted lines indicate nuclease cleavage.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(B)\u003c/strong\u003e Genetic system employed to investigate the role of RecG in DSB repair following replisome inactivation at a single-strand discontinuity.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(C)\u003c/strong\u003e Spot titre analysis in a Δ\u003cem\u003erecG\u003c/em\u003e mutant strain with either AddA\u003csup\u003eΔNUC\u003c/sup\u003e or AddB\u003csup\u003eΔNUC\u003c/sup\u003e engineered to express (+xylose) nCas9. sgRNAs were targeted to a locus at -90° and control strains correspond to parental strains encoding the weak nicking system.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(D)\u003c/strong\u003e Diagram illustrating differential helicase reloading events on a repaired replication fork when AddA or AddB nuclease activity is inactivated in the absence of RecG. Helicase is shown as a red hexagon and PriA in yellow; protein movement is indicated by colour-corresponding arrows. Lagging strand and leading strand templates are represented in black and grey lines, respectively. Blue arrows indicate leading strand synthesis.\u003c/p\u003e","description":"","filename":"6.png","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/41b0ed8de2425c4fac9a78e4.png"},{"id":82235480,"identity":"b94d2595-aa04-4475-8deb-5352267cbf50","added_by":"auto","created_at":"2025-05-08 07:02:08","extension":"png","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":749237,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eModel for DSB repair and helicase reloading following replisome inactivation at a single-strand discontinuity.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eRevised model illustrating molecular events that are required or dispensable for homologous repair of a DSB created after a replication fork encounters a single-strand discontinuity located on either the leading or lagging strand template.\u003c/p\u003e","description":"","filename":"7.png","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/44ee1e4b4c86ef3376cc687c.png"},{"id":99211926,"identity":"dbd44ce9-7bc7-46ac-b531-60d45fc508e1","added_by":"auto","created_at":"2025-12-30 08:21:01","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":4768165,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/e86f297b-9397-4a68-b86c-f581cb1f3923.pdf"},{"id":82235467,"identity":"7ef449d8-29c3-4d31-b3fc-eb5b40346ef4","added_by":"auto","created_at":"2025-05-08 07:02:08","extension":"docx","order_by":1,"title":"","display":"","copyAsset":false,"role":"supplement","size":32486,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cbr\u003e\u003c/p\u003e","description":"","filename":"SUPPLEMENTARYFIGURETITLESANDLEGENDS.docx","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/e0da174ff1adeb817a185d63.docx"},{"id":82235468,"identity":"ac287fb1-f6af-4570-93f0-1235ed3a97b0","added_by":"auto","created_at":"2025-05-08 07:02:08","extension":"xlsx","order_by":2,"title":"","display":"","copyAsset":false,"role":"supplement","size":10524,"visible":true,"origin":"","legend":"Supplementary Table 1","description":"","filename":"20250226TableS1.xlsx","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/2f68a4f21050778dfcf7b613.xlsx"},{"id":82235472,"identity":"faece524-e1c9-4533-81b5-fa392f3d2044","added_by":"auto","created_at":"2025-05-08 07:02:08","extension":"xlsx","order_by":3,"title":"","display":"","copyAsset":false,"role":"supplement","size":24481,"visible":true,"origin":"","legend":"Supplementary Table 2","description":"","filename":"20250226TableS2.xlsx","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/611454382ed7584f9a2658f1.xlsx"},{"id":82235473,"identity":"1810e86c-8418-429e-84bf-bc7e53008bc0","added_by":"auto","created_at":"2025-05-08 07:02:08","extension":"xlsx","order_by":4,"title":"","display":"","copyAsset":false,"role":"supplement","size":147055,"visible":true,"origin":"","legend":"Supplementary Table 3","description":"","filename":"20250226TableS3.xlsx","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/77711eaa6302acbeea9a841c.xlsx"},{"id":82235477,"identity":"bc59f272-e5f6-4f41-88fb-619c1147787f","added_by":"auto","created_at":"2025-05-08 07:02:08","extension":"xlsx","order_by":5,"title":"","display":"","copyAsset":false,"role":"supplement","size":10404,"visible":true,"origin":"","legend":"Supplementary Table 4","description":"","filename":"20250226TableS4.xlsx","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/04f607b53488d4fce1d1f0c4.xlsx"},{"id":82235490,"identity":"a3636c79-4a2c-4402-ae05-4b2bfb754a9c","added_by":"auto","created_at":"2025-05-08 07:02:09","extension":"eps","order_by":6,"title":"","display":"","copyAsset":false,"role":"supplement","size":67002526,"visible":true,"origin":"","legend":"Supplementary Figure 1","description":"","filename":"20250228figures9.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/e147c1888a7d33898eb41bf7.eps"},{"id":82236918,"identity":"6985bf82-1a1d-4a43-bd62-c41e5089c295","added_by":"auto","created_at":"2025-05-08 07:18:09","extension":"eps","order_by":7,"title":"","display":"","copyAsset":false,"role":"supplement","size":3122075,"visible":true,"origin":"","legend":"Supplementary Figure 2","description":"","filename":"20250228figures10.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/28a147e8d8a678b1a773e79e.eps"},{"id":82235486,"identity":"bfce1437-9012-44a3-a4ba-db6c8ef1341b","added_by":"auto","created_at":"2025-05-08 07:02:09","extension":"eps","order_by":8,"title":"","display":"","copyAsset":false,"role":"supplement","size":15156927,"visible":true,"origin":"","legend":"Supplementary Figure 3","description":"","filename":"20250228figures11.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/3182eb5d6edb9d472329e4b0.eps"},{"id":82236512,"identity":"018f7caf-8988-48eb-9627-7f0e2057449f","added_by":"auto","created_at":"2025-05-08 07:10:09","extension":"eps","order_by":9,"title":"","display":"","copyAsset":false,"role":"supplement","size":6636325,"visible":true,"origin":"","legend":"Supplementary Figure 4","description":"","filename":"20250228figures12.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/0c5a1aadab0d143f50f6e22e.eps"},{"id":82235487,"identity":"3b4d0cdf-754c-4500-9021-8e3df693ac26","added_by":"auto","created_at":"2025-05-08 07:02:09","extension":"eps","order_by":10,"title":"","display":"","copyAsset":false,"role":"supplement","size":43281262,"visible":true,"origin":"","legend":"Supplementary Figure 5","description":"","filename":"20250228figures13.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/3746b2bb89cdd5fe37c249b9.eps"},{"id":82235489,"identity":"95300f1a-7bbf-4e77-bf75-14d27c3cc0ca","added_by":"auto","created_at":"2025-05-08 07:02:09","extension":"eps","order_by":11,"title":"","display":"","copyAsset":false,"role":"supplement","size":28210293,"visible":true,"origin":"","legend":"Supplementary Figure 6","description":"","filename":"20250228figures14.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/0d410b7ce4503f004563e1ff.eps"},{"id":82235492,"identity":"377f85ea-8283-48d9-9375-8db092b91504","added_by":"auto","created_at":"2025-05-08 07:02:15","extension":"eps","order_by":12,"title":"","display":"","copyAsset":false,"role":"supplement","size":133191219,"visible":true,"origin":"","legend":"Supplementary Figure 7","description":"","filename":"20250228figures15.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/a21f808a341d121a8ddfcd43.eps"},{"id":82235491,"identity":"2e1b9664-ebd3-48a6-a639-a75748130ec9","added_by":"auto","created_at":"2025-05-08 07:02:11","extension":"eps","order_by":13,"title":"","display":"","copyAsset":false,"role":"supplement","size":82517600,"visible":true,"origin":"","legend":"Supplementary Figure 8","description":"","filename":"20250228figures1616.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/563213c6d64d6c113d348aed.eps"},{"id":82236515,"identity":"74890ef5-7ee0-46b3-aa36-60323f8e0f30","added_by":"auto","created_at":"2025-05-08 07:10:09","extension":"eps","order_by":14,"title":"","display":"","copyAsset":false,"role":"supplement","size":31834733,"visible":true,"origin":"","legend":"Supplementary Figure 9","description":"","filename":"20250228figures17.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/f5f661f9dd496cf4dbe6c604.eps"},{"id":82235479,"identity":"c107c986-3081-419a-b69d-93616c5c654b","added_by":"auto","created_at":"2025-05-08 07:02:08","extension":"eps","order_by":15,"title":"","display":"","copyAsset":false,"role":"supplement","size":7385066,"visible":true,"origin":"","legend":"\u003cp\u003eSupplementary Figure 10\u003c/p\u003e","description":"","filename":"20250228figures18.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/57f72197c94070527282113f.eps"},{"id":82235483,"identity":"3f64cd98-f23f-4bcd-9212-96904541556e","added_by":"auto","created_at":"2025-05-08 07:02:09","extension":"eps","order_by":16,"title":"","display":"","copyAsset":false,"role":"supplement","size":5074590,"visible":true,"origin":"","legend":"Supplementary Figure 11","description":"","filename":"20250228figures19.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/807f614fc407f729b0e7bf33.eps"},{"id":82236513,"identity":"f233edb3-ec7a-41bc-b95c-4de029411d46","added_by":"auto","created_at":"2025-05-08 07:10:09","extension":"eps","order_by":17,"title":"","display":"","copyAsset":false,"role":"supplement","size":13642676,"visible":true,"origin":"","legend":"\u003cp\u003eSupplementary Figure 12\u003c/p\u003e","description":"","filename":"20250228figures20.eps","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/fce6d2a54d75e048e1694c27.eps"},{"id":82235481,"identity":"e000b983-ea83-49e7-ad1f-d47bbef6a78f","added_by":"auto","created_at":"2025-05-08 07:02:09","extension":"png","order_by":18,"title":"","display":"","copyAsset":false,"role":"supplement","size":72774,"visible":true,"origin":"","legend":"","description":"","filename":"GraphicalAbstract.png","url":"https://assets-eu.researchsquare.com/files/rs-6364374/v1/88f093221896a6f9902f3ef9.png"}],"financialInterests":"There is \u003cb\u003eNO\u003c/b\u003e Competing Interest.\nN/A","formattedTitle":"Rescuing bacterial genome replication: essential functions to repair a double-strand break and restart DNA synthesis","fulltext":[{"header":"Highlights","content":"\u003cul\u003e\n \u003cli\u003eSingle-strand discontinuities inactivate the bacterial replisome\u003c/li\u003e\n \u003cli\u003eNicks on the leading or lagging strand template differentially affect fate of the helicase\u003c/li\u003e\n \u003cli\u003eDouble-strand breaks are repaired via homologous recombination and elicit the replication restart pathway\u003c/li\u003e\n \u003cli\u003eResection often generates a DNA strand with a 5\u0026prime;-end that must be processed to ensure productive helicase reloading\u003c/li\u003e\n\u003c/ul\u003e"},{"header":"MAIN","content":"\u003cp\u003eEmergence of antibiotic resistant bacterial pathogens represents an urgent threat to human health and food production\u003csup\u003e\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e\u003c/sup\u003e. Understanding how bacteria respond to antibiotics is necessary to generate effective treatments. While antibiotics generally target specific essential bacterial systems (e.g. cell wall synthesis, translation, transcription) to inhibit bacterial proliferation, there is growing evidence that diverse antibiotics share the potential to disrupt bacterial physiology, thereby generating a second challenge to pathogens\u003csup\u003e\u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e\u003c/sup\u003e. Work in several bacterial species indicates that a range of antibiotics ultimately generate reactive oxygen species (ROS) which can damage biomolecules including DNA\u003csup\u003e\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e,\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e\u003c/sup\u003e. Importantly, antibiotics that generate ROS elicit the bacterial DNA damage response, indicating the presence of double-strand breaks (DSB) under these conditions\u003csup\u003e\u003cspan additionalcitationids=\"CR5 CR6\" citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eThe potency of ROS-mediated DNA damage has been suggested to arise from conflicts between genome repair and replication systems\u003csup\u003e\u003cspan additionalcitationids=\"CR5\" citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e,\u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e\u003c/sup\u003e. Oxidised nucleobases located within the genome must be excised as part of the requisite repair pathway, leading to transient single-strand discontinuities in the DNA template\u003csup\u003e\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e\u003c/sup\u003e. If a bacterial replication fork arrives at a single-strand discontinuity, the encounter generates a DSB\u003csup\u003e\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e10\u003c/span\u003e\u003c/sup\u003e. Elegant experiments using bacteriophage \u003cem\u003eλ\u003c/em\u003e demonstrated that an engineered nick in either strand of the viral genome generated replication-dependent DSBs (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eA)\u003csup\u003e\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e\u003c/sup\u003e. While active DNA replication is necessary to generate DSBs at a nick, the fate of the bacterial DNA replication machinery (replisome) at single-strand discontinuities has not been determined.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eThe replisome is a multicomponent complex of proteins and enzymes that unwinds the chromosome and coordinates synthesis of new DNA polymers on the leading and lagging strands\u003csup\u003e\u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e,\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e\u003c/sup\u003e. The most stable component of the replisome and most pivotal for its processivity is helicase\u003csup\u003e\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e,\u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e\u003c/sup\u003e. Cellular replicative helicases are toroid enzymes that unwind DNA by encircling and translocating along a single client nucleic acid strand, acting as a wedge to disrupt base pairing\u003csup\u003e\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e\u003c/sup\u003e. Studies of bacterial replisomes \u003cem\u003ein vitro\u003c/em\u003e and \u003cem\u003ein vivo\u003c/em\u003e have revealed that the machinery is intrinsically dynamic, with most factors observed to bind and unbind interaction partners multiple times per replication cycle\u003csup\u003e\u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e,\u003cspan citationid=\"CR18\" class=\"CitationRef\"\u003e18\u003c/span\u003e\u003c/sup\u003e. This behaviour enables the replisome to overcome a range of potential roadblocks and remain associated with the template, competent to continue DNA synthesis\u003csup\u003e\u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e\u003c/sup\u003e. However, recent studies in eukaryotic systems have indicated that single-strand discontinuities generate DSBs and arrest DNA replication by impairing helicase progression\u003csup\u003e\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e,\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eDetermining how bacteria react and respond when a replication fork encounters a single-strand discontinuity is crucial to understanding how ROS inhibits bacterial proliferation. Here we have investigated this question by expressing Cas9 nickases (nCas9) in the model bacterium \u003cem\u003eBacillus subtilis\u003c/em\u003e. The results show that a nick located in either template strand blocks downstream DNA synthesis, and that resumption of DNA replication is dependent upon a core set of homologous recombination proteins and the PriA-dependent replication restart system necessary to reload the replicative helicase. Unexpectedly, interrogation of the DNA repair pathway suggests that processing of the DSB often leaves the 5\u0026prime;-terminated DNA strand intact, which can hybridise to the homologous chromosome during recombination and perturb replication restart. We ascribe the enigmatic RecG helicase as the factor that remodels these recombination products containing a 5\u0026prime;-terminated DNA strand, thereby generating the replication fork structure recognised by PriA for productive helicase loading. These findings have critical implications for the fundamental mechanism of homologous recombination in bacteria and the aim of potentiating ROS-dependent antibiotic toxicity.\u003c/p\u003e"},{"header":"RESULTS","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003eSingle-strand discontinuities in a bacterial chromosome arrest DNA synthesis\u003c/h2\u003e \u003cp\u003eOxidised nucleobases generated by ROS can be located randomly throughout a genome, making it a challenge to directly analyse conflicts with DNA replication during base excision repair. To mimic the activity of oxidised base repair, an experimental system capable of generating a site-specific single-strand discontinuity was developed in \u003cem\u003eB. subtilis\u003c/em\u003e using Cas9 nuclease variants and single-guide RNAs (sgRNA)\u003csup\u003e\u003cspan additionalcitationids=\"CR21 CR22\" citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e23\u003c/span\u003e\u003c/sup\u003e. Cas9 carrying single amino acid substitutions (Cas9\u003csup\u003eD\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e10\u003c/span\u003eA\u003c/sup\u003e or Cas9\u003csup\u003eH840A\u003c/sup\u003e) created nickases (nCas9), while the double mutant (Cas9\u003csup\u003eD10A/H840A\u003c/sup\u003e) created a catalytically inactive enzyme (dCas9)\u003csup\u003e\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e24\u003c/span\u003e\u003c/sup\u003e. Each \u003cem\u003ecas9\u003c/em\u003e allele was integrated into the \u003cem\u003eB. subtilis\u003c/em\u003e chromosome under the control of a xylose-inducible promoter (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eB)\u003csup\u003e\u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e25\u003c/span\u003e\u003c/sup\u003e. The location of sgRNA hybridisation is indicated by coordinates between the origin (0\u0026deg;) and terminus (~\u0026thinsp;180\u0026deg;) on either the right (+) or left (-) replicore of the chromosome. To avoid complications arising from potential activation of prophage following DNA damage in the laboratory strain of \u003cem\u003eB. subtilis\u003c/em\u003e, a derivative lacking these genetic elements (Δ6) was used throughout this study\u003csup\u003e\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e26\u003c/span\u003e\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eCell viability was quantified using spot-titre assays, where strains are grown to saturation in liquid medium without xylose, before samples were serially diluted and plated onto solid medium with or without xylose to enumerate the number of colony forming units (CFUs). It was observed that the Cas9\u003csup\u003eD\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e10\u003c/span\u003eA\u003c/sup\u003e nickase elicited a stronger phenotype than Cas9\u003csup\u003eH840A\u003c/sup\u003e upon induction with xylose (Fig. \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003eA-B)\u003csup\u003e\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e24\u003c/span\u003e\u003c/sup\u003e. To achieve comparable nicking efficiency at different sites in the chromosome, the \u003cem\u003ecas9\u003c/em\u003e\u003csup\u003e\u003cem\u003eD\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e10\u003c/span\u003eA\u003c/em\u003e\u003c/sup\u003e allele was used throughout the remainder of this study, in conjunction with distinct sgRNAs targeted to the template of either the leading or the lagging strand (in non-essential regions of the genome). Using sgRNAs to target leading and lagging strands at loci on both left and right chromosome arms, it was observed that expression of nCas9 resulted in severe growth inhibition (Figs.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eC, S1C). In contrast, expression of dCas9 targeted to the same locations did not affect the number of CFUs (Figs.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eC, S1D), indicating that DNA nicking was necessary for the observed growth defect.\u003c/p\u003e \u003cp\u003eNext chromosome replication was investigated using whole genome marker frequency analysis (MFA). Expression of Cas9 variants was induced for one hour in exponentially growing cultures. Genomic DNA (gDNA) was harvested for next-generation sequencing, reads were mapped onto the reference genome sequence, and data is displayed with the chromosome origin (\u003cem\u003eoriC\u003c/em\u003e) at the center of the graphs (Fig. \u003cspan refid=\"MOESM2\" class=\"InternalRef\"\u003eS2\u003c/span\u003e). It was observed that nicks in either the leading or lagging strand resulted in a drop in sequencing coverage from the nick site to the terminus region, whereas expression of dCas9 with the same sgRNAs did not appear to inhibit DNA replication (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eD). Because cells expressing nCas9 are unable to synthetise DNA downstream of the nick, this likely explains the strong growth inhibition observed in spot-titre assays (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eC).\u003c/p\u003e \u003cp\u003ePrevious results indicated that a replication fork encountering a single-strand discontinuity generates a DSB\u003csup\u003e\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e\u003c/sup\u003e. To determine whether DSBs were being formed in \u003cem\u003eB. subtilis\u003c/em\u003e following nCas9 induction, localisation of the reporter mNeonGreen-RecA (mNG-RecA) was observed in live cells using fluorescence microscopy. It has been shown that following artificial endonuclease cleavage to create a DSB, fluorescently labelled RecA recombinase assembles into bundles\u003csup\u003e\u003cspan additionalcitationids=\"CR28\" citationid=\"CR28\" class=\"CitationRef\"\u003e27\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e29\u003c/span\u003e\u003c/sup\u003e. Prior to induction of nCas9, mNG-RecA is observed evenly distributed over the nucleoid (Fig. \u003cspan refid=\"MOESM3\" class=\"InternalRef\"\u003eS3\u003c/span\u003eA). Following expression of nCas9 for one hour during exponential growth, mNG-RecA bundles were detected in 58.5% of cells when the leading strand template was nicked and 48% of cells when the lagging strand template was nicked (Figs.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eE-F, S3B). In contrast, expression of dCas9 did not significantly promote mNG-RecA bundling (Figs.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eE-F, S3B). These results are consistent with the model that collision of a replication fork with a nick in either strand of the parental DNA duplex generates a DSB.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eSingle-strand discontinuities in a bacterial chromosome inactivate the replisome\u003c/h3\u003e\n\u003cp\u003eMFA indicated that under conditions expressing nCas9, DNA synthesis does not proceed beyond the nick. We wondered whether the replisome becomes arrested at a single-strand discontinuity, or rather whether the replication fork collapses. If the latter event occurs, then the bacterial replication restart system would be required to reload helicase and promote replisome assembly\u003csup\u003e\u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e30\u003c/span\u003e,\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e31\u003c/span\u003e\u003c/sup\u003e. In \u003cem\u003eB. subtilis\u003c/em\u003e there is only one known restart system composed of the essential proteins PriA and the helicase loaders DnaD, DnaB, and DnaI (note that in \u003cem\u003eB. subtilis\u003c/em\u003e the replicative helicase is DnaC)\u003csup\u003e\u003cspan additionalcitationids=\"CR33 CR34 CR35\" citationid=\"CR33\" class=\"CitationRef\"\u003e32\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e36\u003c/span\u003e\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eChromatin immunoprecipitation (ChIP) followed by quantitative PCR (qPCR) was used to determine the enrichment of helicase and replication restart proteins surrounding a nick in either the leading or lagging strand template. Following induction of nCas9 for one hour in exponentially growing cells, samples were harvested for ChIP. Along with sites flanking the nick region, protein enrichment was also probed at the origin \u003cem\u003eoriC\u003c/em\u003e as positive control and normalised to a site located on the opposite chromosome arm equidistant from \u003cem\u003eoriC\u003c/em\u003e (+\u0026thinsp;90, Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA). ChIP showed that helicase is specifically enriched upstream of nicks located in either template strand (Figs.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eB, S4). This upstream helicase enrichment was not detected when dCas9 was expressed, indicating that the Cas9-sgRNA nucleoprotein complex alone does not stall replication fork progression (Figs.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eB, S4).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eInterestingly, significant enrichment of helicase was observed downstream of the nick site only when cleavage occurred in the template used for leading strand synthesis (Figs.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eB, S4). Because the bacterial replicative helicase translocates along a single strand in the 5\u0026prime;\u0026loz;3\u0026prime; direction, ChIP results suggest that when the template for the lagging strand is nicked helicase runs-off the DNA substrate, whereas when the template for the leading strand is nicked helicase progresses downstream of the single-strand discontinuity (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eG). Consistent with the ChIP enrichment observed \u003cem\u003ein vivo\u003c/em\u003e (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA-B) and replisome inactivation (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eD), reconstituted unwinding assays \u003cem\u003ein vitro\u003c/em\u003e using bacterial helicase on model replication fork substrates have shown that if a flap is absent from the substrate (i.e. mimicking a nick), helicase transitions from encircling a single DNA strand onto double-stranded DNA (dsDNA) and slides along the duplex, no longer able to unwind\u003csup\u003e\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e37\u003c/span\u003e\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eThe enrichment of helicase upstream of nick sites is consistent with either helicase stalling or helicase reloading via the replication restart pathway. To distinguish between these models, ChIP was used to probe for the presence of replication restart proteins surrounding the nicks. Following nCas9 induction, enrichment of PriA, DnaD, DnaB and DnaI was detected upstream (but not downstream) of the sgRNA target sequences, while these proteins were absent from this region when dCas9 was expressed (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eC-F). These results suggest that nicks in either the leading or lagging strand template elicit the replication restart pathway for helicase reloading, whether helicase falls off the template (lagging strand template nick) or translocates downstream (leading strand template nick) in an inactive state (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eG).\u003c/p\u003e\n\u003ch3\u003eA weakened nCas9 system permits DNA replication restart\u003c/h3\u003e\n\u003cp\u003eWhile replication restart proteins were enriched upstream of nick sites (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eB-F), viability assays and MFA both indicated that downstream DNA synthesis was inhibited (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eC-D). We speculated that persistent nCas9 cleavage activity competes with productive DNA replication restart. Therefore, to allow interrogation of processes occurring downstream of DNA cleavage and replisome collapse, a weaker nCas9 system was developed. An \u003cem\u003essrA\u003c/em\u003e degradation tag\u003csup\u003e\u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e38\u003c/span\u003e\u003c/sup\u003e was fused to \u003cem\u003ecas9\u003c/em\u003e\u003csup\u003e\u003cem\u003eD\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e10\u003c/span\u003eA\u003c/em\u003e\u003c/sup\u003e to promote nCas9 proteolysis, and expression of both \u003cem\u003enCas9-ssrA\u003c/em\u003e and \u003cem\u003esgRNAs\u003c/em\u003e were placed under the control of xylose-inducible promoters (Fig. S5A). In this background, induction of nCas9 did not significantly inhibit cell viability or chromosome content (Fig. S5B-C) and MFA showed that DNA synthesis continued after a one-hour xylose induction, consistent with low tolerable levels of nickase activity (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eH). This weakened nicking system was employed for all experiments described below.\u003c/p\u003e\n\u003ch3\u003ePriA is required to restart DNA replication at single-strand DNA discontinuities\u003c/h3\u003e\n\u003cp\u003eThe master replication restart factor \u003cem\u003epriA\u003c/em\u003e is essential for growth of \u003cem\u003eB. subtilis.\u003c/em\u003e To ascertain whether PriA-dependent replication restart is required following chromosome nicking, a titratable PriA complementation system was developed (Fig. S6A). Here conditions were established to express PriA-ssrA at a low level, sufficient to maintain \u003cem\u003eB. subtilis\u003c/em\u003e viability while limiting the amount of PriA activity within a cell (Fig. S6B-C). When nCas9-ssrA and sgRNA were induced with limiting PriA-ssrA, spot-titre analyses showed that cell growth was inhibited in the absence of the endogenous \u003cem\u003epriA\u003c/em\u003e copy (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eI-J). Yet in this background, higher expression of PriA-ssrA rescued viability (Fig. S6D). Following induction of nCas9 and sgRNA for one hour during exponential growth, MFA showed that limited levels of PriA-ssrA could not sustain chromosome synthesis downstream of the nick site (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eK). Therefore, these data indicate that PriA is essential to restart DNA replication following replisome inactivation at single-strand DNA discontinuities.\u003c/p\u003e \u003cp\u003e \u003cb\u003eThe DSB resulting from a single-strand discontinuity is repaired using a core set of homologous recombination proteins\u003c/b\u003e \u003c/p\u003e \u003cp\u003eCytological analysis indicated that a nick in the template of either the leading or lagging strand generated a DSB (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eE). Strong evidence indicates that in bacteria such DSBs are repaired by homologous recombination, thereby recreating a replication fork to allow resumption of DNA synthesis\u003csup\u003e\u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e39\u003c/span\u003e\u003c/sup\u003e. To determine the factors required to repair DSBs following nCas9 nicking, a targeted reverse genetic analysis was employed, focused on genes previously implicated in homologous recombination and other DNA repair processes (Table \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003e). Using this approach, spot-titre analyses revealed that eight non-essential genes were required for normal growth upon nick induction: DNA end-processing genes \u003cem\u003eaddA\u003c/em\u003e/\u003cem\u003eaddB\u003c/em\u003e and homologous recombination genes \u003cem\u003erecF\u003c/em\u003e/\u003cem\u003erecO\u003c/em\u003e/\u003cem\u003erecR\u003c/em\u003e/\u003cem\u003erecA\u003c/em\u003e/\u003cem\u003erecG\u003c/em\u003e/\u003cem\u003erecU\u003c/em\u003e (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA and S7A). Functional complementation of each synthetic lethal deletion confirmed that these gene products are necessary to support growth in cells expressing nCas9 (Fig. S7B). Surprisingly, the screen revealed that several factors implicated in DNA repair are not essential under these conditions, such as those classed for end-processing (RecQ, RecS, RecJ)\u003csup\u003e\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e,\u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e40\u003c/span\u003e\u003c/sup\u003e, maintaining chromosome structure (\u0026ldquo;SMC-like\u0026rdquo; RecN, SbcCD, SbcEF)\u003csup\u003e\u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e41\u003c/span\u003e,\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e42\u003c/span\u003e\u003c/sup\u003e, recombination mediation (RecD2, RecX)\u003csup\u003e\u003cspan additionalcitationids=\"CR44\" citationid=\"CR44\" class=\"CitationRef\"\u003e43\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR47\" class=\"CitationRef\"\u003e45\u003c/span\u003e\u003c/sup\u003e and Holliday junction migration (RuvA, RuvB)\u003csup\u003e\u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e40\u003c/span\u003e\u003c/sup\u003e (Fig. S8).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eFocusing on the eight mutants that produced a clear growth defect, MFA was used to analyse DNA replication following induction of nCas9 and sgRNA in exponentially growing cells. The results showed that DNA end-processing (\u003cem\u003eaddA\u003c/em\u003e/\u003cem\u003eaddB\u003c/em\u003e) and recombination mutants (\u003cem\u003erecF\u003c/em\u003e/\u003cem\u003erecO\u003c/em\u003e/\u003cem\u003erecR\u003c/em\u003e/\u003cem\u003erecA\u003c/em\u003e/\u003cem\u003erecG\u003c/em\u003e) are defective in chromosome replication downstream of the nick site, with the only exception being \u003cem\u003erecU\u003c/em\u003e (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB; note subtle differences in replication profiles of parental strains containing deletions of homologous recombination genes, consistent with previous findings\u003csup\u003e\u003cspan citationid=\"CR48\" class=\"CitationRef\"\u003e46\u003c/span\u003e,\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e47\u003c/span\u003e\u003c/sup\u003e). These observations support the hypothesis that homologous recombination is necessary to repair the DSB and restart DNA replication.\u003c/p\u003e \u003cp\u003eBased on the proposed activities of these proteins, the following model was developed (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eC). AddAB helicase/nuclease processes the DSB to generate a single-stranded substrate with a 3\u0026prime;-end required for recombination\u003csup\u003e\u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e48\u003c/span\u003e,\u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e49\u003c/span\u003e\u003c/sup\u003e; RecFOR displaces the single-stranded binding protein (SSB) from single-stranded DNA substrates produced by AddAB, thereby allowing RecA to bind\u003csup\u003e\u003cspan citationid=\"CR52\" class=\"CitationRef\"\u003e50\u003c/span\u003e,\u003cspan citationid=\"CR53\" class=\"CitationRef\"\u003e51\u003c/span\u003e\u003c/sup\u003e; RecA is the recombinase that binds to single-stranded DNA (ssDNA) with a free 3\u0026prime;-end, identifies homologous DNA on the non-broken chromosome, and performs strand invasion and exchange\u003csup\u003e\u003cspan citationid=\"CR54\" class=\"CitationRef\"\u003e52\u003c/span\u003e,\u003cspan citationid=\"CR55\" class=\"CitationRef\"\u003e53\u003c/span\u003e\u003c/sup\u003e; RecU resolves the Holliday junction generated by RecA\u003csup\u003e\u003cspan citationid=\"CR56\" class=\"CitationRef\"\u003e54\u003c/span\u003e\u003c/sup\u003e. The role of RecG \u003cem\u003ein vivo\u003c/em\u003e is enigmatic\u003csup\u003e\u003cspan citationid=\"CR57\" class=\"CitationRef\"\u003e55\u003c/span\u003e,\u003cspan citationid=\"CR58\" class=\"CitationRef\"\u003e56\u003c/span\u003e\u003c/sup\u003e and discussed further below.\u003c/p\u003e\n\u003ch3\u003eThe SSB C-terminal tail is required for DNA repair following replisome inactivation\u003c/h3\u003e\n\u003cp\u003eThe requirement of the RecFOR system for repair of a seDSB strongly suggested that the essential single-strand binding protein (SSB) was also involved. To interrogate this hypothesis, recruitment of SSB was probed using ChIP following induction of nCas9 and sgRNA in exponentially growing cells. The results showed that SSB is enriched upstream of either leading or lagging strand nicks (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eA). SSB enrichment\u0026thinsp;~\u0026thinsp;3 kb upstream from the nick site was dependent upon AddAB but independent of the remaining factors required for homologous recombination (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eB), indicating that AddAB is necessary to generate ssDNA for SSB.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eSSB forms a homotetramer that binds ssDNA non-specifically via its N-terminal domain. Each SSB monomer harbours a long disordered C-terminal domain that contains a protein interaction sequence at the terminus (the C-terminal tail or CTT) (Fig. S9A-B)\u003csup\u003e\u003cspan citationid=\"CR59\" class=\"CitationRef\"\u003e57\u003c/span\u003e\u003c/sup\u003e. While the \u003cem\u003eB. subtilis\u003c/em\u003e SSB-CTT is dispensable for cell viability, it is required for efficient repair of DNA lesions created by either UV irradiation or Mitomycin C treatment \u003csup\u003e\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e42\u003c/span\u003e,\u003cspan citationid=\"CR60\" class=\"CitationRef\"\u003e58\u003c/span\u003e\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eTo determine whether the SSB-CTT is required for repair of a DSB following replisome inactivation at a nick, an IPTG-inducible SSB complementation system was generated in a Δ\u003cem\u003essbA\u003c/em\u003e strain, allowing expression of either the wild-type protein or a variant lacking the last six amino acids (SSB\u003csup\u003eΔCTT\u003c/sup\u003e, Fig. S9C). Spot-titre analyses confirmed that SSB\u003csup\u003eΔCTT\u003c/sup\u003e can complement a Δ\u003cem\u003essbA\u003c/em\u003e mutant similar to wild-type SSB and that viability was dependent upon IPTG (Fig. S9D), and immunoblots showed that the inducible SSB proteins were being expressed at levels comparable to the endogenous protein (Fig. S9E).\u003c/p\u003e \u003cp\u003eThe SSB complementation system was then moved into strains expressing nCas9 and sgRNA (Fig. S9F). Spot titre analyses showed that wild-type SSB was able to support viability following induction of nCas9 and sgRNA, whereas the SSB\u003csup\u003eΔCTT\u003c/sup\u003e variant could not (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eC). MFA confirmed that chromosome replication from the nick site to the terminus is arrested when SSB lacks its C-terminal tail (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eD). Therefore, the results indicate that the SSB-CTT is necessary for DNA repair following replisome inactivation at a nick.\u003c/p\u003e \u003cp\u003e \u003cb\u003eThe SSB-CTT is required to recruit RecO to the site of DNA repair during homologous recombination\u003c/b\u003e \u003c/p\u003e \u003cp\u003eThe SSB-CTT acts as a protein interaction hub\u003csup\u003e\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e42\u003c/span\u003e\u003c/sup\u003e that directly interacts with several factors found to be necessary for repair of a DSB, including RecO, RecG, and PriA (Figs. S9G, 2I-K, 3A-B). To investigate the essential role of the SSB-CTT, ChIP was employed to probe the recruitment of SSB variants, His-RecO, and His-RecA upstream of nick sites. ChIP showed that the SSB\u003csup\u003eΔCTT\u003c/sup\u003e variant remained enriched upstream of nicks at similar levels to the wild-type protein (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eE). However, enrichment in His-RecO and His-RecA was lost in cells harbouring SSB\u003csup\u003eΔCTT\u003c/sup\u003e (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eE), indicating that the SSB:RecO interaction depends on the SSB-CTT (Fig. S9H). In cells encoding wild-type SSB, enrichment of His-RecA was dependent upon both AddAB and RecFOR (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eF), consistent with the model that RecFOR is necessary for RecA loading onto ssDNA substrates bound by SSB. Taken together, these results indicate that an essential role of the SSB-CTT is to recruit RecO to repair sites following replisome inactivation at single-strand DNA discontinuities.\u003c/p\u003e \u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003ch2\u003eWhat is the essential role of RecG for restart of DNA replication?\u003c/h2\u003e \u003cp\u003eWhile the role of most factors required for resuming DNA replication following replisome inactivation at a single-strand discontinuity could be rationalised, the role of RecG was unclear. RecG has been implicated in a myriad of DNA transactions including reversal of replication forks, migration of Holliday junctions, regulation of RecA strand exchange, processing of flaps generated when DNA replication forks converge, and stabilisation of D-loops\u003csup\u003e\u003cspan citationid=\"CR58\" class=\"CitationRef\"\u003e56\u003c/span\u003e,\u003cspan additionalcitationids=\"CR60 CR61\" citationid=\"CR61\" class=\"CitationRef\"\u003e59\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e62\u003c/span\u003e\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eExamination of the MFA in a Δ\u003cem\u003erecG\u003c/em\u003e mutant showed that there was an increase in sequence read depth upstream of a nick targeted to either the leading or lagging strand template (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eA), suggesting that DNA synthesis increases in this region in the absence of RecG. This observation was reminiscent of results reported in \u003cem\u003eEscherichia coli\u003c/em\u003e during DSB repair in a Δ\u003cem\u003erecG\u003c/em\u003e mutant, where it was proposed that RecG remodels intermediate DNA structures formed during homologous recombination to ensure PriA is correctly orientated to load helicase (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eB, in the absence of RecG helicase reloading occurs on the opposite strand normally used for DNA replication, thus directing inappropriate DNA synthesis from the terminus towards \u003cem\u003eoriC\u003c/em\u003e)\u003csup\u003e\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e47\u003c/span\u003e\u003c/sup\u003e. This hypothesis diverges from the generally accepted model for homologous recombination, in which resection of a DSB involves degradation of the strand with a 5\u0026prime;-end, thereby generating a single-stranded 3\u0026prime;-end substrate for recombinases (e.g. RecA, Fig. S10). The revised model instead suggests that following DNA end-processing, the strand containing a 5\u0026prime;-end remains intact and can hybridise the displaced strand (D-loop) formed as a consequence of RecA strand invasion\u003csup\u003e\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e47\u003c/span\u003e\u003c/sup\u003e. It was suggested that this intermediate DNA structure was the substrate for RecG, such that the enzyme would remodel the strands by unwinding the annealed 5\u0026prime;-end until the annealed 3\u0026prime;-end abutted the fork junction, thus generating the correct substrate for PriA to reload helicase (Fig. S11)\u003csup\u003e\u003cspan citationid=\"CR65\" class=\"CitationRef\"\u003e63\u003c/span\u003e\u003c/sup\u003e. Biochemical data using purified proteins has shown that RecG and PriA can perform these activities on model replication fork substrates\u003csup\u003e\u003cspan citationid=\"CR65\" class=\"CitationRef\"\u003e63\u003c/span\u003e,\u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e64\u003c/span\u003e\u003c/sup\u003e. Therefore, we sought to test this hypothesis \u003cem\u003ein vivo\u003c/em\u003e using the nCas9 system.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eRecG is required to remodel replication forks formed when a DSB contains a 5′-end\u003c/h3\u003e\n\u003cp\u003eThe genetic analysis above indicated that the AddAB helicase/nuclease was responsible of end processing of the DSB (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA). While both AddA and AddB subunits harbour nuclease activity, the two enzymes cleave distinct strands of the DNA duplex following unwinding by AddA helicase activity (AddA cleaves the strand terminating with a 3\u0026prime;-end, AddB cleaves the strand terminating with a 5\u0026prime;-end)\u003csup\u003e\u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e49\u003c/span\u003e\u003c/sup\u003e. Therefore, it was hypothesised that inactivating either AddA or AddB nuclease activity would bias DSB processing to favour the generation of either a 3\u0026prime;-tail (AddA nuclease mutant) or a 5\u0026prime;-tail (AddB nuclease mutant).\u003c/p\u003e \u003cp\u003eTo begin, ChIP was used to confirm that AddAB was acting directly on the DSB generated following replisome inactivation at a nick. AddA and AddB were independently fused to the fluorescent protein \u003cem\u003emNeonGreen\u003c/em\u003e and enrichment of each subunit was determined. The results showed that mNG-AddA and mNG-AddB are both enriched upstream of nicks, consistent with their direct role in end-processing under these conditions (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eC).\u003c/p\u003e \u003cp\u003eNext, the enzymatic activities of AddA and AddB were reduced through substitution of catalytic residues\u003csup\u003e\u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e49\u003c/span\u003e\u003c/sup\u003e. Spot titre analyses showed that the helicase defective AddA\u003csup\u003eK\u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e36\u003c/span\u003eA\u003c/sup\u003e (AddA\u003csup\u003eΔHEL\u003c/sup\u003e) was unable to sustain bacterial growth upon induction of nicks, whereas nuclease defective variants of either AddA\u003csup\u003eD1172A\u003c/sup\u003e (AddA\u003csup\u003eΔNUC\u003c/sup\u003e) or AddB\u003csup\u003eD961A\u003c/sup\u003e (AddB\u003csup\u003eΔNUC\u003c/sup\u003e) grew similar to a strain with wild-type AddAB (Figs.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eD, S12A). Notably, the double nuclease variant AddA\u003csup\u003eΔNUC\u003c/sup\u003eAddB\u003csup\u003eΔNUC\u003c/sup\u003e also supported normal growth (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eD), suggesting that under these conditions in live cells the DSB may not require resection. MFA confirmed that DNA synthesis was blocked downstream of a nick in the strain expressing AddA\u003csup\u003eΔHEL\u003c/sup\u003e, while the strains expressing AddA\u003csup\u003eΔNUC\u003c/sup\u003e, AddB\u003csup\u003eΔNUC\u003c/sup\u003e or AddA\u003csup\u003eΔNUC\u003c/sup\u003eAddB\u003csup\u003eΔNUC\u003c/sup\u003e were comparable to wild-type (Figs.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eE, S12B-C). Moreover, ChIP demonstrated that SSB is not enriched upstream of nicks in a strain expressing AddA\u003csup\u003eΔHEL\u003c/sup\u003e, whereas SSB was enriched in AddA\u003csup\u003eΔNUC\u003c/sup\u003e and AddB\u003csup\u003eΔNUC\u003c/sup\u003e backgrounds (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eF). Together these results indicate that AddAB helicase activity is directly required to process a DSB generated after replisome inactivation at a nick, and that nuclease activity can be abolished without compromising DNA replication restart.\u003c/p\u003e \u003cp\u003eFinally, the Δ\u003cem\u003erecG\u003c/em\u003e mutant was moved into strains expressing either AddA\u003csup\u003eΔNUC\u003c/sup\u003e or AddB\u003csup\u003eΔNUC\u003c/sup\u003e and nicking was induced (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eA-B). Spot-titre assays showed that RecG remained essential in the strains containing AddB\u003csup\u003eΔNUC\u003c/sup\u003e, where degradation of the strand terminating with a 5\u0026prime;-end is inhibited (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eC). Strikingly, in the AddA\u003csup\u003eΔNUC\u003c/sup\u003e strain where degradation of the strand terminating with a 3\u0026prime;-end is inhibited, the absence of RecG was well tolerated (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eC-D). These results are consistent with the model that RecG is required to remodel recombination intermediates that contain a single-strand with a 5\u0026prime;-end annealed to a D-loop. Importantly, the critical role of RecG in strains with single-strand discontinuities (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA) implies that wild-type processing of a DSB often generates a single-strand with a 5\u0026prime;-end, which is inconsistent with most current models of bacterial homologous recombination.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e"},{"header":"DISCUSSION","content":"\u003cp\u003eIn this study we have characterised the fate of the bacterial replisome \u003cem\u003ein vivo\u003c/em\u003e after it encounters a single-strand discontinuity in the DNA template (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eA). The results indicate that a nick in the template for either the leading or lagging strand arrests DNA synthesis, generates a DSB, and elicits replication restart following repair of the DNA break through homologous recombination (Figs.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003e\u0026ndash;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003e). These findings are consistent with the observation that a nick in a replicating bacteriophage chromosome generates DSBs\u003csup\u003e\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e\u003c/sup\u003e and with recent reports of the impact of nicks in eukaryotic replication systems\u003csup\u003e\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e,\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e,\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e23\u003c/span\u003e\u003c/sup\u003e. Importantly, while replisomes are thought to be generally intolerant to many types of DNA damage\u003csup\u003e\u003cspan citationid=\"CR67\" class=\"CitationRef\"\u003e65\u003c/span\u003e\u003c/sup\u003e, it appears that the bacterial replication machinery cannot readily bypass single-strand interruptions as these lesions affect helicase progression (Figs.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e, \u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e(I)).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eRepair of the DSB to facilitate replication restart requires homologous recombination (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003e). Genetic analysis combined with ChIP indicates that a single pathway is utilised under these experimental conditions (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e): AddAB unwinds the DNA; SSB binds the ssDNA; RecFOR counteracts SSB and allows RecA to bind ssDNA; RecA promotes strand invasion of the homologous chromosome and generates a D-loop, which often anneals to a single-strand with a 5\u0026prime;-end; RecG regresses the fork and simultaneously separates the annealed strand with a 5\u0026prime;-end; PriA binds to the remodelled fork where it interacts with the strand harbouring a 3\u0026prime;-end and reloads helicase to promote replisome assembly and DNA replication restart. The observed lack of redundancy between proteins involved in recombinational DNA repair suggests that the pathway identified here is the major route for fixing this type of collapsed replication fork in \u003cem\u003eB. subtilis\u003c/em\u003e\u003csup\u003e\u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e41\u003c/span\u003e,\u003cspan citationid=\"CR68\" class=\"CitationRef\"\u003e66\u003c/span\u003e\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eBuilding upon these results, we investigated specific protein activities to help elucidate their roles during recombinational repair of a DSB. Below we discuss these findings in what we propose is chronological order.\u003c/p\u003e \u003cdiv id=\"Sec11\" class=\"Section2\"\u003e \u003ch2\u003eThe replicative helicase is inactivated after it encounters a nick\u003c/h2\u003e \u003cp\u003eIn bacteria the replicative helicase encircles and tracks along the template for lagging strand synthesis. ChIP data suggests that there are distinct fates of the helicase depending on the location of a nick (Figs.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA-B, S4). As expected for a nick in the lagging strand template, helicase enrichment was only observed at the cleavage site and upstream, consistent with the model that helicase runs-off the template when it encounters the single-strand discontinuity (Figs.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eG, \u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e(I)). In contrast, when the nick was in the leading strand template, helicase was also enriched downstream of the cleavage site, suggesting that the enzyme continued translocation. We propose that upon encountering a nick in the leading strand template, helicase slides onto the dsDNA template in an inactive conformation that cannot perform further DNA unwinding (Figs.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eG, \u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e(I)). This model is consistent with \u003cem\u003ein vitro\u003c/em\u003e experiments using \u003cem\u003eThermus aquaticus\u003c/em\u003e helicase, where after unwinding up to a nick on the leading strand template, the enzyme can continue to translocate with the two strands of DNA passing through the central channel with no resultant unwinding\u003csup\u003e\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e37\u003c/span\u003e\u003c/sup\u003e. Whether the bacterial helicase is actively removed from dsDNA under these conditions, akin to what has been observed for the eukayotic CMG helicase in \u003cem\u003eXenopus\u003c/em\u003e extracts\u003csup\u003e\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e\u003c/sup\u003e, requires further investigation.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec12\" class=\"Section2\"\u003e \u003ch2\u003eAddAB helicase activity is necessary and sufficient for recombinational repair\u003c/h2\u003e \u003cp\u003eAddAB and related enzymes have been shown to bind blunt end dsDNA or dsDNA with short overhangs (4\u0026ndash;20 nucleotides with either a 5\u0026prime;- or a 3\u0026prime;-end)\u003csup\u003e\u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e49\u003c/span\u003e,\u003cspan citationid=\"CR69\" class=\"CitationRef\"\u003e67\u003c/span\u003e\u003c/sup\u003e. When the template for leading strand synthesis is nicked, the DNA polymerase likely runs-off the end of the substrate to produce a blunt or nearly blunt end, suitable for recognition by AddAB. However, when the template for lagging strand synthesis is nicked, a substrate with a 3\u0026prime;-tail would likely be produced. Because the reverse genetic screen suggested that no additional nucleases are required for this recombinational repair pathway (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003e, S8), we speculate that the high local concentration of replication factors would facilitate priming near the end of the 3\u0026prime;-tail, thereby filling the ssDNA with an Okazaki fragment to create a suitable substrate for AddAB.\u003c/p\u003e \u003cp\u003eThe master recombination factor RecA requires ssDNA to promote homologous pairing. A long-standing question is how this ssDNA is generated \u003cem\u003ein vivo\u003c/em\u003e. Most current models suggest that enzymes resect a DSB to generate a single-strand terminating with a 3\u0026prime;-end\u003csup\u003e\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e,\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e31\u003c/span\u003e\u003c/sup\u003e. Here we tested this model by mutating the nuclease activities of AddA and AddB. The results showed that neither nuclease activity is required for enzyme function under the experimental conditions tested (Figs.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eD-E, S12B-C). In contrast, AddAB helicase activity was required for recombinational repair, consistent with a previous study of the ancestral AdnAB resection machinery in \u003cem\u003eMycobacteria tuberculosis\u003c/em\u003e\u003csup\u003e\u003cspan citationid=\"CR70\" class=\"CitationRef\"\u003e68\u003c/span\u003e\u003c/sup\u003e. These results suggest that the key role of AddAB is not to digest the dsDNA ends, but rather to separate the strands and create the ssDNA substrate for SSB (Figs.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eF, \u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e(II-III)).\u003c/p\u003e \u003cp\u003eThe local concentration of SSB is high at the site of DNA replication\u003csup\u003e\u003cspan citationid=\"CR71\" class=\"CitationRef\"\u003e69\u003c/span\u003e,\u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e70\u003c/span\u003e\u003c/sup\u003e, potentially facilitating its rapid association with the ssDNA emerging from AddAB. In fact, \u003cem\u003ein vitro\u003c/em\u003e the AddA\u003csup\u003eNUC\u003c/sup\u003eAddB\u003csup\u003eNUC\u003c/sup\u003e nuclease-dead enzyme does not unwind dsDNA in the absence of SSB\u003csup\u003e\u003cspan citationid=\"CR73\" class=\"CitationRef\"\u003e71\u003c/span\u003e\u003c/sup\u003e, suggesting that AddAB unwinding and SSB binding ssDNA may be coupled. We propose that AddAB nuclease activity represents a secondary method of generating a ssDNA substrate for SSB, following initial AddAB helicase activity required to separate the DNA duplex.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec13\" class=\"Section2\"\u003e \u003ch2\u003eInteraction of the SSB C-terminal tail with RecO is necessary for replication repair\u003c/h2\u003e \u003cp\u003eThe SSB-CTT is an established protein interaction hub that has been shown to directly bind several of the essential replication and repair factors required at nicks, including RecO\u003csup\u003e\u003cspan additionalcitationids=\"CR73 CR74\" citationid=\"CR74\" class=\"CitationRef\"\u003e72\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR77\" class=\"CitationRef\"\u003e75\u003c/span\u003e\u003c/sup\u003e, RecG\u003csup\u003e\u003cspan citationid=\"CR78\" class=\"CitationRef\"\u003e76\u003c/span\u003e,\u003cspan citationid=\"CR79\" class=\"CitationRef\"\u003e77\u003c/span\u003e\u003c/sup\u003e, and PriA\u003csup\u003e\u003cspan citationid=\"CR60\" class=\"CitationRef\"\u003e58\u003c/span\u003e,\u003cspan citationid=\"CR80\" class=\"CitationRef\"\u003e78\u003c/span\u003e\u003c/sup\u003e. However, the critical activities performed by the SSB-CTT \u003cem\u003ein vivo\u003c/em\u003e have not yet been established\u003csup\u003e\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e42\u003c/span\u003e\u003c/sup\u003e. Based on the results above, we conclude that the SSB-CTT interaction with RecO is necessary for replication repair. RecO is part of the RecFOR system required for RecA loading onto ssDNA \u003cem\u003ein vitro\u003c/em\u003e\u003csup\u003e\u003cspan citationid=\"CR81\" class=\"CitationRef\"\u003e79\u003c/span\u003e\u003c/sup\u003e and RecA-GFP foci formation \u003cem\u003ein vivo\u003c/em\u003e\u003csup\u003e\u003cspan citationid=\"CR82\" class=\"CitationRef\"\u003e80\u003c/span\u003e\u003c/sup\u003e. Because RecO activity occurs before the activities of RecG and PriA (i.e. RecA binding ssDNA is blocked, Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eF), this precludes further conclusions regarding the importance of these protein:protein interactions at this time.\u003c/p\u003e \u003cp\u003e \u003cem\u003eIn vitro\u003c/em\u003e the RecO:RecR complex facilitates loading of RecA onto SSB-coated ssDNA\u003csup\u003e\u003cspan citationid=\"CR83\" class=\"CitationRef\"\u003e81\u003c/span\u003e,\u003cspan citationid=\"CR84\" class=\"CitationRef\"\u003e82\u003c/span\u003e\u003c/sup\u003e. Because RecO, RecR, and RecF were all required for replication repair, we propose that the essential role of RecF here is to recruit RecR and generate the RecO:RecR complex. This model is consistent with a hand-off mechanism for RecR observed \u003cem\u003ein vitro\u003c/em\u003e\u003csup\u003e\u003cspan citationid=\"CR83\" class=\"CitationRef\"\u003e81\u003c/span\u003e\u003c/sup\u003e. RecF is a dimeric ATPase that adopts a clamp conformation around dsDNA\u003csup\u003e\u003cspan citationid=\"CR53\" class=\"CitationRef\"\u003e51\u003c/span\u003e\u003c/sup\u003e. Recent studies in \u003cem\u003eE. coli\u003c/em\u003e have shown that recruitment of RecF to sites of repair is mediated by the DNA polymerase sliding clamp processivity factor DnaN\u003csup\u003e\u003cspan citationid=\"CR84\" class=\"CitationRef\"\u003e82\u003c/span\u003e\u003c/sup\u003e. As was the case for SSB\u003csup\u003e\u003cspan citationid=\"CR72\" class=\"CitationRef\"\u003e70\u003c/span\u003e\u003c/sup\u003e, the local concentration of DnaN is high near the sites of DNA replication collapse\u003csup\u003e\u003cspan citationid=\"CR71\" class=\"CitationRef\"\u003e69\u003c/span\u003e\u003c/sup\u003e, thereby potentially facilitating RecF recruitment. Importantly for this model, since both SSB and DnaN are enriched at a replication fork, we predict that there is a mechanism to regulate RecO:RecR complex formation until it is required, thus safeguarding the SSB-coated strand from promiscuous RecA loading (Figs.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eB, \u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eF, \u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e(III-V)).\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003ch2\u003eRecG remodels a repaired replication fork by unwinding dsDNA with an exposed 5\u0026prime;-end\u003c/h2\u003e \u003cp\u003eThe biological role of RecG has been a long-standing question\u003csup\u003e\u003cspan citationid=\"CR57\" class=\"CitationRef\"\u003e55\u003c/span\u003e,\u003cspan citationid=\"CR58\" class=\"CitationRef\"\u003e56\u003c/span\u003e\u003c/sup\u003e. \u003cem\u003eIn vitro\u003c/em\u003e RecG is a DNA translocase that unwinds a myriad of DNA substrates, including Holliday junctions, D-loops, and model replication forks\u003csup\u003e\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e47\u003c/span\u003e,\u003cspan additionalcitationids=\"CR61\" citationid=\"CR62\" class=\"CitationRef\"\u003e60\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR64\" class=\"CitationRef\"\u003e62\u003c/span\u003e,\u003cspan citationid=\"CR66\" class=\"CitationRef\"\u003e64\u003c/span\u003e\u003c/sup\u003e. Here MFA indicates that a \u003cem\u003eΔrecG\u003c/em\u003e mutant cannot efficiently restart DNA replication at a nick directed to either the leading or the lagging strand template (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB, \u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eA-B), consistent with a role in fork remodelling\u003csup\u003e\u003cspan citationid=\"CR48\" class=\"CitationRef\"\u003e46\u003c/span\u003e\u003c/sup\u003e. Based on genetic experiments and supported by biochemical assays \u003cem\u003ein vitro\u003c/em\u003e, a model has been presented for RecG binding to a replication fork and unwinding dsDNA on the lagging strand template to allow correct orientation of PriA for helicase loading\u003csup\u003e\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e47\u003c/span\u003e,\u003cspan citationid=\"CR65\" class=\"CitationRef\"\u003e63\u003c/span\u003e\u003c/sup\u003e. Critically, this model requires that the single strand with a 5\u0026prime;-end from the DSB can hybridise with ssDNA of the D-loop on the homologous chromosome.\u003c/p\u003e \u003cp\u003eHow this predicted intermediate structure is constructed remains to be determined, but the implication is that the separated strands of the DSB are either the same length or the strand with a 5\u0026prime;-end is longer (Figs.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eA, \u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eD, \u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e(VI-VII)). This model contrasts with most current views of bacterial DSB end-processing where a single strand with a 3\u0026prime;-end is produced for RecA (Fig. S10)\u003csup\u003e\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e,\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e31\u003c/span\u003e,\u003cspan citationid=\"CR85\" class=\"CitationRef\"\u003e83\u003c/span\u003e\u003c/sup\u003e. Moreover, because RecG is essential in strains expressing Cas9 nickases, it suggests that during wild-type DSB end-processing the separated strands often contain a 5\u0026prime;-end that requires remodelling by RecG (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e(VII)). These unanticipated results, together with those of the accompanying manuscript analysing resolution of collapsed eukaryotic replication forks\u003csup\u003e\u003cspan citationid=\"CR86\" class=\"CitationRef\"\u003e84\u003c/span\u003e\u003c/sup\u003e, necessitate further evaluation of DSB management following replisome collapse across all domains of life.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec15\" class=\"Section2\"\u003e \u003ch2\u003eHolliday junction migration and resolution are not necessary for replication restart\u003c/h2\u003e \u003cp\u003eBranch migration of a Holliday junction is generally considered a hallmark of homologous recombination, therefore it was notable that the canonical Holliday junction translocase RuvAB was not required for replication repair at nicks (Fig. S8D). In fact, the \u003cem\u003eΔrecU ΔruvAB\u003c/em\u003e knockout attenuated the penetrance of the \u003cem\u003eΔrecU\u003c/em\u003e mutant upon introduction of single-strand discontinuities, suggesting that either another enzyme can functionally complement RuvAB activities or that branch migration is not required for this repair pathway.\u003c/p\u003e \u003cp\u003eWhile genetic analysis shows that RecU is required for wild-type growth following expression of Cas9 nickases, MFA indicates that RecU is not necessary for replication restart \u003cem\u003eper se\u003c/em\u003e (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003e). These results are consistent with the model that RecU resolves the Holliday junction formed during recombinational repair\u003csup\u003e\u003cspan citationid=\"CR87\" class=\"CitationRef\"\u003e85\u003c/span\u003e\u003c/sup\u003e. We hypothesise that the growth defect observed in the \u003cem\u003eΔrecU\u003c/em\u003e mutant may be caused by aberrant chromosome segregation\u003csup\u003e\u003cspan citationid=\"CR87\" class=\"CitationRef\"\u003e85\u003c/span\u003e\u003c/sup\u003e, as the recombined DNA molecules need to be separated prior to completing cell division. It is known that multiple systems can facilitate chromosome separation following replication (e.g. Topoisomerase IV, XerD)\u003csup\u003e\u003cspan citationid=\"CR88\" class=\"CitationRef\"\u003e86\u003c/span\u003e,\u003cspan citationid=\"CR89\" class=\"CitationRef\"\u003e87\u003c/span\u003e\u003c/sup\u003e, potentially explaining why the growth defect of the \u003cem\u003eΔrecU\u003c/em\u003e mutant under these experimental conditions was less severe compared to the other recombination mutants identified.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec16\" class=\"Section2\"\u003e \u003ch2\u003ePotentiating ROS-mediated antibiotic toxicity by perturbing DNA repair\u003c/h2\u003e \u003cp\u003eIn this work, the nicks generated by Cas9 enzymes were used as proxies for single-strand discontinuities formed during removal of damaged DNA from a bacterial genome. We find that when a bacterial replication fork encounters a single strand discontinuity it becomes inactivated, likely contributing to the toxic effect of ROS-dependent damage induced by antimicrobials. By identifying the essential factors and activities required to repair the DSB formed after a replisome encounters a nick, we have provided a set of molecular targets whose disruption could potentiate the activity of antibiotics that produce ROS-dependent DNA damage.\u003c/p\u003e \u003c/div\u003e"},{"header":"Methods","content":"\u003cp\u003e\u003cem\u003eExperimental model and growth\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003eThe following bacterial organisms were used: \u003cem\u003eE. coli \u003c/em\u003eand\u003cem\u003e B. subtilis. \u003c/em\u003eUnless otherwise stated in method details, strains were grown at 37\u0026deg;C using nutrient agar (NA) or lysogeny broth (LB: 10 g/L tryptone, 10 g/L NaCl, 5 g/L yeast extract) for routine selection and maintenance of bacterial strains. Supplements were added as required: ampicillin (100 \u0026micro;g/ml), erythromycin (1 \u0026micro;g/ml) in conjunction with lincomycin (25 \u0026mu;g/ml), chloramphenicol (5 \u0026micro;g/ml), kanamycin (5 \u0026micro;g/ml), spectinomycin (50 \u0026micro;g/ml), zeocin (10 \u0026mu;g/ml), xylose (1% w/v) and IPTG (0.05 mM, 0.1 mM or 1 mM).\u003c/p\u003e\n\u003cp\u003e\u003cem\u003e \u003c/em\u003e\u003cem\u003eBacterial strains\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003eBacterial strains used in this study are listed in Table S2.\u003c/p\u003e\n\u003cp\u003e\u003cem\u003e \u003c/em\u003e\u003cem\u003eE. coli plasmid construction\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003e\u003cem\u003eE. coli \u003c/em\u003etransformations for constructs harbouring \u003cem\u003epriA\u003c/em\u003e were performed in CW198 via heat-shock following the Hanahan method\u003csup\u003e88\u003c/sup\u003e and propagated in LB with appropriate antibiotics at 37\u0026deg;C. DH5\u0026alpha; [\u003cem\u003eF- \u0026Phi;80lacZ\u0026Delta;M15 \u0026Delta;(lacZYA-argF) U169 recA1 endA1 hsdR17(rk-, mk+) phoA supE44 thi-1 gyrA96 relA1 \u0026lambda;-\u003c/em\u003e] was used for other plasmids construction. Plasmids were purified using QIAprep spin miniprep kits and recombinant DNA confirmed by sequencing (Table S3). Descriptions, where necessary, are provided below\u003cem\u003e.\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003epCW367, pCW368, pCW478, pCW479, pCW510, pCW717, pCW734, pCW736 and pCW850 were generated by Quickchange mutagenesis using mutagenic primer pairs containing an overlap of 10-15 bp. PCR products were treated with DpnI for at least 3 hours to digest template DNA and 10 \u0026mu;l were used for \u003cem\u003eE. coli\u003c/em\u003e transformation.\u003c/p\u003e\n\u003cp\u003epCW300, pCW301, pCW327, pCW460, pCW471, pCW472, pCW575, pCW734, pCW736 and pCW766 were generated by Gibson assembly using the NEB Hi-Fi cloning kit according to manufacturer instructions following purification of individual PCR fragments using QIAquick PCR purification kits.\u003c/p\u003e\n\u003cp\u003e\u003cem\u003eLinear recombinant DNA assembly\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003eFor recombinant DNA\u003cem\u003e \u003c/em\u003eassembly without \u003cem\u003eE. coli\u003c/em\u003e propagation, pCW573, pCW574, pCW577, pCW578, pCW605, pCW640, pCW672, pCW702, pCW720, pCW732, pCW758, pCW759, pCW760, pCW777, pCW779, pCW781, pCW783, pCW785, pCW802, pCW803, pCW804, pCW809, pCW834, pCW848, pCW854, pCW855, pCW857 and pCW859 were assembled linearly using a multistep \u003cem\u003ein vitro\u003c/em\u003e assembly process followed by nested PCR amplification prior to \u003cem\u003eB. subtilis\u003c/em\u003e transformation. Briefly, individual PCR fragments corresponding to homology regions and insert DNA were amplified, purified and DNA ends ligated using the NEB Hi-Fi cloning kit. Raw Gibson assembly products were then amplified using nested primer pairs annealing towards the end of homology regions and PCR products were used for \u003cem\u003eB. subtilis\u003c/em\u003e transformation. Following isolation of \u003cem\u003eB. subtilis\u003c/em\u003e colonies resistant to the appropriate antibiotic markers, genomic DNA was extracted using the DNeasy blood and tissue kit, regions corresponding to recombinant DNA were amplified via PCR using oligonucleotides annealing outside homology fragments to ensure double crossover had occurred and results were confirmed via PCR product sequencing. Note that CW2045 and derivatives harbour the additional \u003cem\u003eaddA\u003csup\u003eD682E\u003c/sup\u003e\u003c/em\u003e mutation necessary to propagate \u003cem\u003eaddB\u003csup\u003eD961A\u003c/sup\u003e-addA\u003csup\u003eD1172A\u003c/sup\u003e\u003c/em\u003e in live cells, which is surface exposed and structurally unlikely to impact AddAB end-processing activities (Fig. S12E).\u003c/p\u003e\n\u003cp\u003e\u003cem\u003eB. subtilis strain construction\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003eTransformation of competent \u003cem\u003eB. subtilis\u003c/em\u003e cells was performed using an optimized two-step starvation procedure as previously described\u003csup\u003e89\u003c/sup\u003e. Briefly, recipient strains were grown overnight at 37\u0026deg;C in transformation medium (Spizizen salts: 0.2% w/v ammonium sulphate, 1.4% w/v dipotassium phosphate, 0.6% w/v potassium phosphate, 0.1% w/v tri-sodium citrate, 0.02% w/v magnesium sulphate supplemented with 1 \u0026mu;g/ml ammonium iron(III) citrate, 6 mM MgSO\u003csub\u003e4\u003c/sub\u003e, 0.5% w/v glucose, 0.02 mg/ml tryptophan and 0.02% w/v casein peptone) supplemented with IPTG where required. Overnight cultures were diluted 1:20 into fresh transformation medium supplemented with IPTG where required and grown at 37\u0026deg;C for 2.5-3 hours with continual shaking. An equal volume of prewarmed starvation medium (Spizizen salts supplemented with 6 mM MgSO\u003csub\u003e4\u003c/sub\u003e and 0.5% w/v glucose) was added and the culture was incubated at 37\u0026deg;C for two hours with continual shaking. DNA was added to 350 \u0026mu;l cells and the mixture was incubated at 37\u0026deg;C for one hour with continual shaking. 20-200 \u0026mu;l of each transformation was plated onto selective media supplemented with IPTG where required and incubated at 37\u0026deg;C for 24-48 hours. Results were validated by propagating cells on relevant antibiotic markers and sequencing.\u003c/p\u003e\n\u003cp\u003e\u003cem\u003eSpot titre assays\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003eCells were grown in LB overnight in the presence of 0.5% w/v glucose. Overnight cultures were diluted in fresh LB medium using 10-fold serial dilutions and 5 \u0026mu;L aliquots were spotted onto NA plates with or without xylose. NA plates were supplemented with IPTG when required as indicated on individual figure panels. All plates were incubated for 24 hours or up to 48 hours when indicated. Experiments were performed independently at least three times and representative data are shown.\u003c/p\u003e\n\u003cp\u003e\u003cem\u003eImmunoblot analyses\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003eCultures were grown overnight in LB and diluted 1:100 in fresh medium the next morning. Diluted cells were grown to an absorbance at 600 nm (A\u003csub\u003e600\u003c/sub\u003e) of 0.2-0.3, induced with 1% w/v xylose or 1 mM IPTG where required and further incubated for one hour. Cells were harvested using centrifugation, resuspended in 1x PBS supplemented with one tenth of a peptidase inhibitor tablet and sonicated (40 amp) twice for 12 seconds with 3 second pulses at 4\u0026deg;C. Samples were adjusted to 1X LDS buffer and 125 mM DTT, fixed for 5 minutes at 95\u0026deg;C and centrifuged for 2 minutes prior to loading the supernatant on polyacrylamide gels. Proteins were separated by electrophoresis using NuPAGE 4-12% Bis-Tris gradient gels run in MES SDS buffer and transferred to methanol-activated Hybond-P PVDF membranes using Wypall X60 cloths for blotting in a semi-dry apparatus (Bio-rad Trans-Blot Turbo, transfer buffer: 300 mM Tris-HCl, 300 mM glycine, 140 mM tricine, 0.05% w/v SDS, 2.5 mM EDTA). Membranes were washed twice using PBST (1X PBS supplemented with 0.1% v/v Tween-20) prior to blocking for 90 minutes at room temperature (blocking buffer: PBST supplemented with 7.5% w/v milk). Individual polypeptides were probed using the following primary antibodies diluted in blocking buffer following overnight incubation at 4\u0026deg;C: anti-Cas9 (1:1000), anti-FtsZ (1:5000), anti-PriA (1:1000), anti-SSB (1:1000) and anti-mNeonGreen (1:1000). Detection was performed with anti-rabbit (PriA, SSB), anti-sheep (FtsZ) or anti-mouse (Cas9, mNeonGreen) horseradish peroxidase-linked secondary antibodies in combination with the Clarity ECL substrate using an ImageQuant LAS 4000 mini digital imaging system. Detection of Cas9, FtsZ, PriA, SSB and mNeonGreen was within a linear range. Experiments were independently performed at least twice and representative data are shown.\u003c/p\u003e\n\u003cp\u003e\u003cem\u003eWhole genome sequencing\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003eCells were grown in rich chemically defined medium (RCDM: Spizizen salts supplemented with 1 \u0026mu;g/ml ammonium iron(III) citrate, 0.1 mM CaCl\u003csub\u003e2\u003c/sub\u003e, 0.13 mM MnSO\u003csub\u003e4\u003c/sub\u003e, 6 mM MgSO\u003csub\u003e4\u003c/sub\u003e, 0.5% w/v glucose, 0.1% w/v glutamate, 0.02 mg/ml tryptophan and 0.02% w/v casein peptone) overnight and diluted 1:100 in fresh RCDM the next morning. Diluted cells were grown to an absorbance at 600 nm (A\u003csub\u003e600\u003c/sub\u003e) of 0.2-0.3, induced with 1% w/v xylose and further incubated for one hour. Growth was arrested by adding 0.05% w/v sodium azide to cultures followed by vigorous mixing, cells were collected by centrifugation, the supernatant discarded, and pellets were flash frozen in liquid nitrogen before genomic DNA extraction using the DNeasy blood and tissue kit and a final elution volume of 50 \u0026micro;l in ultrapure water. Individual gDNA libraries were prepared using the DNA Prep tagmentation kit and pre-paired i5/i7 indices prior to pooling and DNA quantification with the 1X dsDNA high sensitivity kit using a Qubit instrument. Pooled libraries were diluted to 100 pM in resuspension buffer and 20 \u0026micro;l were loaded onto iSeq 100 i1 Reagent v2 sequencing chips for whole-genome sequencing using an iSeq100 sequencer.\u003c/p\u003e\n\u003cp\u003eFollowing whole-genome sequencing and demultiplexing embedded within the iSeq100 indexed sequencing workflow, .FASTQ files were extracted and loaded onto the CLC Genomics Workbench for initial data processing. Paired .FASTQ files were mapped onto the \u003cem\u003eB. subtilis\u003c/em\u003e \u0026Delta;6 reference genome using the \u0026lsquo;Map Reads to Reference\u0026rsquo; function and a .BAM file was created from mapped sequencing reads for each sequenced genome. An in-house script written in R using the Rsamtools library was employed to deconvolute .BAM files to validate even read distribution for sense/antisense strands prior to calculating density coverage and extracting cumulated densities per chromosome position.\u003c/p\u003e\n\u003cp\u003e\u003cem\u003eFluorescence microscopy\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003eStrains were grown overnight at 37\u0026deg;C in RCDM and the following day, cultures were diluted 1:100 into fresh imaging medium (RCDM where glucose is replaced by 0.5% v/v glycerol) in the presence of 0.1 mM IPTG for induction of mNeonGreen-RecA expression where required. Cells were allowed to grow to an A\u003csub\u003e600\u003c/sub\u003e of 0.3, induced with 1% w/v xylose and further incubated for 90 minutes. DAPI (5 \u0026micro;g/ml) and Nile red (1 \u0026micro;g/ml) stains were used to visualise the nucleoid and cell membrane following a 10 minute incubation on a benchtop incubator prior to imaging.\u003c/p\u003e\n\u003cp\u003eCells were mounted on ~1.25% agar pads (in sterile ultrapure water) and a 0.13- to 0.17-mm glass coverslip was placed on top. Microscopy was performed on an inverted epifluorescence microscope (Nikon Ti) fitted with a Plan Apochromat Objective 100x/1.40 NA Oil Ph3. Light was transmitted from a CoolLED pE-300 white light source through a Sutter Instruments liquid light guide and images were collected using a Photometrics Prime camera. Chroma fluorescence filter sets were used with 100 ms GFP, 250 ms DAPI and 50 ms mCherry exposure times at 100% LED power. Digital images were acquired using NIS Elements and analysed via the Fiji software\u003csup\u003e90\u003c/sup\u003e. Quantification of mNeonGreen-RecA features was manually curated from the count of 100 individual cells. All experiments were independently performed at least twice and representative data are shown.\u003c/p\u003e\n\u003cp\u003e\u003cem\u003eChromatin immunoprecipitation\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003eChromatin immunoprecipitation (ChIP) was performed as previously described with minor modifications\u003csup\u003e32\u003c/sup\u003e. Strains were grown overnight at 37\u0026deg;C in RCDM and the following day, cultures were diluted 1:100 into fresh medium and allowed to grow to an A\u003csub\u003e600\u003c/sub\u003e of 0.2-0.3. Cells were induced with 1% w/v xylose and further incubated for one hour prior to ChIP. Samples were adjusted to 1X PBS and crosslinked with 1% v/v formaldehyde for 10 minutes on a roller at room temperature, then quenched with 0.1 M glycine. Cells were pelleted at 4\u0026deg;C, washed three times with ice-cold 1X PBS (pH 7.3), frozen in liquid nitrogen and stored at -80\u0026deg;C. Frozen cell pellets were resuspended in 500 \u0026micro;l of lysis buffer (50 mM NaCl, 10 mM Tris-HCl pH 8.0, 20% w/v sucrose, 10 mM EDTA, 100 \u0026micro;g/ml RNase A, one quarter of a peptidase inhibitor tablet, 4 mg/ml lysozyme) and incubated at 37\u0026deg;C for 30 min to degrade the cell wall. Protoplasts were supplemented with 500 \u0026micro;l of immunoprecipitation buffer (300 mM NaCl, 100 mM Tris-HCl pH 7.0, 2% v/v Triton X-100, one quarter of a peptidase inhibitor tablet) to lyse cells and the mixture was incubated at 37\u0026deg;C for a further 10 minutes before cooling on ice for 5 minutes. Lysis and immunoprecipitation buffer volumes were multiplied by the number of antibodies to probe per sample. DNA samples were sonicated (40 amp) three times for 2 minutes at 4\u0026deg;C to obtain an average fragment size of 500 to 1000 base pairs. Cell debris were removed by centrifugation at 4\u0026deg;C and the supernatant transferred to a fresh Eppendorf tube. To determine the relative amount of DNA immunoprecipitated compared to the total amount of DNA, 100 \u0026micro;l of supernatant was removed, treated with 0.5 mg/ml pronase at 37\u0026deg;C for four hours then stored on ice. To immunoprecipate protein-DNA complexes, 800 \u0026micro;l of the remaining supernatant was incubated with individual antibodies (2 \u0026micro;l anti-SSB, 2 \u0026micro;l anti-PriA, 2 \u0026micro;l anti-DnaD, 2 \u0026micro;l anti-DnaB, 2 \u0026micro;l anti-DnaI, 2 \u0026micro;l anti-DnaC, 2 \u0026micro;l anti-mNeonGreen or 4 \u0026micro;l anti-His) for 90 minutes at room temperature. 750 \u0026micro;g of protein G Dynabeads were equilibrated by washing with bead buffer (1X PBS, 0.01% v/v Tween 20), resuspended in 25 \u0026micro;l of bead buffer and incubated with the sample supernatant for 1 hour at room temperature. Immunoprecipitated complexes were collected by applying the mixture to a magnet and washed with the following buffers in the respective order: 0.5X immunoprecipitation buffer for 15 min, 0.5X immunoprecipitation buffer supplemented with 500 mM NaCl for 15 min, stringent wash buffer (250 mM LiCl, 10 mM Tris-HCl pH 8.0, 0.5% v/v Igepal, 0.5% w/v sodium deoxycholate, 10 mM EDTA) for 20 min. Finally, protein-DNA complexes were washed a further three times with TET buffer (10 mM Tris-HCl pH 8.0, 1 mM EDTA, 0.01% v/v Tween 20) and resuspended in 100 \u0026micro;l of TEN buffer (10 mM Tris-HCl pH 8.0, 1 mM EDTA, 300 mM NaCl). Formaldehyde crosslinks of both the immunoprecipitate and total DNA were reversed by incubation at 65\u0026deg;C for 18 hours in the presence of 1 mg/ml proteinase K. DNA was then removed from the magnetic beads, cleaned using QIAquick PCR purification columns and used for quantitative PCR analyses.\u003c/p\u003e\n\u003cp\u003e\u003cem\u003eQuantitative PCR\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003eQuantitative PCR (qPCR) was performed using the Luna qPCR master mix to measure relative amounts of DNA bound by SSB, PriA, DnaD, DnaB, DnaI, DnaC, mNeonGreen-AddA, mNeonGreen-AddB, His-RecO and His-RecA at specific genomic locations (e.g. near the nick site located at -90\u0026deg; on the chromosome or the origin \u003cem\u003eoriC\u003c/em\u003e compared to a control site located at +90\u0026deg; on the chromosome). All PCR reactions were assembled using the QIAgility robotic workstation in 20 \u0026mu;l reaction volumes in a Rotor-Disc 100 and qPCRs were run on a Rotor-Gene Q instruments. Standard curves were obtained using the Rotor-Gene Q software to calculate the efficiency of each primer pair, which varied \u0026sim;5% between sets. Oligonucleotides were designed to amplify specific genomic regions\u003cem\u003e \u003c/em\u003eusing the Primer3Plus tool\u003csup\u003e91\u003c/sup\u003e, were typically 20-25 bases in length (Table S4) and amplified a ~100 bp PCR product. Individual fold enrichment ratios were obtained as follows: first, every Ct value was converted to 1/2\u003csup\u003eCt\u003c/sup\u003e and technical triplicates were averaged to generate a single enrichment value; second, genomic location specific enrichment was normalised by corresponding values obtained at +90\u0026deg; on the chromosome. Error bars indicate the standard error of the mean for three biological replicates.\u003c/p\u003e\n\u003cp\u003e\u003cem\u003eProtein structure prediction and representation\u003c/em\u003e\u003c/p\u003e\n\u003cp\u003eProtein models for SSB:ssDNA and SSB:RecO were generated using AlphaFold 3 and polypeptides corresponding to individual protein entities as input\u003csup\u003e92\u003c/sup\u003e. Main and alternative models were manually examined and Model 0 was chosen to highlight key features using the Pymol Molecular Graphics 2.1 software\u003csup\u003e93\u003c/sup\u003e. Structural highlights of the AddAB complex were derived from the crystal structure PDB: 4CEH\u003csup\u003e94\u003c/sup\u003e.\u003cbr\u003e \u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eDATA AVAILABILITY\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eLead contact\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eFor additional information and requests for resources and reagents, please contact the lead contact: Charles Winterhalter ([email protected]).\u003c/p\u003e\n\u003cp\u003e\u0026nbsp;\u003cstrong\u003eMaterials availability\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eGenetic materials generated in this study can be obtained directly from the lead contact.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eData and code availability\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eR code (DOI: \u0026nbsp;10.5281/zenodo.14793122) and protein structures for SSB:ssDNA and SSB:RecO complexes (DOI: 10.5281/zenodo.14793919) are available from Zenodo.\u003c/p\u003e\n\u003cp\u003e\u0026nbsp;\u003cstrong\u003eACKNOWLEDGEMENTS\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis work was supported by a Wellcome Trust Early-Career Award [226338/Z/22/Z] to CW and Wellcome Trust Discovery Award [225811/Z/22/Z] to HM. We would also like to express our gratitude to\u0026nbsp;Frances Davison for technical assistance, James Grimshaw for microscope maintenance and Frederic Schramm for insightful discussions prior to submission.\u003c/p\u003e\n\u003cp\u003e\u0026nbsp;\u003cstrong\u003eAUTHOR CONTRIBUTIONS\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eCW and HM contributed to the conception/design of the work. CW and SF generated results presented in the manuscript. CW created figures. CW and HM wrote/edited the manuscript.\u003c/p\u003e\n\u003cp\u003e\u0026nbsp;\u003cstrong\u003eETHICS DECLARATION\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare no competing interests.\u003c/p\u003e\n\u003cp\u003e\u0026nbsp;\u003cstrong\u003eDECLARATION OF GENERATIVE AI AND AI-ASSISTED TECHNOLOGIES\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eDuring the preparation of this work, the authors used AlphaFold 3 to generate SSB:ssDNA and SSB:RecO protein models. After using this tool, the authors reviewed the models and edited their content as needed and take full responsibility for the content of the publication.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n\u003cli\u003eMurray, C.J.L., Ikuta, K.S., Sharara, F., Swetschinski, L., Aguilar, G.R., Gray, A., Han, C., Bisignano, C., Rao, P., Wool, E., et al. (2022). Global burden of bacterial antimicrobial resistance in 2019: a systematic analysis. Lancet \u003cem\u003e399\u003c/em\u003e, 629-655. 10.1016/S0140-6736(21)02724-0.\u003c/li\u003e\n\u003cli\u003eButler, M.S., Vollmer, W., Goodall, E.C.A., Capon, R.J., Henderson, I.R., and Blaskovich, M.A.T. (2024). A Review of Antibacterial Candidates with New Modes of Action. Acs Infect Dis \u003cem\u003e10\u003c/em\u003e, 3440-3474. 10.1021/acsinfecdis.4c00218.\u003c/li\u003e\n\u003cli\u003eDwyer, D.J., Kohanski, M.A., and Collins, J.J. (2009). Role of reactive oxygen species in antibiotic action and resistance. 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Nature \u003cem\u003e508\u003c/em\u003e, 416-419. 10.1038/nature13037.\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":true,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":true,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"nature-portfolio","isNatureJournal":true,"hasQc":false,"allowDirectSubmit":false,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"","title":"Nature Portfolio","twitterHandle":"","acdcEnabled":false,"dfaEnabled":false,"editorialSystem":"ejp","reportingPortfolio":"","inReviewEnabled":true,"inReviewRevisionsEnabled":false},"keywords":"DNA, replication, nick, helicase, fork, homologous repair, recombination, PriA, restart","lastPublishedDoi":"10.21203/rs.3.rs-6364374/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-6364374/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003eMany antibiotics indirectly generate reactive oxygen species (ROS) that can damage bacterial genomes. Oxidised nucleobases become genotoxic when they are targeted for repair through excision, generating a single-strand discontinuity that can be converted to a double-strand break (DSB) by an oncoming replication fork. Because the genomic location of nucleobase oxidation is stochastic, investigating the fate of DNA replication machinery (replisome) at single-strand discontinuities has been limited. Here we have addressed this issue by expressing Cas9 nickases in \u003cem\u003eBacillus subtilis\u003c/em\u003e to create site specific single-strand discontinuities in a bacterial chromosome. We find that nicks in either leading or lagging strand arrest bacterial replication fork progression and generate a DSB that requires repair using homologous recombination to allow replication restart. These discoveries provoke reassessment of the fundamental mechanism of bacterial homologous recombination and provide insights to the development of alternative antimicrobials by identifying a specific pathway that can potentiate ROS-dependent bacterial killing.\u003c/p\u003e","manuscriptTitle":"Rescuing bacterial genome replication: essential functions to repair a double-strand break and restart DNA synthesis","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2025-05-08 07:02:03","doi":"10.21203/rs.3.rs-6364374/v1","editorialEvents":[],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"nature-communications","isNatureJournal":true,"hasQc":false,"allowDirectSubmit":false,"externalIdentity":"NCOMMS","sideBox":"Learn more about [Nature Communications](http://www.nature.com/ncomms/)","snPcode":"","submissionUrl":"https://mts-ncomms.nature.com/","title":"Nature Communications","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"ejp","reportingPortfolio":"Nature Communications","inReviewEnabled":true,"inReviewRevisionsEnabled":false}}],"origin":"","ownerIdentity":"d7b7da42-449e-4e2a-93f9-0a5d8298bb4c","owner":[],"postedDate":"May 8th, 2025","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"published-in-journal","subjectAreas":[{"id":47150156,"name":"Biological sciences/Microbiology/Cellular microbiology"},{"id":47150157,"name":"Biological sciences/Molecular biology/DNA damage and repair"},{"id":47150158,"name":"Biological sciences/Genetics/Genomic instability"},{"id":47150159,"name":"Biological sciences/Molecular biology/DNA replication/Replisome"},{"id":47150160,"name":"Biological sciences/Biological techniques/Genomic analysis/Chromatin immunoprecipitation"}],"tags":[],"updatedAt":"2025-12-30T08:20:54+00:00","versionOfRecord":{"articleIdentity":"rs-6364374","link":"https://doi.org/10.1038/s41467-025-66550-w","journal":{"identity":"nature-communications","isVorOnly":false,"title":"Nature Communications"},"publishedOn":"2025-11-26 05:00:00","publishedOnDateReadable":"November 26th, 2025"},"versionCreatedAt":"2025-05-08 07:02:03","video":"","vorDoi":"10.1038/s41467-025-66550-w","vorDoiUrl":"https://doi.org/10.1038/s41467-025-66550-w","workflowStages":[]},"version":"v1","identity":"rs-6364374","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-6364374","identity":"rs-6364374","version":["v1"]},"buildId":"8U1c8b4HqxoKbykW_rLl7","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}