Is RecG a general guardian of the bacterial genome?

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ARTICLE IN PRESS DNA Repair xxx (2010) xxx–xxx

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DNA Repair journal homepage: www.elsevier.com/locate/dnarepair

Mini-review

Is RecG a general guardian of the bacterial genome? Christian J. Rudolph, Amy L. Upton, Geoffrey S. Briggs, Robert G. Lloyd ∗ Institute of Genetics, University of Nottingham, Queen’s Medical Centre, Nottingham, NG7 2UH, United Kingdom

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Article history: Available online xxx Keywords: Stable DNA replication Holliday junctions Replication fork collision Pathological replication

a b s t r a c t The RecG protein of Escherichia coli is a double-stranded DNA translocase that unwinds a variety of branched DNAs in vitro, including Holliday junctions, replication forks, D-loops and R-loops. Coupled with the reported pleiotropy of recG mutations, this broad range of potential targets has made it hard to pin down what the protein does in vivo, though roles in recombination and replication fork repair have been suggested. However, recent studies suggest that RecG provides a more general defence against pathological DNA replication. We have postulated that this is achieved through the ability of RecG to eliminate substrates that the replication restart protein, PriA, could otherwise exploit to re-replicate the chromosome. Without RecG, PriA triggers a cascade of events that interfere with the duplication and segregation of chromosomes. Here we review the studies that led us to this idea and to conclude that RecG may be both a specialist activity and a general guardian of the genome. © 2010 Elsevier B.V. All rights reserved.

Contents 1. 2. 3. 4. 5. 6. 7. 8.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Replication initiation and termination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Replication in the face of DNA damage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A role for RecG in recombination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A role for RecG in replication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The role of RecG in stable DNA replication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A model for pathological replication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conflict of interest . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction The RecG protein of Escherichia coli is a double-stranded DNA translocase that can target a variety of branched DNA substrates including Holliday junctions, three-strand junctions, D-loops and R-loops [1–4]. RecG homologues are found in most bacterial species with the only known exceptions being within the mollicutes (e.g. Mycoplasma genitalium and M. pneumoniae) and the chlamydiae (e.g. Chlamydophila pneumonia and Chlamydia trachomatis) [5,6]. In E. coli, cells lacking RecG show a pleiotropic phenotype, with defects in various aspects of DNA metabolism, making it rather difficult to pin down exactly what the protein does in vivo.

∗ Corresponding author. Tel.: +44 115 823 03 03; fax: +44 115 823 03 38. E-mail address: [email protected] (R.G. Lloyd).

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The dramatic increase in sensitivity to DNA damaging agents observed when recG null alleles are combined with ruv mutations inactivating the RuvABC Holliday junction resolvase has led to the suggestion that RecG and RuvABC are part of two overlapping pathways to overcome damage to DNA via a recombination mechanism [7,8]. In addition, the low viability of strains lacking both RecG and the replication restart protein PriA [9,10] has indicated that RecG might have a role in the restart of arrested replication forks. Furthermore, the presence of RecG appears to limit damage-inducible stable DNA replication (SDR), a form of synthesis that is independent of the main replication initiator DnaA and can initiate away from the normal origin (oriC) [11]. SDR has been shown to be dependent on recombination [11] and is dramatically exacerbated in the absence of RecG [12,13]. It has also been shown that SDR can occur constitutively in cells lacking certain proteins, including RecG and RNase HI [11,12]. The potential role of RecG in recombination and in the rescue of stalled replication forks has been discussed exten-

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sively in the literature [6,14–20]. In contrast, the reported role of RecG in SDR has received relatively little attention. Recently we presented evidence suggesting that RecG has an important function in the control of chromosome replication and segregation in bacteria [13] and that this is linked to RecG’s role in limiting DnaA-independent SDR [11,13,21]. Our data led to the hypothesis that in cells lacking RecG, fork collisions outside of the normal terminus area can trigger a cascade of pathological replication and extensive recombination, which causes major problems for chromosome segregation and cell cycle progression. Thus, it appears that RecG has a crucial role in the regulation and prevention of origin-independent pathological DNA synthesis [13,21]. Our data support the idea that one important function of the specialised termination area of bacteria is to limit the potentially pathological consequences of chromosome re-replication [13,21–23]. Here we review the studies that have led to this conclusion and discuss other advances in our understanding of the multifunctional RecG protein. To place this in context, we first provide a brief overview of the relevant features of chromosome replication and segregation in bacteria, both during normal growth and in the face of damage to DNA.

2. Replication initiation and termination Accurate replication of DNA and faithful transmission of duplicated chromosomes are major challenges for dividing cells and require precise coordination and regulation of chromosome replication and segregation. In eukaryotic cells, initiation of replication at multiple origins per chromosome needs to be tightly regulated to ensure that replication of any given chromosomal area occurs exactly once per cell cycle [24]. A complex licensing system ensures that origins fire no more than once per cell cycle [24–26]. Origins are licensed by the assembly of the pre-replication complex before entry into S-phase. Initiation of DNA synthesis causes the assembled replisome components to move away from the origin. Licensing is inhibited once S-phase has been initiated, which prevents formation of another pre-replication complex, thereby limiting firing of every origin until duplication of the chromosomes has been completed and segregation has been initiated [26]. Without this regulation the danger of under- and overreplication of the chromosome is increased, with both situations leaving active replication forks spread over the chromosomes, causing interference with chromosomal segregation [25]. Furthermore, over-replication can result in head-to-tail collisions of forks, leading to the accumulation of linear DNA fragments, which might engage in genome-destabilising recombination events [27]. Similar to eukaryotes, bacterial cells must initiate replication once per cell cycle to allow successful transmission of the chromosome [28,29]. But in contrast, most bacteria initiate synthesis at a single chromosomal origin (oriC). However, the timing of origin firing remains of crucial importance. Initiate too soon and there is risk of run-off replication and fork collapse after head-to-tail collisions when the primary forks stall [30]. Nevertheless, origin firing is far less restrictive than in eukaryotes. Rapidly growing E. coli cells can re-initiate replication at the origin before the previous round of replication has terminated, allowing a generation time of about 20 min, which is approximately half the time normally required for replication of the entire chromosome [31]. In E. coli, replication initiates at oriC under the control of DnaA protein [32] (Fig. 1A). The two forks established proceed around the circular chromosome in opposite directions until they meet within a broad termination zone opposite the origin. This zone is flanked by polar sequences (ter) bound by Tus protein. The Tus–ter complex allows forks to enter, but not to leave [22,33,34] (Fig. 1B and C), dividing the chromosomes into two halves termed ‘replichores’ and

restricting termination to a specialised area that contains additional genetic elements involved in orchestrating chromosome segregation [22,35] (Fig. 1A–C). But the path of the two forks traversing each replichore is not always smooth. If one of the forks becomes blocked and fails to be rescued, the remaining portion of the replichore will not be duplicated since the second fork will be trapped in the terminus area. Although there is little doubt that the block imposed by Tus–ter complexes can be overcome eventually [36], especially if the primary arrested fork is met by a secondary following fork [37], the efficiency of replisome blockage is quite extensive. Sharma and Hill used a strain background in which a 2 kb region was excluded from replication by introduction of ter sites in a blocking orientation either side of dif. Expression of Tus protein in this background caused a rather dramatic decrease in the survival of cells [38]. Given that there are 5 ter sites per replichore [34], it seems likely that Tus–ter should impose quite a harsh constraint. This constraint may explain why mechanisms promoting fork rescue and replication restart appear to be so important for bacteria [15,39,40]. In eukaryotes, the multiple origins per chromosome and lack of termination sites that are a permanent block to progression of replication forks mean that such incomplete replication would be less likely. Any chromosome segment left un-replicated by a blocked fork could be duplicated by another fork coming from an adjacent origin. Even if converging forks were both blocked, replication could be completed if one of the many dormant origins present in eukaryotic chromosomes was located in between and was induced to fire [41]. However, a question remains over why fork movement has to be restricted in bacteria. A potential explanation comes from the observation that in many bacterial species almost all highly expressed genes are transcribed in the same direction as replication, whereas weakly expressed genes show no such correlation [23,42–47]. The same is true for most of the genes essential for viability. In E. coli at least, this polarity appears to be maintained by the organisation of the chromosome into the two replichores. This arrangement strongly implies that co-orientation of transcription and replication has evolved under selective pressure. Indeed, in vitro data suggest that a head-on collision of replication and transcription complexes leads to increased pause times of the complexes [48]. Furthermore, in vivo data using a plasmid-based assay revealed that the topological stress caused by head-on collision events can lead to knotting of the daughter duplexes behind the fork [49], consistent with head-on collisions being a serious impediment to chromosome replication. More support for this idea comes from experiments conducted with Bacillus subtilis. Wang et al. measured replication fork progression in a strain in which oriC had been moved from its original position to a site half way to the terminus area. As a consequence of moving oriC, one replication fork has to move through a quarter of the chromosome in the opposite direction to normal [50]. Measurement of replication fork progression revealed that replication was significantly delayed. This delay was only observed in the chromosomal area in which replication opposed the normal direction and was detectable only when transcription was active [50], thus supporting the idea that co-orientation of transcription and replication has evolved to allow rapid and efficient replication of the chromosome. Although a general co-directionality of transcription and replication is not observed in eukaryotic cells, co-directionality seems to be the rule in those genomic areas with exceptionally high transcription activity [47]. Thus, in eukaryotic cells replication fork movement seems to be restricted by only a few strategically placed replication fork barriers, which stands in stark contrast to the situation in bacteria. In E. coli the 10 ter sites (terA–terJ) [34] span an area of almost 50% of the chromosome (Fig. 1B). In line with the



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Fig. 1. DNA replication and termination in E. coli. (A) Diagram illustrating initiation and termination of chromosome replication. (B) Schematic representation of the locations of the ter sites (terA–terJ), and oriC on the E. coli chromosome. Each group of ter sites per replichore is semi-permissive, only allowing continuation of the replisome duplicating the replichore they are located on. Thus, terC, terB, terF, terG and terJ are orientated to block clockwise moving forks, whereas terA, terD, terE, terI and terH are orientated to block only anticlockwise forks. (C) Replisomes converging on Tus–ter complexes (permissive orientation of the complex is indicated by an arrow).

high efficiency of Tus–ter [38], the majority of replication forks are stopped at the innermost ter sites [51]. It remains unclear as to why so many ter sites are needed and why it should be necessary for them to span such an enormous area of the chromosome. 3. Replication in the face of DNA damage In eukaryotic cells a variety of surveillance mechanisms (checkpoints) make sure cells progress through the cell cycle only when appropriate to do so. For instance, the G1 –S transition checkpoint inhibits initiation of DNA replication and therefore entry into S-phase if lesions are detected in the template. This delay provides time for repair activities to restore the template, after which replication might proceed unhindered. Without such coordination, there is increased risk of mutation, genomic instability and cell death [52]. In contrast to eukaryotic cells, origin firing does not appear to be restricted by a G1 –S checkpoint in bacteria following UV irradiation. Replication forks stall at UV-induced lesions and undergo time-consuming processing before replication restarts, providing an opportunity for repair activities to clear the path ahead [53,54]. However, the delay in finishing chromosome replication does not lead to a delay in origin firing. New rounds of replication are initiated (almost) at the normal rate, enabling a cluster of replication forks to traverse the chromosome, causing a quite substantial overrepresentation of the origin over the terminus region at early time points following irradiation [53] (Fig. 2A). Once all the lesions have been repaired, the whole cluster of replication forks is capable of multiplying the chromosome very rapidly, creating multiple

complete copies of the genome [53]. Analysis of cellular replication of UV-irradiated wild type cells showed that a mild UV dose caused a filamentation period of 60 min in which no increase in the formation of viable cells is observed. However, following this division delay, division of wild type cells occurs more frequently than in unirradiated cells, leading to a reduced generation time of ∼15 min compared to 21 min measured in the absence of irradiation. After approximately 60 min of very rapid divisions, cellular replication continues with a rate similar to unirradiated cells with a total number of viable cells that is not much reduced in comparison to the mock-irradiated sample (Fig. 2B) [54]. Thus, it appears that re-firing of the origin compensates for the delay required for processing of forks blocked by UV-induced lesions [53,54]. Although such a cluster of replisomes will most likely lead to formation of complex chromosomal structures, the general directionality of replication is unchanged and strictly follows the replichore arrangement. However, UV induces further events which can interfere with this directionality. Despite the regulatory mechanisms limiting initiation of replication to oriC, DNA damage has been shown to cause SDR in other chromosomal areas after induction of the SOS response [11]. It is not clear whether such stress-induced or unscheduled replication contributes to the successful completion of replication. In theory, new initiation events could provide a solution to blocked forks in bacteria, similar to the dormant origins in eukaryotic cells. However, whilst SDR could obviate the need to rescue a stalled fork, its initiation at various sites in the chromosome would increase the incidence of fork collisions. Moreover, studies of DNA replication in vitro


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Fig. 2. Effect of UV on cell cycle progression in E. coli cells. (A) Schematic representing the events following induction of UV damage. Replication forks stall at UV-induced lesions and undergo time-consuming processing before replication restarts, providing an opportunity for repair activities to clear the path ahead. New rounds of replication are initiated almost at the normal rate, enabling a cluster of replication forks to traverse the chromosomes once all the lesions have been repaired, allowing rapid creation of multiple complete copies of the genome. (B) Viable cell replication of wild type E. coli cells following irradiation. Cells were UV- or mock-irradiated, diluted in conditioned medium and incubated in a 37 ◦ C shaking water bath, as described [53,54]. At each time point samples were removed and the viable titre determined. Estimated generation times at various stages of the experiment are indicated. Image has been reproduced from [54].

revealed that without Tus to curb fork movement, considerable rereplication of DNA that had already been replicated was detectable [55]. It was postulated that re-replication is initiated by a replisome displacing the 3 end of the nascent leading strand made by the fork coming in the other direction, thus generating a 3 flap at which a new replisome could be loaded. This scenario is supported by in vivo data in E. coli showing that a lack of terminators leads to DNA over-replication [56,57]. Although these events seem to appear at low frequencies under normal conditions [57], the data available suggest that collision events nevertheless might pose a threat for the genomic integrity if not contained within a specialised area. Strand displacement is not observed with DnaB alone in vitro [58], indicating that re-replication is a particular risk following collisions between fully fledged replisomes. Restriction of replication fork collisions within a replication fork trap might provide therefore an additional explanation for the importance of a defined termination area [22,23]. Any overreplication caused by collision of forks would quickly be stopped by a Tus–ter complex (Fig. 3). Indeed, recent data [13] provide evidence that in the absence of RecG an increased number of fork collisions occur outside of the terminus area, triggering a cascade of pathological replication and extensive recombination with rather catastrophic consequences. The data support the idea that the terminus area might provide the means to prevent pathological replication caused by head-on collision events of forks coming from the origin during the normal cell cycle. Any overreplication triggered by the collision of two forks would not be allowed to escape the terminus area, thus allowing the replichore arrangement to be maintained and avoiding collision events with transcription complexes or following forks coming from the origin. However, the idea that RecG might limit pathological replication and avoid unnecessary recombination seems at odds with earlier evidence indicating that RecG is needed to promote recombination.

4. A role for RecG in recombination The recG gene was initially identified in a screen for mutants showing some degree of recombination deficiency, suggesting a role of recG in recombination and DNA repair [59]. Further investigations revealed that recG mutants show a slight reduction of recombination in conjugational Hfr crosses and a mild increase in sensitivity to UV, mitomycin C and ionising radiation [7,8]. However, combining a recG null allele with a ruv mutation inactivating the RuvABC Holliday junction resolvase effectively blocks recombination and greatly enhances sensitivity to UV light [7], suggesting that RecG and RuvABC are part of two overlapping pathways for promoting the late stages of recombination and for overcoming UV-damaged DNA [7]. This idea was supported further by in vitro studies demonstrating that RecG catalyses branch migration of Holliday junctions [60] and by in vivo studies showing that activation of the normally quiescent rusA gene suppresses a ruv deletion very effectively, but only in the presence of RecG protein [61,62]. RusA targets Holliday junctions with high specificity and efficiency [63], but does not catalyse branch migration [63,64], suggesting that the branch migration activity of RecG is critical in the absence of RuvAB to allow resolution of Holliday junctions by RusA [61]. Although RecG drives branch migration very efficiently, it also unwinds a variety of other branched DNA structures, including Dloops and R-loops [1–3]. If RecG were to unwind D-loops in vivo, it would be expected to reduce recombination, which does not fit well with the idea of RecG being a recombinase that helps to complete the recombination reaction. Another particular feature of RecG is that it can unwind a forked DNA structure to create a Holliday junction [65], and can do so in an in vitro replication system [66]. The crystal structure of Thermatoga maritima RecG has revealed how this is accomplished [67] (Fig. 4A). RecG has conserved helicase domains linked to a ‘wedge’ domain, which provides specificity for binding a branched DNA structure. It has been proposed that the helicase motor acts as a dsDNA translocase, pulling the parental



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Fig. 3. The terminus area can prevent extensive over-replication caused by fork collisions. Opposing replisome complexes cause displacement of the nascent leading strand as forks collide. The 3 flap can be used for PriA-dependent formation of new replication forks. Progression of forks would cause over-replication (highlighted in red) of the chromosome. Progression of the newly formed replication forks within the termination area will be quickly stopped by a Tus–ter complex, allowing degradation of the over-replicated area by exonucleases (RecBCD). Ligation will then lead to successful termination of replication.

strands of a replication fork through separate channels flanking the wedge, neither wide enough to accommodate duplex DNA [67]. This has the effect of stripping off the nascent strands and allowing the parental strands to re-anneal, as suggested by biochemical studies [66]. The unwound nascent strands may then also anneal, so that, as the protein continues to translocate along the parental duplex, a Holliday junction forms around the wedge and a ‘nascent strand duplex’ is spooled out in front (Fig. 4B). This final stage is most likely equivalent to the observed Holliday junction branch migration reaction catalysed by RecG [60]. However, there is accumulating experimental evidence suggesting that the idea of two overlapping recombination pathways is too simple. ruv deletion mutations are either synthetically lethal or very sick if combined with dam, polA or uvrD mutations, all of which are involved in DNA metabolism and repair [68–70]. The synthetic lethality suggests that in the absence of dam, polA or uvrD cells rely more on the Holliday junction resolvase. But why should RuvABC be so vital when the postulated RecG pathway for Holliday junction resolution is intact? The synthetic lethality or sickness of ruv dam,

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ruv polA and ruv uvrD double mutants can be efficiently suppressed if rusA expression is activated, strongly supporting the idea that the lethality is caused by the accumulation of Holliday junction intermediates. Furthermore, the suppression by RusA critically depends on the presence of RecG, in line with the idea that RecG catalyses branch migration of Holliday junctions (J. Zhang, A.A. Mahdi, G.S. Briggs, R.G. Lloyd, unpublished). However, if RecG would act in an alternative pathway for resolving late recombination intermediates, this pathway clearly is not very efficient, given that it should be fully functional in the double mutants tested. These results suggest that the branch migration activity of RecG might lack an associated Holliday junction resolvase activity, implying that RecG on its own does not provide an efficient alternative pathway to RuvABC for resolution of late recombination intermediates. This idea is further supported by experimental data from a plasmid-based assay showing the persistence of specific recombination intermediates following UV irradiation in ruv mutants but not in recG mutants [71]. In the absence of RecG, increased formation of branched intermediates is observed, but these intermediates disappear almost as quickly as in the wild type control [71]. In addition, in a recent study Wardrope et al. were able to show that 2-aminopurine treatment in cells lacking ClpX protease leads to EcoKI-dependent fragmentation of the chromosome. While clpX recG double mutants showed accumulation of linear fragments in the presence of EcoKI, these fragments were absent in a ruvC background unless RusA was expressed, indicating that without the Holliday junction resolution activity of RuvC the linear fragments are tied together by Holliday junctions [72]. This lack of accumulation of recombination intermediates in recG mutant cells seems to be in line with the work by Donaldson et al. [71]. Thus, there is little doubt that RecG can process recombination intermediates in the absence of RuvAB. However, the data described suggest that RecG must also work in a pathway which has very little overlap with the RuvABC-dependent Holliday junction resolution pathway. The original idea of RecG being primarily a recombinase that promotes recombination is further complicated by the fact that different experimental assays produce contradictory results. Lovett and co-workers were able to show that the deletion frequency of direct repeats located either on a plasmid or on the chromosome was increased in the absence of RecG, whereas ruv deletions caused a reduction [73,74]. Additional data from our laboratory confirmed that the increase observed with the plasmid-based assay in the absence of RecG is dependent on RecA but not on RuvABC (CJR, RGL, unpublished). Thus, while the decrease in recombination observed in conjugational Hfr crosses in recG mutant cells [8] argues that RecG can stimulate recombination, the increased recombination at tandem repeats observed in cells lacking RecG [73,74] suggests that RecG prevents certain types of recombination. Therefore it appears that RecG can act both as an activator as well as a suppressor of certain recombination events. However, the finding that recombination events causing tandem repeat deletions in recG mutant cells are independent of RuvABC (CJR, RGL, unpublished) supports the idea that RecG might be involved in a RuvABC-independent recombination pathway. An increase in the number of recombination events in the absence of RecG was confirmed by other assays [75,76]. Grove et al. designed a system in which an I-SceI-induced double strand break can result in a recombination event in which one or both ends of the broken chromosome might engage either the intact sister chromatid or a plasmid. Recombination events involving the plasmid can produce detectable recombinants [76]. This system revealed an increased number of recombination events in the absence of RecG. However, in contrast to the tandem repeat recombination assay, the recombination events observed were entirely dependent on RuvC, suggesting that RecG is involved in the disruption of



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Fig. 4. (A) Structure of RecG bound to a lagging strand fork. E. coli RecG has been modelled onto the X-ray crystal structure of Thermotoga maritima RecG [67]. Domain 1 is shown in grey, with the wedge domain highlighted in green. The helicase domains 2 and 3 are shown in blue. (B) Schematic representation of the reactions catalysed by RecG in vitro. (i) RecG can interconvert forked structures and Holliday junctions and efficiently branch migrates Holliday junctions. (ii) RecG catalyses the dissociation of Rand D-loops and (iii) rapidly converts 3 flaps into 5 flaps.

structures which otherwise cause formation of Holliday junctions [76]. Although the genetic interaction of ruv and recG suggests some degree of overlap in overcoming UV-damaged DNA, both pathways appear to have different specificities with one not always being an alternative to the other. While RuvABC is clearly acting late in recombination to resolve Holliday junctions, the precise role of the RecG is less clear. 5. A role for RecG in replication The observation that strains lacking both RecG and PriA are very poorly viable [9,10] revealed another facet of the role of RecG in the cell. PriA is the main replication restart protein and facilitates loading of the DnaB replicative helicase and subsequent replisome assembly at branched DNAs [39]. It is required for initiating SDR and also for restarting DNA synthesis following exposure to genotoxic agents [11,13,77]. Cells lacking PriA show a reduced viability and an increased sensitivity to DNA damage, phenotypes which are generally attributed to the deficiency in rescuing stalled or damaged forks [39]. However, the failure to curb unnecessary recombination

is another possibility [78]. PriA also has a helicase activity and can unwind any 5 strand at the branch point of a fork to create a landing pad for DnaB [79]. PriA helicase activity is thought to be part of a PriC-dependent re-initiation pathway and is not essential for PriA’s function in replication restart [39,80–82]. The genetic interaction of RecG and PriA led to the suggestion that RecG might have a direct role in the repair or restart of damaged replication forks. Support for this idea came from a search for suppressors of the recG mutant phenotype, in which a series of strains were identified that had single amino acid substitutions within or near the helicase motifs of PriA [83]. These PriA helicase mutations suppress not only the damage sensitivity of recG mutants quite effectively [10,83,84], but also the extreme filamentation phenotype that was observed in cells lacking RecG following irradiation with mild UV doses [13,85]. This suppression suggests that the helicase activity of PriA is involved in the generation or persistence of a toxic intermediate in the absence of RecG [13]. recG deletion mutants carrying a plasmid which allows controlled expression of wild type recG show fast and efficient break-down of filaments into small and healthily growing cells if expression of recG is induced 60 min after irradiation. Thus, the intermediates formed or stabilised by PriA helicase



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activity remain accessible for processing via RecG over long periods despite the presence of RuvABC in these cells [13]. The fact that RecG can interconvert replication forks and Holliday junctions seemed to provide a link between RecG’s role in recombination on the one hand and in replication on the other, and has led to the suggestion that RecG might be involved in the reversal of replication forks stalled at small lesions [14,16]. Exactly what happens when a fork does stall remains far from clear but is likely to vary according to the blocking lesion [15,39,53,86]. Meneghini and Hanawalt [87] suggested that a lesion in the leading strand template blocks fork progression whereas a lesion in the lagging strand template does not. The lagging strand polymerase simply skips an Okazaki fragment, leaving a gap, an idea that was strongly supported by in vitro as well as in vivo data [88–90]. For the leading strand, however, arrest of the DNA polymerase does not necessarily cause an immediate arrest of the complete replication fork. Synthesis of the lagging strand can still continue [88,89,91,92], producing a structure where the newly synthesized lagging strand is longer than the leading strand. In this situation the blocking lesion might still be masked by the stalled polymerase or cannot be repaired because the template is unwound. A solution was postulated by Higgins et al. [93] and Fujiwara and Tatsumi [94]. Both groups presented evidence that, in mammalian cells, blocked forks reverse to form a Holliday junction structure. This replication fork reversal would transfer the lesion back into double-stranded DNA, enabling excision by one of the many repair systems or error-free bypass. Several mechanisms have been discussed for replication fork reversal, including positive supercoiling in front of the replication fork [95] as well as a RecA-driven reaction [15,19,86]. However, the biochemical analysis of RecG showing that the protein is able to catalyse the interconversion of fork and Holliday junction structures in vitro [65,66] in combination with the results suggesting a role of RecG in replication restart made it an attractive candidate. A role of RecG in replication fork reversal therefore would provide a link between replication restart and recombination [15,17–19]. But the fate of replication forks arrested at UV-induced damage is still poorly understood. The stalling of replication by inactivation of components of the replisome has been shown to induce replication fork reversal in vivo. Several pathways have been described by which this reversal can be achieved. However, none of these pathways appear to require RecG [40] and direct in vivo evidence supporting a role of RecG in replication fork reversal following UV irradiation is lacking. Indirect support for a role of RecG in replication comes from in vivo studies in B. subtilis showing that the repair helicases PriA, RecG and RecQ interact with SSB and co-localise with replisome components. Co-localisation was not observed in strains expressing a C-terminal truncated form of SSB, suggesting that localisation is achieved via a specific interaction with SSB [96]. Given that PriA is the primary replication restart protein [39], an attractive hypothesis is that localisation near replication forks reflects an increased rescue potential for forks that might get stuck at DNA lesions [96]. This fits very well with in vitro as well as in vivo data. Replication restart in UV-irradiated E. coli cells depends critically on DnaC [53], a protein that is essential for loading of the DnaB replicative helicase on ssDNA, the first step in the assembly of a replisome [32,97]. This fits with in vitro data showing that a block to synthesis by the leading strand polymerase due to an abasic site uncouples DNA synthesis and unwinding. DnaB loses contact with the polymerases and moves ahead, exposing the leading strand template as it leaves the polymerase stably bound at the arrest site [98]. RecA may then be loaded on the exposed template strand, displacing any bound single-strand DNA binding (SSB) protein with the aid of the RecF, RecO and RecR proteins [98–100]. The loading of RecA and other recombination proteins at dislocated

7

forks has been implicated in the removal of replisome components remaining on the template and in reshaping the fork [15,78,98]. Additional factors act to limit any unnecessary recombination as a result of RecA loading [70,78,101–103]. However, once the replicative helicase is removed, replication restart depends critically on PriA- or PriC-mediated loading of DnaB and subsequent assembly of a new replisome complex [39,104]. Thus, having an increased concentration of PriA near active forks might allow cells to quickly and efficiently restart replication following arrest of forks at lesions [96]. Given that both RecG and RecQ have been postulated to be involved in the processing of forks stalled at lesions [16,54,105], the same rationale applies, leading to the suggestion that SSB is a platform for coordinating repair at active replication forks [106], similar to the idea of BASC, a super-complex which coordinates multiple repair activities in eukaryotic cells [107]. This is greatly supported by an increasing number of studies showing that numerous proteins involved in replication and repair, including RecG, RecQ and ExoI, interact with SSB in B. subtilis, E. coli and other bacteria [106,108–115].

6. The role of RecG in stable DNA replication A prediction of the hypothesis that RecG might be involved in the quick and efficient processing of stalled replication forks would be that the rate of replication should be reduced following UV irradiation in cells lacking RecG, since forks should be stalled at lesions for longer than normal. To investigate this, Donaldson and co-workers measured the rate of [3 H]thymidine incorporation in UV-irradiated recG cells. The incorporation pattern of wild type and recG cells looked almost identical, which led to the conclusion that RecG might not be involved in restart of replication [86,116]. However, our recent studies revealed that net [3 H]thymidine incorporation is a poor measure for replication restart [53,54]. They demonstrated that at least three processes contribute significantly to thymidine incorporation after UV irradiation: DnaA-dependent initiation of replication at the normal origin of replication (oriC), DnaA-independent initiation of SDR at other positions in the chromosome and, thirdly, replication associated with rescue of the forks initially stalled at lesions, consistent with previous studies [11,53,54,117]. Hence, measurements of net [3 H]thymidine incorporation could be misleading as a delay in one of the three processes might be masked entirely by an increased activity of the other. Indeed, it was shown by Hong et al. that, similar to cells lacking RNase HI, recG mutant cells exhibit constitutive levels of SDR (cSDR) [12]. RecG has been shown to unwind the D-loop and Rloop structures [1–3] (Fig. 4B), whereas RNase HI degrades RNA from RNA:DNA duplexes [11], and it was speculated that the persistence of R-loops in both rnhA and recG mutants is responsible for the observed levels of cSDR [12]. In addition, the levels of SDR observed in cells lacking RecG are partially caused by an increase in damage-induced SDR (iSDR) [12]. iSDR has been reported to be dependent on the activation of the SOS response [11] and cells lacking RecG exhibit a mild SOS-constitutive phenotype [8,118,119], which might be responsible for the increased iSDR. Additional SOS induction by thymine starvation leads to iSDR levels, which, in the absence of RecG, exceed the levels observed in wild type cells significantly [118]. When we examined replication in greater detail in a strain carrying the thermosensitive dnaA46 allele, we found that the absence of RecG has a rather dramatic effect (Fig. 5). In mock-irradiated dnaA46 single mutants shifted to 42 ◦ C, a temperature at which the mutant DnaA protein is inactive, incorporation of [3 H]thymidine continues for some time before reducing severely, consistent with synthesis by the majority of existing replication forks coming to an end and



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Fig. 5. Comparison of DNA synthesis in the presence and absence of RecG. (A–C) [3 H]thymidine incorporation in wild type, recG, dnaA46 and recG dnaA46 cells [13,53]. Cultures were grown at 30 ◦ C in Davis medium, filtered onto a 0.22 ␮m cellulose acetate membrane and UV- or mock-irradiated directly on the membrane. Cultures were shifted to 42 ◦ C and replication monitored by addition of [3 H]thymidine [13]. Data are means (±SE) of three or more experiments. Data have been reproduced from [13].

the failure to initiate new rounds from oriC in the absence of DnaA. UV irradiation increases the level of incorporation quite significantly [13,53] (Fig. 5B). This increase is consistent with induction of SDR, which is defined by its independence of DnaA [11]. In mock-irradiated dnaA46 recG cells the level of incorporation after the shift to 42 ◦ C is already significantly higher than in mockirradiated dnaA46 cells (Fig. 5C). This is in line with the increased levels of SDR reported in recG mutants [12]. UV irradiation results in a much higher level of incorporation in dnaA46 recG cells at restrictive temperature, almost as high as in a recG single mutant. Thus, replication is clearly much affected in the absence of RecG, both with and without UV irradiation [13]. The idea of SDR contributing substantially to replication in cells lacking RecG was supported by the investigation of replication patterns in dnaA46 temperature sensitive derivatives of strains in which the origin and terminus areas of the chromosome were tagged with fluorescent proteins [13,21]. UV irradiation led to a moderate accumulation of origin as well as terminus foci over several hours in dnaA46 cells at restrictive temperature [13] (Fig. 6A), in line with reports showing that SDR can continue over extensive periods [11]. Eliminating RecG from dnaA46 cells exacerbated this effect drastically. In dnaA46 recG cells UV irradiation led to a rapid and very dramatic multiplication of both origin and terminus foci, consistent with the observed increase in SDR. Furthermore, in contrast to the regularly interspersed origin and terminus foci observed in irradiated dnaA46 (recG+ ) cells, the amplified origin and terminus foci observed in dnaA46 recG cells frequently were forming extensive and discrete clusters within the filaments [13], an effect also seen in recG cells in which origin firing is fully functional [13] (Fig. 6B). The persistent and extensive replication observed in the absence of RecG led us to consider the possibility that DNA damage might be only the initial trigger for the formation of additional replication forks. The perpetuation of SDR would then be the result of secondary replication forks initiated as a result of pathological events arising from unscheduled collisions between opposing forks [13,55–57]. RecG might normally limit this secondary pathology, hence explaining the much-elevated SDR and much-prolonged defects in chromosome segregation and cell division in recG mutant cells [13]. This theory was supported by results showing that SDR was able to continue over extensive periods in recG cells regardless of the applied UV dose, provided enough lesions were introduced to trigger the initial events. A substantial reduction of the UV dose made very little difference to the amplification of foci [21].

Irradiation with a reduced UV dose revealed another characteristic phenotype of cells lacking RecG. Estimation of the number of origin and terminus foci revealed a dramatic fluctuation of the calculated ori/ter ratio, with some filaments showing a more than 6-fold over-representation of origin over terminus foci, while others showed a greatly increased number of terminus foci (Fig. 6C). These data led to the conclusion that damage-induced SDR is far more pathological in the absence of RecG and can lead to rapid and uncontrolled over-replication of limited chromosomal areas [21]. As mentioned before, the sensitivity to DNA damaging agents as well as the severe filamentation phenotype following UV irradiation in cells lacking RecG are very effectively suppressed in cells carrying a priA300 allele, expressing a helicase-deficient PriA protein which is functional in replication restart. The suppression suggests that PriA helicase activity is responsible for either the formation or the persistence of some DNA intermediates that could result in this pathological replication. Indeed, when we investigated the amplification of chromosomal areas in UV-irradiated dnaA46 recG priA300 cells at 42 ◦ C, amplification of the origin and terminus was dramatically reduced [13] (Fig. 6A), in line with a study by Tanaka et al. showing that strains expressing helicase defective PriA show reduced levels of constitutive as well as damage-induced SDR [77]. Together these results strongly suggest that much of the pathology observed in recG cells may actually be due to the increased SDR, which is further amplified by UV-induced damage and continues long after the primary lesions have been repaired. The reduction of SDR in cells lacking PriA helicase activity therefore limits the pathology caused by the lack of RecG. 7. A model for pathological replication Why should an increase in the number of replication forks established following initiation of SDR have such severe consequences? Does this have something to do with the way replication is then terminated? Although the events associated with termination have not been fully dissected, it would seem to require little more than a merging of forks, dissociation of the replisomes and ligation of the nascent strands after the filling of any remaining gaps (Fig. 7Ai–iii). However, the collision of two forks might not always proceed so smoothly. We considered the possibility that when two forks collide one replisome may sometimes displace the nascent leading strand of the fork coming in the other direction, establishing a 3 ssDNA flap with the potential to support further replication. In wild type cells, such flaps might be eliminated by 3 –5 ssDNA exonucleases (Fig. 7E), or converted to 5 flaps by RecG translocase and then



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9

Fig. 6. Effect of RecG on replication and segregation of origin and terminus areas. (A) Fluorescence microscopy showing UV-induced DnaA-independent synthesis in dnaA46 temperature sensitive strains at restrictive temperature following UV irradiation (combined phase contrast and fluorescence images are shown). In cells lacking RecG UV irradiation leads to a PriA helicase dependant increase in origin (red) and terminus foci (green) in the absence of oriC firing [13]. For these experiments strains were used in which origin and terminus areas of the chromosome were tagged with lacO and tetO arrays, respectively. The strains carried a plasmid encoding LacI-eCFP and TetR-eYFP repressors to decorate these arrays [13,53,138]. Cells were grown at permissive temperature prior to UV irradiation and shifted to 42 ◦ C directly after UV. (B) Replication and segregation of origin (red foci) and terminus (green foci) areas of the chromosome in fully replication proficient cells (dnaA+ ) (combined phase contrast and fluorescence images are shown). (C) UV-induced SDR leads to uncontrolled DNA amplification in the absence of RecG. Irradiation with a low UV dose leads to a drastic over-representation of either origin (red) or terminus (green) foci in the absence of oriC firing (combined phase contrast and fluorescence images are shown) [21]. The light arrows point to filaments with the number of termini exceeding the number of origins whereas the dark arrow indicates a filament with the number of origins exceeding the number of termini. Cells were grown at permissive temperature prior to UV treatment and shifted to 42 ◦ C directly after irradiation. Images have been reproduced from [13] and [21].

eliminated by 5 –3 ssDNA exonucleases. RecG has a particularly high affinity for a 3 flap [66,120] (Fig. 7D). However, 3 flaps provide a target for PriA helicase activity [3,10,120,121], which could lead to the assembly of new replisomes that re-replicate the newly synthesised strands, generating highly recombinogenic dsDNA flaps as they proceed (Fig. 7Ci–iii). 3 flaps in particular would require the presence of PriA helicase in order to unwind the nascent lagging strand, creating a single-stranded area onto which DnaB can be loaded. Given that in vitro the affinity of RecG for 3 flaps is 10fold higher than the affinity of PriA [120], 3 flaps might normally be processed almost exclusively by RecG, explaining the low frequency of over-replication events in tus mutant cells which have RecG available [57]. However, in the absence of RecG, 3 flap formation and unwinding via PriA helicase might become frequent enough to pose a problem.

Such events would explain the much-extended PriA helicasedependent delay in chromosome segregation and cell division observed in the absence of RecG [13,85]. The model outlined is consistent with the chromosomal over-replication observed in the absence of a functional termination system as well as the RecBCDdependent hotspot for recombination associated with the terminus area [56,57,122]. It fits particularly well with the over-replication of the leading strand observed in vitro with an oriC plasmid template and reconstituted replisomes [55]. The model is also in good agreement with the data by Donaldson et al., showing that branched DNA structures thought to reflect Holliday junctions can be detected in UV-irradiated wild type, ruv and recG cells in a plasmid-based assay [71]. However, while the intermediates observed in ruv cells persisted, those detected in recG cells disappeared as in wild type cells, as our model would predict. We



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Fig. 7. Models depicting possible outcomes of replication fork collision. (A) Fork merging and nascent strand ligation. (B) Pathological replication resulting from unscheduled replication fork collisions or during normal termination: B(i) schematic of the E. coli chromosome showing normal replication from oriC and the presence of several additional replication forks initiated as a result of SDR induction. The opposed arrowheads indicate the positions of unscheduled fork collisions outside of the normal termination zone (for simplicity, only two ter sites are depicted). B(ii) Nascent strand displacement following unscheduled collisions triggered by SDR, or at a lower frequency in the absence of SDR. (C) and (F) Pathological, PriA-dependent replication in the absence of RecG generates a dsDNA branch that can provoke recombination. (D) Termination achieved via a 5 ssDNA exonuclease after RecG converts a 3 flap to a 5 flap. (E) Termination achieved via a 3 ssDNA exonuclease. The model was revised from [21].

would expect the formation of some high molecular weight intermediates as a result of repeated recombinational exchanges, some of which may involve the over-replicated DNA invading the sister chromosome (Fig. 7Civ–v). Thus, in this case Holliday junction resolution by RuvABC may lead to the formation of chromosomal multimers. Further repetitions could produce a network of multiple chromosomal copies tied together. This interpretation is consistent with the observed synergistic phenotype of recG with xerC [123]. The XerC protein is required for the site-specific recombination at dif, which is needed to convert chromosome dimers and higher multimers to monomers [124]. The equivalent site-specific recombination system of B. subtilis is required in order to avoid chromosomal instability following elimination of the Tus analogue, RTP [125], indicating that uncontrolled termination of replication in this species might have similar pathological consequences. The model outlined predicts that 3 –5 exonucleases might be rather vital in the absence of RecG as they would provide the only means to avoid 3 flaps from being targeted by PriA, enabling replication to terminate normally. In recent studies we have shown that the viability of recG mutant cells does indeed depend on the presence of ExoI, ExoVII or the SbcCD nuclease, all of which can remove 3 flaps. We have also shown that this requirement can be overcome by eliminating the helicase activity of PriA (CJR, ALU, RGL, in preparation). The scenario of a replication catastrophe fits well with the major phenotypes associated with a recG deletion. The model predicts that the initial lesions caused by UV irradiation are only the trigger of the pathological cascade. The increased number of fork collision events

following UV irradiation in cells lacking RecG lead to formation of 3 flaps that PriA subsequently targets, at least in some cases. Once triggered, the re-replication of the chromosome perpetuates the problem even when the initiating lesions are long gone, explaining how doses as low as 1 J/m2 suffice to induce substantial delays in cell division and pathological amplification of sometimes limited chromosomal areas [13,21,85]. This explains why the expression of recG is able to efficiently alleviate the problem at almost any time following irradiation. Firstly, it would reduce the amount of SDR [12,13] and therefore fork collisions. Secondly it would help eliminate both 3 flaps and D-loop intermediates that trigger and perpetuate the cascade, enabling ongoing replication forks to finish multiplication of the chromosomes present, allowing subsequent break-down of filaments to normal dividing cells, as observed [13]. The co-localisation of RecG with SSB [96,113] will help to guide RecG to regions where it is needed. Since there is no indication that SDR forks are structurally different from regular forks [11], the concentration of RecG will be increased, specifically if two forks collide. Additionally, SSB will cover single-stranded DNA at locations where recombination is taking place [100], potentially allowing RecG to limit some exchange events and thus to reduce replication initiated at recombination intermediates. It seems likely that the specific localisation of RecG makes it effective in preventing such a pathological cascade. In the absence of RecG, ExoI will be available since it also interacts with SSB [111,126]. However, PriA will also be localised near replication forks, providing a competition with other enzymes which might result in the initiation of the cascade described.



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11

Fig. 8. UV-induced dnaA-independent replication reaches all chromosomal areas. (A) Schematic NotI restriction pattern of the E. coli chromosome. The distance from oriC to each end of the fragments is indicated. Fragments clockwise and anticlockwise of oriC are shown in red and blue, respectively; the fragment containing oriC is shown in black. (B) and (C) Visualisation of BrdU incorporation in UV-irradiated or mock-irradiated cells in the absence of oriC firing. Cells were grown at permissive temperature, UV-irradiated or mock irradiated and subsequently shifted to 42 ◦ C. At the time points indicated the cells were pulse labelled with BrdU for 10 min. Cells were lysed within agarose plugs, digested with the rare cutter NotI and the chromosomal fragments separated via pulse field gel electrophoresis. The image was reproduced from [21].

The idea of a replication catastrophe also predicts that some areas of the chromosome may be re-replicated several times if formation of multiple forks happens to be induced in close proximity. As described before, irradiation with mild UV doses does not inhibit origin firing, leading to formation of a whole cluster of three or four rounds of replisomes traversing the chromosome. If an initial collision event leads to formation of a 3 flap, the following forks coming from the origin would then collide relatively rapidly, each time with the danger of re-iterating the problem. Such events may account for the frequent asymmetric amplification of the origin and terminus areas of the chromosome apparent in the absence of RecG, which can be quite extreme (Fig. 6). An alternative explanation might be the repeated firing of distinct damage-inducible origins, as previously described [11]. However, when we tried to analyse where iSDR initiates, we found no strong evidence of such origins in our analysis of DNA synthesis mediated via SDR [21], although we cannot dismiss their existence. Our data rather suggest that UV-induced synthesis can be initiated at any chromosomal location, at least at a population level [21] (Fig. 8). Furthermore, over-initiation at oriC has been shown to result in replication fork collapse, since secondary forks are capable of catching up with primary forks, causing the formation of linear DNA fragments due to replication runoff [30]. The same rationale should apply to rapidly firing SDR origins, thereby limiting the ability of a single origin to cause over-replication. Finally, UV-induced synthesis continues after all detectable lesions have been removed [13] and it is not obvious why firing of specific SDR origins should occur once the initial trigger is gone. SDR might also contribute to yet another phenotype observed in cells lacking RecG. It has been observed that if populations of microorganisms are subjected to non-lethal selection, cells acquire mutations even though they are only slowly growing, if at all. This phenomenon is called stress-induced or ‘adaptive’ mutation

[127,128]. In E. coli, Pol IV and recombination proteins are required for these mutations to appear. In cells lacking RecG, the rate of these stress-induced mutations is dramatically increased [129,130], an effect which is appears to be due to the fact that Pol IV is overexpressed in recG mutant cells [131]. However, there is some evidence that SDR is error-prone [11], although the contribution of Pol IV to this mutagenesis was never investigated. Furthermore, SDR has also been linked to entry into stationary phase, at least under certain conditions [11]. An increased level of SDR in recG mutant cells might therefore contribute to stress-induced mutations, specifically under conditions where cells are entering stationary phase due to the growth restriction. Although our data are in line with the idea that pathological replication, rather than an accumulation of recombination intermediates, is primarily responsible for the phenotype of cells lacking RecG, they do not rule out the idea that RecG can also help to resolve Holliday junctions, as initially suggested [7]. Indeed, there is robust in vivo data suggesting that E. coli RecG is capable of catalysing branch migration, at least in the absence of RuvABC [61] (J. Zhang, A.A. Mahdi, G.S. Briggs and R.G. Lloyd, unpublished). Furthermore, experiments in Neisseria gonorrhoeae have demonstrated that ruv as well as recG are involved in pilin variation, which is necessary to achieve immune evasion by antigenic variation [132]. The fact that deletion of priA has no effect on pilin variation [132] makes it likely that the effect observed is caused by a limitation of one or several recombination pathways involving either RecG or RuvABC. However, our results in E. coli raise the possibility that any recombination reaction that increases the number of replication fork collisions may have pathological consequences that reduce viability. Hence, the observed reduction of recombination rates in conjugational Hfr crosses in the absence of RecG might be caused by a reduced ability to recover recombinant products due to the induction of the pathological cascade described rather than a direct effect of recombination.



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8. Concluding remarks Our knowledge of the genetics and biochemistry of RecG helicase has grown considerably since the gene was first described in E. coli nearly 40 years ago. However, the pleiotropic phenotype of recG mutants coupled with the wide spectrum of substrates unwound by the protein has made it difficult to pin down what the protein does in vivo. What has become apparent from the structure of RecG and from recent genetic and cytological studies is that the pleiotropic phenotype can be explained by the particular affinity of RecG not only for Holliday junctions but also for any dsDNA substrates with a 3 ssDNA branch as a component of the structure (D-loops, R-loops, stalled replication forks and repair intermediates). Such structures are common intermediates of DNA metabolism, and therefore it is hardly surprising that inactivating RecG can lead to such a variety of phenotypic effects. The replication catastrophe model we have presented for RecG is one particular example where the absence of RecG triggers a cascade of pathological replication and recombination. Similar pathological events may occur in other bacteria, but the phenotypes observed may vary according to the availability of other factors (e.g. helicases and nucleases) that enable the relevant substrates to be processed by other means. Thus, RecG may be regarded both as a specialist activity and as a general guardian of the genome. Conflict of interest The authors declare that there are no conflicts of interest. Acknowledgements The authors wish to thank Roxane Lestini and Stéphane Delmas for critical comments on the manuscript. CJR, GSB and RGL are supported by the Medical Research Council. CJR is also supported by The Leverhulme Trust. Fig. 9. Termination of replication in eukaryotic cells. The unwinding mechanism of eukaryotic helicases is still not clear and several models are discussed [133]. If the helicase encircles single-stranded DNA similar to DnaB in E. coli, strand displacement will generate a 5 flap (Ai–iii) due to the 5 –3 polarity of MCM2–7 [133,135]. The formed 5 flap can be efficiently cleaved by flap endonuclease 1 (FEN-1) [136] (Bi–ii). Alternatively, given that Okazaki fragments are relatively short in eukaryotic cells [137], a collision event may result in the complete displacement of one or several Okazaki fragments (Ci–ii). Subsequent nascent strand ligation will achieve successful termination in both cases (D).

The severity of the effects observed in recG mutants begs the question of how replication is brought to an end in eukaryotic cells. It follows from the multiple origins per chromosome that a high frequency of replication fork collisions is the norm, but there is no evidence of the pathology described here. The replicative helicase appears to have the opposite polarity to E. coli DnaB [133–135]. However, it is not clear whether it encircles the (single stranded) leading strand template, similar to the mode of function of DnaB in E. coli, or whether it encircles double-stranded DNA [133]. In the latter case the problem might never arise. However, even in the case that only the leading strand template is encircled, any strand displacement when forks merge would involve the nascent lagging strand and create a 5 flap (Fig. 9Ai–iii) rather than a 3 flap, and would likely be processed via the flap endonuclease, FEN-1 [136] (Fig. 9Bi–ii). Alternatively, given Okazaki fragments in eukaryotes are short [137], the helicase might simply unwind one or more unligated Okazaki fragments completely, leaving no flap (Fig. 9Ci–ii). Thus, the difference in polarity of the replicative helicase may avoid the pathological replication we suggest is a significant risk in bacteria.

References [1] S.D. Vincent, A.A. Mahdi, R.G. Lloyd, The RecG branch migration protein of Escherichia coli dissociates R-loops, J. Mol. Biol. 264 (1996) 713–721. [2] A. Fukuoh, H. Iwasaki, K. Ishioka, H. Shinagawa, ATP-dependent resolution of R-loops at the ColE1 replication origin by Escherichia coli RecG protein, a Holliday junction-specific helicase, EMBO J. 16 (1997) 203–209. [3] P McGlynn, A.A. Al-Deib, J. Liu, K.J. Marians, R.G. Lloyd, The DNA replication protein PriA and the recombination protein RecG bind D-loops, J. Mol. Biol. 270 (1997) 212–221. [4] M.C. Whitby, R.G. Lloyd, Targeting Holliday junctions by the RecG branch migration protein of Escherichia coli, J. Biol. Chem. 273 (1998) 19729–19739. [5] E.P. Rocha, E. Cornet, B. Michel, Comparative and evolutionary analysis of the bacterial homologous recombination systems, PLoS Genet. 1 (2005) e15. [6] G.J. Sharples, S.M. Ingleston, R.G. Lloyd, Holliday junction processing in bacteria: insights from the evolutionary conservation of RuvABC, RecG, and RusA, J. Bacteriol. 181 (1999) 5543–5550. [7] R.G Lloyd, Conjugational recombination in resolvase-deficient ruvC mutants of Escherichia coli K-12 depends on recG, J. Bacteriol. 173 (1991) 5414–5418. [8] R.G. Lloyd, C. Buckman, Genetic analysis of the recG locus of Escherichia coli K-12 and of its role in recombination and DNA repair, J. Bacteriol. 173 (1991) 1004–1011. [9] J.D. McCool, S.J. Sandler, Effects of mutations involving cell division, recombination, and chromosome dimer resolution on a priA2::kan mutant, Proc. Natl. Acad. Sci. U.S.A. 98 (2001) 8203–8210. [10] A.V. Gregg, P. McGlynn, R.P. Jaktaji, R.G. Lloyd, Direct rescue of stalled DNA replication forks via the combined action of PriA and RecG helicase activities, Mol. Cell 9 (2002) 241–251. [11] T. Kogoma, Stable DNA replication: interplay between DNA replication, homologous recombination, and transcription, Microbiol. Mol. Biol. Rev. 61 (1997) 212–238. [12] X. Hong, G.W. Cadell, T. Kogoma, Escherichia coli RecG and RecA proteins in R-loop formation, EMBO J. 14 (1995) 2385–2392. [13] C.J. Rudolph, A.L. Upton, L. Harris, R.G. Lloyd, Pathological replication in cells lacking RecG DNA translocase, Mol. Microbiol. 73 (2009) 352–366. [14] G.S. Briggs, A.A. Mahdi, G.R. Weller, Q. Wen, R.G. Lloyd, Interplay between DNA replication, recombination and repair based on the structure of RecG helicase, Philos. Trans. R. Soc. Lond. B Biol. Sci. 359 (2004) 49–59.



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