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Chromosome replication origins: Do we really need them? Be´ne´dicte Michel1)2) and Rolf Bernander3) This article is dedicated to the memory of Dr. Rolf Bernander who passed away in January 2014.
Replication of the main chromosome in the halophilic archaeon Haloferax volcanii was recently reported to continue despite deletion of all active replication origins. Equally surprising, the deletion strain grew faster than the parent strain. It was proposed that origin-less H. volcanii duplicate their chromosomes via recombination-dependent replication. Here, we recall our present knowledge of this mode of chromosome replication in different organisms. We consider the likelihood that it accounts for the viability of H. volcanii deleted for its main specific replication origins, as well as possible alternative interpretations of the results. The selective advantages of having defined chromosome replication origins are discussed from a functional and evolutionary perspective.
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Keywords: archaea; bacteria; RadA; recombination; replication initiation; replication origin
DOI 10.1002/bies.201400003 1) 2) 3)
Centre de Ge´ne´tique Mole´culaire, CNRS, Gif sur Yvette, France Universite´ Paris-Sud, Orsay, France Department of Molecular Biosciences, The Wenner-Gren Institute, Stockholm University, Stockholm, Sweden
*Corresponding author: Be´ne´dicte Michel E-mail: [email protected] Abbreviations: BIR, break-induced replication; cSDR, constitutive stable DNA replication; D-loop, displacement loop; dsDNA, double-stranded DNA; DSB(R), doublestrand break (repair); HT, high throughput; iSDR, induced stable DNA replication; MF, marker frequency; ORC, origin recognition complex; RDR, recombination-dependent replication; R-loops, RNA-loops.
Bioessays 36: 585–590, ß 2014 WILEY Periodicals, Inc.
Introduction In all organisms the genetic material needs to be duplicated (replicated) and then segregated to obtain two viable daughter cells after cell division. In bacteria and archaea, chromosome replication is initiated at specific sites, replication origins [1]. These contain different DNA elements including short repeated sequences to which replication initiator proteins bind. In bacteria the DnaA protein binds to DnaA boxes within the origin [2, 3], while in archaea the corresponding boxes are denoted Orbs (origin recognition boxes) and are recognised by the Orc1/Cdc6 proteins [4]. In both lineages, binding of multiple initiator monomers results in the formation of a nucleoprotein complex that promotes assembly of the rest of the replication machinery, the replisome, and subsequent unwinding of the origin DNA followed by initiation of chromosome replication (Fig. 1) [1]. The situation is more complex in eukaryotic organisms. In yeast, replication initiation also occurs at defined origins, ARSs (autonomously replicating sequences; [5, 6]) while higher eukaryotes, including human, do not appear to utilise specific DNA sequence elements for replication initiation. Instead, initiation is correlated with overall chromatin structure, such that loosely packed DNA regions are more accessible to the origin recognition complex (ORC), while tightly packed chromatin is less prone to initiation [7, 8]. In addition, not all assembled ORCs are used at each replication cycle. Eukaryotic ORCs and archaeal Orc/Cdc6 proteins are homologous, while the bacterial DnaA protein is more distantly related, although similarity to the Orc family is apparent at the structural and functional levels [9]. Chromosome sizes range from less than 0.2 Mb in certain parasitic bacteria to several hundred megabases (Mb) in many eukaryotic species. In all bacteria experimentally investigated to date, chromosomes contain a single replication origin, while eukaryotic replication may initiate from hundreds of positions in order to replicate
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Modes of replication initiation in bacteria DnaA sequence specific: oriC (DnaA boxes)
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Replication initiation in archaea Orc1/Cdc6 sequence specific: oriC
PriA structure specific: •replication fork, •recombination-made D-loop
(orbs)
helicase loader (DnaC in E. coli) helicase (MCM) helicase (DnaB in E. coli) primase primase (DnaG)
polymerase / replication
polymerase III / replication Figure 1. Replication initiation in bacteria and archaea. Replication initiation in bacteria requires binding of the DNA-binding protein DnaA to DnaA-boxes at the chromosome origin oriC. The DnaA initiator melts DNA and forms a nucleoprotein complex. In E. coli, the hexameric helicase DnaB is loaded onto the single-stranded DNA region at oriC by the concerted action of DnaA and DnaC (helicase loader). Two DnaB hexamers loaded on the two DNA strands in opposite orientations interact with primase. Primer synthesis drives the binding of the polymerase III holoenzyme and the initiation of bi-directional replication [2, 3]. In archaea, Orc1 binding to orbs motifs in oriC melts DNA and may directly promote the loading of the double-hexameric helicase MCM, which in turn drives bi-directional replication initiation [1]. A second mode of replication initiation in bacteria is independent of DNA sequence but relies on the recognition of a specific structure, either an abandoned replication fork or a D-loop formed by homologous recombination [24, 25]. PriA specifically recognises these DNA structures and together with partner proteins including the helicase loader DnaC, promotes the loading of a DnaB helicase. Only one DnaB hexamer is loaded and replication restart is unidirectional (Fig. 2). DnaA, PriA and the hexameric helicase are ubiquitous in bacteria, as well as various versions of helicase loader proteins. To date, no PriA structural or functional homologue has been identified in archaea or in eukaryotes.
large chromosomes within a limited time period [1]. Among the archaea, some organisms contain a single origin while up to four origins on the same chromosome have been reported in other species [10]. In accordance with general principles for macromolecular biosynthesis, chromosome replication is regulated at the initiation stage, to reduce unwarranted cellular investment of energy, and to prevent excessive initiation events. A second main reason for careful control of initiation is to coordinate replication with cellular growth and thereby ensure that replication is terminated before chromosome segregation which, in turn, needs to be concluded before cell division can occur. Consequently, impairment of the tightly regulated and sophisticated system for chromosome replication initiation usually results in severe consequences, including induction of mechanisms that block downstream cell cycle processes (SOS and checkpoint systems in bacteria and eukaryotes, respectively), aberrant cell morphology owing to loss of coordination with cell division, as well as a significantly reduced growth rate [11].
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Haloferax volcanii main chromosome counts three replication origins
In a recent study, Hawkins et al. [12] reported replication characteristics of the halophilic (salt-loving) archaeon Haloferax volcanii. This organism thrives in environments characterised by high osmotic pressure, and was originally isolated from a sediment sample retrieved from the Dead Sea [13]. Halophilic archaea are usually polyploid and the copy number of the genome in Haloferax cells has been estimated at around 20 [14], which complicates experimental analyses of chromosome replication patterns. The genome is, in addition, distributed among several replicons, including a main chromosome, two megaplasmids and two small plasmids [15, 16]. Consequently, an intricate system for prevention of replicon loss is in operation during the replication, genome segregation and cell division processes. In all complete genome sequences of halophilic archaea, including H. volcanii [15, 16], multiple orc1/cdc6 replication initiation genes have been identified, together with Orb-like repeat elements in the intergenic region immediately upstream of these genes. This has spurred the question of how many of these loci actually represent active replication origins. An early genetic investigation indicated that the main chromosome of H. volcanii contains two active origins [15], recently corroborated by high throughput (HT) sequencing-based marker frequency (MF) analysis [17]. Two active origins have also been mapped in the closely related Haloferax mediterranei species [17], as well as in Haloarcula hispanica [18]. Also in the Hawkins study, HT sequencing-based MF analysis was applied but, in contrast to the previous investigations, three replication origins were mapped to the main chromosome. A fourth origin, from an integrated Bioessays 36: 585–590, ß 2014 WILEY Periodicals, Inc.
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RecA
Replication can be initiated from recombination intermediates in several organisms
Chi
exo
RecA loading
strand exchange
RecBCD
resolution
RuvABC
replisome reloading
PriA Pol
replication restart Pol Figure 2. Model for the concerted action of double-strand break repair and replication restart proteins. RecBCD degrades double-stranded DNA ends until it encounters a Chi site. Its helicase-nuclease activity is then modified to produce a 30 single-stranded DNA on which RecBCD loads RecA. The synaptic step (homology search and strand exchange) is performed by RecA and results in the formation of a Holliday junction (X structure) adjacent to a D-loop. The post-synaptic steps are the migration and resolution of the Holliday junction, migration being performed by RuvAB and resolution by RuvABC. PriA binding to the D-loop promotes loading of a new helicase, which, in turn, triggers the binding of a new replisome. In E. coli, RecBCD-mediated recombination is always coupled with PriA-dependent replication restart. The red and blue lines are the two homologous recombining molecules and arrowheads indicate the 30 ends of the DNA strands. Indented pink circle: RecBCD; small yellow circles: RecA; big yellow oval: Pol III.
plasmid, was also found to be present in certain isolates. The authors then proceeded to create deletion mutants, demonstrating that each origin could be individually deleted, in most cases without major effects on the growth rate of liquid Bioessays 36: 585–590, ß 2014 WILEY Periodicals, Inc.
Certain types of origin-independent replication are dependent on homologous recombination, mediated by the RecA protein in bacteria, UvsW in bacteriophage T4, and Rad51 in eukaryotes [20]. A possible role of the archaeal RadA recombinase, the ortholog of these proteins, was therefore tested in H. volcanii. In contrast with recA inactivation in bacteria, H. volcanii radA gene deletion is in itself highly deleterious for growth. However, it allows viability while it was found to be lethal for the originless strain, where replication was thus proposed to depend on homologous recombination for initiation. At the molecular level, recombination-dependent replication (RDR) starts by the formation of a DNA double-stranded end, followed by the resection of the 50 DNA end to form a 30 extension on which the recombinase can bind to form a nucleoprotein filament. This filament is competent for homology search, strand-invasion and strand-exchange with a homologous double-stranded DNA molecule. In RDR, the D-loop formed by strand-exchange is used for the assembly of a replisome and replication initiation (Fig. 2). RDR was first discovered in bacteriophage T4 [21]. During infection of Escherichia coli by T4 and certain other bacteriophages, origin-dependent replication is programmed to cease and RDR becomes the main pathway for phage multiplication. RDR creates a network of chromosomes that needs specific T4-encoded enzymes for Holliday junction branch migration and Y-structure debranching, allowing the disentangling and packaging of phage DNA molecules [22, 21]. Therefore, it can be predicted that if H. volcanii lacking all its main origins relies on this mode of replication, disentangling of recombination intermediates should be essential for viability. Following the discovery of a replication restart machinery in E. coli, it was found that the key protein for this process, PriA, is essential for double-strand break repair (DSBR) by homologous recombination [23, 24]. It was proposed that replication initiated from homologous recombination intermediates (D-loops, Fig. 1) is essential for the completion of the repair process, and the reaction was later fully reconstituted in vitro [25]. The tight coupling between DSBR
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batch cultures. Similar results have been obtained in studies in which each of the two replication origins in H. hispanica chromosome [18], or three origins in the crenarchaeal Sulfolobus islandicus species [19], was individually rendered inactive by deletion of the specific orc1/cdc6 gene adjacent to each particular origin. The big surprise in the Hawkins study is the observation that replication of the main chromosome in H. volcanii continues even when all replication origins are deleted. In contrast, in both the H. hispanica and S. islandicus studies, at least one active origin was required for chromosome replication and cell survival [18, 19]. Furthermore, and equally surprising, not only did the H. volcanii cells survive, but they actually grew faster than the parent strain, as determined by OD measurements and competition assays in liquid culture.
repair of ds DNA ends helicase
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by homologous recombination and replication initiation was suggested to have evolved because DSBs occur predominantly at replication forks, and this coupling allows replication to restart from where it was interrupted. RDR was proposed to account for a mode of replication of the E. coli chromosome called induced stable DNA replication (iSDR [26]). iSDR is observed after induction of the SOS response, i.e. the induction of a set of DNA repair genes following DNA damage. iSDR takes place in the absence of the chromosomal origin, in the absence of protein synthesis, and in mutants deprived of the initiator protein DnaA. iSDR initiation requires DSBR proteins (RecBCD, RecA, Holliday junction resolvases) and the replication restart protein PriA. A second type of origin-independent replication in E. coli is called constitutive stable DNA replication (cSDR). It was originally observed in a mutant that lacks RNase H (rnh), in which R-loops can form during transcription by annealing of RNA with the complementary DNA strand, and can be used for PriA-dependent replication initiation [27]. rnh dnaA double mutants initiate DNA replication from different regions of the chromosome, a reaction that requires RecA [28, 29]. However, it was verified in the Hawkins et al. study that all rnh genes are intact in the origin-deleted strain, excluding this hypothesis. RDR was also proposed to account for origin-independent initiation of replication in a recG mutant, in which erroneous merging of replication forks in the terminus region leads to initiation of divergent replication forks via homologous recombination [30], but this process requires replication initiation in the first place, to create the forks that will be converted into a recombination-dependent origin when they meet at the terminus. To date, uncoordinated replication initiation from nonoriC sites in bacteria has been shown to be detrimental for growth and viability. Actually, iSDR does not allow the formation of colonies while the rnh dnaA mutant forms slowgrowing colonies. Replication initiation from homologous recombination intermediates has also been described in yeast, known as break-induced replication (BIR [31, 32]). As in bacteria, BIR/RDR starts by homologous recombination from a DNA double-strand end, followed by replication initiation at the D-loop formed by strand invasion. BIR is thought to occur when the second end of a DNA DSB is not available. It allows replication of megabases of DNA, and uses the normal replication fork polymerases and helicase [33]. However, BIR is independent of pre-replicative complex proteins, required for origin-dependent initiation [33]. In contrast with origin-dependent replication, BIR is highly mutagenic: recombination-initiated replication switches template efficiently, resulting in gross chromosome rearrangements, deletions, insertions and point mutations [34–36].
H. volcanii lacking the three main chromosome replication origins is viable In the following, selected results and inferences from the Hawkins study are discussed and compared to other observations and previously characterised systems, as outlined above.
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Three active replication origins were identified in the main chromosome, whereas two were mapped in an independent study, as well as in two other related haloarchaeal species [17, 18]. As many as ten origins have been suggested in a theoretical study, as well as 15 Cdc6 proteins [4]. Whether these disparities might result from biological (e.g. strain or growth phase) or technical differences (e.g. sequencing technology) is not clear at present, but it has to be noted that the Hawkins et al. study, which identified a third origin called oriC3 as a weak origin, was performed at the physiological temperature for H. volcanii, 45 ˚C, while the study that did not detect this third origin was performed at room temperature. The MF peak corresponding to oriC3 is lower in height as compared to other peaks, and the authors deduce that oriC3 is less frequently used than the other origins. Paradoxically, oriC3 deletion had a more pronounced effect on growth rate than deletion of any of the other origins, for reasons that are unknown at present. As also commented upon by the authors, the peak is not associated with a shift in GC skew polarity, as is usually the case for chromosome replication origins, which suggests late acquirement of oriC3 during evolutionary processes. Replication did not appear to be uniform across the chromosome in the various deletion mutants, as evidenced by their MF gradients. Uneven replication of different chromosome regions, in combination with the randomness of initiation of RDR, would be expected to generate difficulties in resolving fully replicated chromosomes before cell division, with consequent deleterious effects on division and growth rate as usually observed in RDR. The essentiality of the radA gene when all origins had been deleted was taken as evidence that the strain had become dependent upon homologous recombination for replication initiation. However, RadA could also be needed during the elongation stage, if the use of non-canonical origins creates collisions between replication and other processes such as transcription, together with loss of coordination with cellular growth. An interesting related question is how RadA-dependent replication would be prevented from interfering with replication in wild-type cells. It was proposed that the replicative helicase MCM complexes are titrated by active origins, preventing RDR in wild-type cells, a hypothesis that could be tested by over-producing MCM. Finally, the genome copy number was reported to be unchanged, raising the question of how RDR could be regulated to generate on average one replication event per replicon per cell generation. In general, the putative existence of an archaeal version of RDR is of significant interest but it would deserve additional experimental support. For example, RDR implies that viability should be dependent on Holliday junction resolution or on debranching enzymes. Also, profiles are less sharp in all strains where one ori is deleted, which is interpreted as RadA-dependent replication taking place. This hypothesis could be tested by MF analysis of delta-oriC1 radA or delta-oriC2 radA mutants, as the observation of sharper profiles at the remaining origins when RadA is absent would be expected. An alternative explanation for the replication and survival of the origin-deleted strain is activation of cryptic or dormant origins, such as the two remaining non-active loci Bioessays 36: 585–590, ß 2014 WILEY Periodicals, Inc.
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Replication origins are essential for replication initiation control and shape chromosome sequences With the exception of certain bacteriophages and viruses, all organisms duplicate their chromosome(s) by bi-directional replication from sites that are determined by the assembly of a pre-replicative complex on a specific or non-specific DNA sequence. The temporal control of the assembly of prereplicative complexes and, in the case of eukaryotes, the temporal control of their use, ensure the coordination of chromosome replication with the cell cycle, so that each chromosome is duplicated once and only once before cell division. In bacteria, over-replication in a dnaA mutant called dnaAcos causes increased DNA double-strand break formation by over-replication, resulting in loss of cell viability [38, 39]. Mutations in genes that control replication initiation increase the sensitivity to DNA damaging agents [40], although the inactivation of one of the multiple regulation systems for replication initiation may not be sufficient to affect growth in otherwise wild-type strains and in the absence of genotoxic treatment [41]. In eukaryotes, in which the number and position of active origins is not determined by a specific sequence but by several other factors, disrupting the temporal regulation of replication initiation is also detrimental for growth ([42] and references therein). The second evolutionary advantage of replication origins is their contribution to chromosome organisation. On circular chromosomes such as in bacteria, the presence of a fixed replication origin ensures the meeting of the two replication forks in a specific region called the terminus, and triggers several levels of chromosome organisation: (i) genes closer to the origin are more highly expressed (gene dosage effect), (ii) replication and transcription are most often co-directional, which improves gene stability ([43] and references therein), and (iii) the chromosome DNA sequence bears short DNA motifs that can be oriented compared to the replication direction, can be specific for the origin or terminus region, and play a role in DNA recombination and/or chromosome segregation [44]. For example, partition sites (ParS) are often close to the origin. In the terminus region, Ter sites create a Bioessays 36: 585–590, ß 2014 WILEY Periodicals, Inc.
replication fork trap in which replication forks can enter but from which they cannot exit, and a terminus-specific site called MatS governs the structure and dynamic properties of a 1 Mb terminus domain. In contrast, Noc sites that prevent septum closure on DNA are specifically absent from the terminus region. Chi sites that stimulate homologous recombination are over-represented in the orientation that favours the repair of DNA double-stand breaks occurring at replication forks, and Kops sites, recognised by the septum protein FtsK, are oriented to direct the segregation of chromosomes into two daughter cells. In eukaryotes, although replication termination and initiation regions are broader than in prokaryotes, replication and transcription directions are also biased towards co-directionality [45]. In archaea, replication-biased genome organisation has also been reported [46].
Conclusion At least in the long-term – if not within a few generations under laboratory conditions – it is clear that replication initiation from locations that have not been selected during evolution, and cell cycle-independent initiation, are bound to decrease fitness. The provocative work by Hawkins et al. raises several interesting questions. Either an archaeal version of RDR can replicate chromosomes efficiently and even improve growth rate, or dormant or newly integrated origins are responsible for the viability of H. volcanii deleted of its main replication origins. If confirmed, any of these two hypotheses would be highly interesting. According to the former, H. volcanii would be the first example of a free-living organism able to duplicate its chromosome from recombination intermediates only. And the latter hypothesis proposes that unidentified sequences could turn into active replication origins if needed.
Acknowledgements We are very grateful to Dr. Linda Sperling for helpful reading of the manuscript. We thank Dr. Thorsten Allers for helpful discussion. This work was supported by grant 621-2010-5551 from the Swedish Research Council to R.B., and ANR 11-BSV-006-01 from the Agence Nationale de la Recherche to B.M.
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containing orc1/cdc6 genes with immediately adjacent Orblike repeats [16, 15]. Indeed, in the origin-deleted strains the orc1/cdc6 genes adjacent to the deleted origins are still intact. If different dormant origins were randomly activated in different cells in the population, a flatter profile would be obtained as compared to a situation in which only one or a few origins were used by all cells. Deletion of all putative origin regions would provide a means to investigate this possibility. Alternatively, replication in the origin-deleted strain could result from the RadA-dependent insertion of a mini-chromosome or plasmid origin, via homologous recombination between IS sequences for example. The strain used in the Hawkins et al. study harbours several replicons and has the particularity of carrying a plasmid insertion on the main chromosome; origin capture was also previously reported in Sulfolobus [37].
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7. Mesner LD, Valsakumar V, Cieslik M, Pickin R, et al. 2013. Bubble-seq analysis of the human genome reveals distinct chromatin-mediated mechanisms for regulating early- and late-firing origins. Genome Res 23: 1774–88. 8. Mechali M, Yoshida K, Coulombe P, Pasero P. 2013. Genetic and epigenetic determinants of DNA replication origins, position and activation. Curr Opin Genet Dev 23: 124–31. 9. Kawakami H, Katayama T. 2010. DnaA, ORC, and Cdc6: similarity beyond the domains of life and diversity. Biochem Cell Biol 88: 49–62. 10. Pelve EA, Lindas AC, Knoppel A, Mira A, et al. 2012. Four chromosome replication origins in the archaeon Pyrobaculum calidifontis. Mol Microbiol 85: 986–95. 11. Mankouri HW, Huttner D, Hickson ID. 2013. How unfinished business from S-phase affects mitosis and beyond. EMBO J 32: 2661–71. 12. Hawkins M, Malla S, Blythe MJ, Nieduszynski CA, et al. 2013. Accelerated growth in the absence of DNA replication origins. Nature 503: 544–7. 13. Mullakhanbhai MF, Larsen H. 1975. Halobacterium volcanii spec. nov., a Dead Sea halobacterium with a moderate salt requirement. Arch Microbiol 104: 207–14. 14. Breuert S, Allers T, Spohn G, Soppa J. 2006. Regulated polyploidy in halophilic archaea. PLoS One 1: e92. 15. Norais C, Hawkins M, Hartman AL, Eisen JA, et al. 2007. Genetic and physical mapping of DNA replication origins in Haloferax volcanii. PLoS Genet 3: e77. 16. Hartman AL, Norais C, Badger JH, Delmas S, et al. 2010. The complete genome sequence of Haloferax volcanii DS2, a model archaeon. PLoS One 5: e9605. 17. Pelve EA, Martens-Habbena W, Stahl DA, Bernander R. 2013. Mapping of active replication origins in vivo in thaum- and euryarchaeal replicons. Mol Microbiol 90: 538–50. 18. Wu Z, Liu J, Yang H, Liu H, et al. 2014. Multiple replication origins with diverse control mechanisms in Haloarcula hispanica. Nucleic Acids Res 42: 2282–94. 19. Samson RY, Xu Y, Gadelha C, Stone TA, et al. 2013. Specificity and function of archaeal DNA replication initiator proteins. Cell Rep 3: 485–96. 20. Yang S, VanLoock MS, Yu X, Egelman EH. 2001. Comparison of bacteriophage T4 UvsX and human Rad51 filaments suggests that RecAlike polymers may have evolved independently. J Mol Biol 312: 999–1009. 21. Mosig G. 1998. Recombination and recombination-dependent DNA replication in bacteriophage T4. Annu Rev Genet 32: 379–413. 22. Liu J, Morrical SW. 2010. Assembly and dynamics of the bacteriophage T4 homologous recombination machinery. Virol J 7: 357. 23. Kogoma T, Cadwell GW, Barnard KG, Asai T. 1996. The DNA replication priming protein, PriA, is required for homologous recombination and double-strand break repair. J Bacteriol 178: 1258–64. 24. Sandler SJ, Samra HS, Clark AJ. 1996. Differential suppression of priA2: kan phenotypes in Escherichia coli K-12 by mutations in priA, lexA, and dnaC. Genetics 143: 5–13. 25. Xu L, Marians KJ. 2003. PriA mediates DNA replication pathway choice at recombination intermediates. Mol Cell 11: 817–26. 26. Kogoma T. 1997. Stable DNA replication: interplay between DNA replication, homologous recombination, and transcription. Microbiol Mol Biol Rev 61: 212–38.
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Bioessays 36: 585–590, ß 2014 WILEY Periodicals, Inc.
Chromosome replication origins: Do we really need them? Be´ne´dicte Michel1)2) and Rolf Bernander3) This article is dedicated to the memory of Dr. Rolf Bernander who passed away in January 2014.
Replication of the main chromosome in the halophilic archaeon Haloferax volcanii was recently reported to continue despite deletion of all active replication origins. Equally surprising, the deletion strain grew faster than the parent strain. It was proposed that origin-less H. volcanii duplicate their chromosomes via recombination-dependent replication. Here, we recall our present knowledge of this mode of chromosome replication in different organisms. We consider the likelihood that it accounts for the viability of H. volcanii deleted for its main specific replication origins, as well as possible alternative interpretations of the results. The selective advantages of having defined chromosome replication origins are discussed from a functional and evolutionary perspective.
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Keywords: archaea; bacteria; RadA; recombination; replication initiation; replication origin
DOI 10.1002/bies.201400003 1) 2) 3)
Centre de Ge´ne´tique Mole´culaire, CNRS, Gif sur Yvette, France Universite´ Paris-Sud, Orsay, France Department of Molecular Biosciences, The Wenner-Gren Institute, Stockholm University, Stockholm, Sweden
*Corresponding author: Be´ne´dicte Michel E-mail: [email protected] Abbreviations: BIR, break-induced replication; cSDR, constitutive stable DNA replication; D-loop, displacement loop; dsDNA, double-stranded DNA; DSB(R), doublestrand break (repair); HT, high throughput; iSDR, induced stable DNA replication; MF, marker frequency; ORC, origin recognition complex; RDR, recombination-dependent replication; R-loops, RNA-loops.
Bioessays 36: 585–590, ß 2014 WILEY Periodicals, Inc.
Introduction In all organisms the genetic material needs to be duplicated (replicated) and then segregated to obtain two viable daughter cells after cell division. In bacteria and archaea, chromosome replication is initiated at specific sites, replication origins [1]. These contain different DNA elements including short repeated sequences to which replication initiator proteins bind. In bacteria the DnaA protein binds to DnaA boxes within the origin [2, 3], while in archaea the corresponding boxes are denoted Orbs (origin recognition boxes) and are recognised by the Orc1/Cdc6 proteins [4]. In both lineages, binding of multiple initiator monomers results in the formation of a nucleoprotein complex that promotes assembly of the rest of the replication machinery, the replisome, and subsequent unwinding of the origin DNA followed by initiation of chromosome replication (Fig. 1) [1]. The situation is more complex in eukaryotic organisms. In yeast, replication initiation also occurs at defined origins, ARSs (autonomously replicating sequences; [5, 6]) while higher eukaryotes, including human, do not appear to utilise specific DNA sequence elements for replication initiation. Instead, initiation is correlated with overall chromatin structure, such that loosely packed DNA regions are more accessible to the origin recognition complex (ORC), while tightly packed chromatin is less prone to initiation [7, 8]. In addition, not all assembled ORCs are used at each replication cycle. Eukaryotic ORCs and archaeal Orc/Cdc6 proteins are homologous, while the bacterial DnaA protein is more distantly related, although similarity to the Orc family is apparent at the structural and functional levels [9]. Chromosome sizes range from less than 0.2 Mb in certain parasitic bacteria to several hundred megabases (Mb) in many eukaryotic species. In all bacteria experimentally investigated to date, chromosomes contain a single replication origin, while eukaryotic replication may initiate from hundreds of positions in order to replicate
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Modes of replication initiation in bacteria DnaA sequence specific: oriC (DnaA boxes)
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Replication initiation in archaea Orc1/Cdc6 sequence specific: oriC
PriA structure specific: •replication fork, •recombination-made D-loop
(orbs)
helicase loader (DnaC in E. coli) helicase (MCM) helicase (DnaB in E. coli) primase primase (DnaG)
polymerase / replication
polymerase III / replication Figure 1. Replication initiation in bacteria and archaea. Replication initiation in bacteria requires binding of the DNA-binding protein DnaA to DnaA-boxes at the chromosome origin oriC. The DnaA initiator melts DNA and forms a nucleoprotein complex. In E. coli, the hexameric helicase DnaB is loaded onto the single-stranded DNA region at oriC by the concerted action of DnaA and DnaC (helicase loader). Two DnaB hexamers loaded on the two DNA strands in opposite orientations interact with primase. Primer synthesis drives the binding of the polymerase III holoenzyme and the initiation of bi-directional replication [2, 3]. In archaea, Orc1 binding to orbs motifs in oriC melts DNA and may directly promote the loading of the double-hexameric helicase MCM, which in turn drives bi-directional replication initiation [1]. A second mode of replication initiation in bacteria is independent of DNA sequence but relies on the recognition of a specific structure, either an abandoned replication fork or a D-loop formed by homologous recombination [24, 25]. PriA specifically recognises these DNA structures and together with partner proteins including the helicase loader DnaC, promotes the loading of a DnaB helicase. Only one DnaB hexamer is loaded and replication restart is unidirectional (Fig. 2). DnaA, PriA and the hexameric helicase are ubiquitous in bacteria, as well as various versions of helicase loader proteins. To date, no PriA structural or functional homologue has been identified in archaea or in eukaryotes.
large chromosomes within a limited time period [1]. Among the archaea, some organisms contain a single origin while up to four origins on the same chromosome have been reported in other species [10]. In accordance with general principles for macromolecular biosynthesis, chromosome replication is regulated at the initiation stage, to reduce unwarranted cellular investment of energy, and to prevent excessive initiation events. A second main reason for careful control of initiation is to coordinate replication with cellular growth and thereby ensure that replication is terminated before chromosome segregation which, in turn, needs to be concluded before cell division can occur. Consequently, impairment of the tightly regulated and sophisticated system for chromosome replication initiation usually results in severe consequences, including induction of mechanisms that block downstream cell cycle processes (SOS and checkpoint systems in bacteria and eukaryotes, respectively), aberrant cell morphology owing to loss of coordination with cell division, as well as a significantly reduced growth rate [11].
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Haloferax volcanii main chromosome counts three replication origins
In a recent study, Hawkins et al. [12] reported replication characteristics of the halophilic (salt-loving) archaeon Haloferax volcanii. This organism thrives in environments characterised by high osmotic pressure, and was originally isolated from a sediment sample retrieved from the Dead Sea [13]. Halophilic archaea are usually polyploid and the copy number of the genome in Haloferax cells has been estimated at around 20 [14], which complicates experimental analyses of chromosome replication patterns. The genome is, in addition, distributed among several replicons, including a main chromosome, two megaplasmids and two small plasmids [15, 16]. Consequently, an intricate system for prevention of replicon loss is in operation during the replication, genome segregation and cell division processes. In all complete genome sequences of halophilic archaea, including H. volcanii [15, 16], multiple orc1/cdc6 replication initiation genes have been identified, together with Orb-like repeat elements in the intergenic region immediately upstream of these genes. This has spurred the question of how many of these loci actually represent active replication origins. An early genetic investigation indicated that the main chromosome of H. volcanii contains two active origins [15], recently corroborated by high throughput (HT) sequencing-based marker frequency (MF) analysis [17]. Two active origins have also been mapped in the closely related Haloferax mediterranei species [17], as well as in Haloarcula hispanica [18]. Also in the Hawkins study, HT sequencing-based MF analysis was applied but, in contrast to the previous investigations, three replication origins were mapped to the main chromosome. A fourth origin, from an integrated Bioessays 36: 585–590, ß 2014 WILEY Periodicals, Inc.
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RecA
Replication can be initiated from recombination intermediates in several organisms
Chi
exo
RecA loading
strand exchange
RecBCD
resolution
RuvABC
replisome reloading
PriA Pol
replication restart Pol Figure 2. Model for the concerted action of double-strand break repair and replication restart proteins. RecBCD degrades double-stranded DNA ends until it encounters a Chi site. Its helicase-nuclease activity is then modified to produce a 30 single-stranded DNA on which RecBCD loads RecA. The synaptic step (homology search and strand exchange) is performed by RecA and results in the formation of a Holliday junction (X structure) adjacent to a D-loop. The post-synaptic steps are the migration and resolution of the Holliday junction, migration being performed by RuvAB and resolution by RuvABC. PriA binding to the D-loop promotes loading of a new helicase, which, in turn, triggers the binding of a new replisome. In E. coli, RecBCD-mediated recombination is always coupled with PriA-dependent replication restart. The red and blue lines are the two homologous recombining molecules and arrowheads indicate the 30 ends of the DNA strands. Indented pink circle: RecBCD; small yellow circles: RecA; big yellow oval: Pol III.
plasmid, was also found to be present in certain isolates. The authors then proceeded to create deletion mutants, demonstrating that each origin could be individually deleted, in most cases without major effects on the growth rate of liquid Bioessays 36: 585–590, ß 2014 WILEY Periodicals, Inc.
Certain types of origin-independent replication are dependent on homologous recombination, mediated by the RecA protein in bacteria, UvsW in bacteriophage T4, and Rad51 in eukaryotes [20]. A possible role of the archaeal RadA recombinase, the ortholog of these proteins, was therefore tested in H. volcanii. In contrast with recA inactivation in bacteria, H. volcanii radA gene deletion is in itself highly deleterious for growth. However, it allows viability while it was found to be lethal for the originless strain, where replication was thus proposed to depend on homologous recombination for initiation. At the molecular level, recombination-dependent replication (RDR) starts by the formation of a DNA double-stranded end, followed by the resection of the 50 DNA end to form a 30 extension on which the recombinase can bind to form a nucleoprotein filament. This filament is competent for homology search, strand-invasion and strand-exchange with a homologous double-stranded DNA molecule. In RDR, the D-loop formed by strand-exchange is used for the assembly of a replisome and replication initiation (Fig. 2). RDR was first discovered in bacteriophage T4 [21]. During infection of Escherichia coli by T4 and certain other bacteriophages, origin-dependent replication is programmed to cease and RDR becomes the main pathway for phage multiplication. RDR creates a network of chromosomes that needs specific T4-encoded enzymes for Holliday junction branch migration and Y-structure debranching, allowing the disentangling and packaging of phage DNA molecules [22, 21]. Therefore, it can be predicted that if H. volcanii lacking all its main origins relies on this mode of replication, disentangling of recombination intermediates should be essential for viability. Following the discovery of a replication restart machinery in E. coli, it was found that the key protein for this process, PriA, is essential for double-strand break repair (DSBR) by homologous recombination [23, 24]. It was proposed that replication initiated from homologous recombination intermediates (D-loops, Fig. 1) is essential for the completion of the repair process, and the reaction was later fully reconstituted in vitro [25]. The tight coupling between DSBR
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batch cultures. Similar results have been obtained in studies in which each of the two replication origins in H. hispanica chromosome [18], or three origins in the crenarchaeal Sulfolobus islandicus species [19], was individually rendered inactive by deletion of the specific orc1/cdc6 gene adjacent to each particular origin. The big surprise in the Hawkins study is the observation that replication of the main chromosome in H. volcanii continues even when all replication origins are deleted. In contrast, in both the H. hispanica and S. islandicus studies, at least one active origin was required for chromosome replication and cell survival [18, 19]. Furthermore, and equally surprising, not only did the H. volcanii cells survive, but they actually grew faster than the parent strain, as determined by OD measurements and competition assays in liquid culture.
repair of ds DNA ends helicase
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by homologous recombination and replication initiation was suggested to have evolved because DSBs occur predominantly at replication forks, and this coupling allows replication to restart from where it was interrupted. RDR was proposed to account for a mode of replication of the E. coli chromosome called induced stable DNA replication (iSDR [26]). iSDR is observed after induction of the SOS response, i.e. the induction of a set of DNA repair genes following DNA damage. iSDR takes place in the absence of the chromosomal origin, in the absence of protein synthesis, and in mutants deprived of the initiator protein DnaA. iSDR initiation requires DSBR proteins (RecBCD, RecA, Holliday junction resolvases) and the replication restart protein PriA. A second type of origin-independent replication in E. coli is called constitutive stable DNA replication (cSDR). It was originally observed in a mutant that lacks RNase H (rnh), in which R-loops can form during transcription by annealing of RNA with the complementary DNA strand, and can be used for PriA-dependent replication initiation [27]. rnh dnaA double mutants initiate DNA replication from different regions of the chromosome, a reaction that requires RecA [28, 29]. However, it was verified in the Hawkins et al. study that all rnh genes are intact in the origin-deleted strain, excluding this hypothesis. RDR was also proposed to account for origin-independent initiation of replication in a recG mutant, in which erroneous merging of replication forks in the terminus region leads to initiation of divergent replication forks via homologous recombination [30], but this process requires replication initiation in the first place, to create the forks that will be converted into a recombination-dependent origin when they meet at the terminus. To date, uncoordinated replication initiation from nonoriC sites in bacteria has been shown to be detrimental for growth and viability. Actually, iSDR does not allow the formation of colonies while the rnh dnaA mutant forms slowgrowing colonies. Replication initiation from homologous recombination intermediates has also been described in yeast, known as break-induced replication (BIR [31, 32]). As in bacteria, BIR/RDR starts by homologous recombination from a DNA double-strand end, followed by replication initiation at the D-loop formed by strand invasion. BIR is thought to occur when the second end of a DNA DSB is not available. It allows replication of megabases of DNA, and uses the normal replication fork polymerases and helicase [33]. However, BIR is independent of pre-replicative complex proteins, required for origin-dependent initiation [33]. In contrast with origin-dependent replication, BIR is highly mutagenic: recombination-initiated replication switches template efficiently, resulting in gross chromosome rearrangements, deletions, insertions and point mutations [34–36].
H. volcanii lacking the three main chromosome replication origins is viable In the following, selected results and inferences from the Hawkins study are discussed and compared to other observations and previously characterised systems, as outlined above.
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Three active replication origins were identified in the main chromosome, whereas two were mapped in an independent study, as well as in two other related haloarchaeal species [17, 18]. As many as ten origins have been suggested in a theoretical study, as well as 15 Cdc6 proteins [4]. Whether these disparities might result from biological (e.g. strain or growth phase) or technical differences (e.g. sequencing technology) is not clear at present, but it has to be noted that the Hawkins et al. study, which identified a third origin called oriC3 as a weak origin, was performed at the physiological temperature for H. volcanii, 45 ˚C, while the study that did not detect this third origin was performed at room temperature. The MF peak corresponding to oriC3 is lower in height as compared to other peaks, and the authors deduce that oriC3 is less frequently used than the other origins. Paradoxically, oriC3 deletion had a more pronounced effect on growth rate than deletion of any of the other origins, for reasons that are unknown at present. As also commented upon by the authors, the peak is not associated with a shift in GC skew polarity, as is usually the case for chromosome replication origins, which suggests late acquirement of oriC3 during evolutionary processes. Replication did not appear to be uniform across the chromosome in the various deletion mutants, as evidenced by their MF gradients. Uneven replication of different chromosome regions, in combination with the randomness of initiation of RDR, would be expected to generate difficulties in resolving fully replicated chromosomes before cell division, with consequent deleterious effects on division and growth rate as usually observed in RDR. The essentiality of the radA gene when all origins had been deleted was taken as evidence that the strain had become dependent upon homologous recombination for replication initiation. However, RadA could also be needed during the elongation stage, if the use of non-canonical origins creates collisions between replication and other processes such as transcription, together with loss of coordination with cellular growth. An interesting related question is how RadA-dependent replication would be prevented from interfering with replication in wild-type cells. It was proposed that the replicative helicase MCM complexes are titrated by active origins, preventing RDR in wild-type cells, a hypothesis that could be tested by over-producing MCM. Finally, the genome copy number was reported to be unchanged, raising the question of how RDR could be regulated to generate on average one replication event per replicon per cell generation. In general, the putative existence of an archaeal version of RDR is of significant interest but it would deserve additional experimental support. For example, RDR implies that viability should be dependent on Holliday junction resolution or on debranching enzymes. Also, profiles are less sharp in all strains where one ori is deleted, which is interpreted as RadA-dependent replication taking place. This hypothesis could be tested by MF analysis of delta-oriC1 radA or delta-oriC2 radA mutants, as the observation of sharper profiles at the remaining origins when RadA is absent would be expected. An alternative explanation for the replication and survival of the origin-deleted strain is activation of cryptic or dormant origins, such as the two remaining non-active loci Bioessays 36: 585–590, ß 2014 WILEY Periodicals, Inc.
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Replication origins are essential for replication initiation control and shape chromosome sequences With the exception of certain bacteriophages and viruses, all organisms duplicate their chromosome(s) by bi-directional replication from sites that are determined by the assembly of a pre-replicative complex on a specific or non-specific DNA sequence. The temporal control of the assembly of prereplicative complexes and, in the case of eukaryotes, the temporal control of their use, ensure the coordination of chromosome replication with the cell cycle, so that each chromosome is duplicated once and only once before cell division. In bacteria, over-replication in a dnaA mutant called dnaAcos causes increased DNA double-strand break formation by over-replication, resulting in loss of cell viability [38, 39]. Mutations in genes that control replication initiation increase the sensitivity to DNA damaging agents [40], although the inactivation of one of the multiple regulation systems for replication initiation may not be sufficient to affect growth in otherwise wild-type strains and in the absence of genotoxic treatment [41]. In eukaryotes, in which the number and position of active origins is not determined by a specific sequence but by several other factors, disrupting the temporal regulation of replication initiation is also detrimental for growth ([42] and references therein). The second evolutionary advantage of replication origins is their contribution to chromosome organisation. On circular chromosomes such as in bacteria, the presence of a fixed replication origin ensures the meeting of the two replication forks in a specific region called the terminus, and triggers several levels of chromosome organisation: (i) genes closer to the origin are more highly expressed (gene dosage effect), (ii) replication and transcription are most often co-directional, which improves gene stability ([43] and references therein), and (iii) the chromosome DNA sequence bears short DNA motifs that can be oriented compared to the replication direction, can be specific for the origin or terminus region, and play a role in DNA recombination and/or chromosome segregation [44]. For example, partition sites (ParS) are often close to the origin. In the terminus region, Ter sites create a Bioessays 36: 585–590, ß 2014 WILEY Periodicals, Inc.
replication fork trap in which replication forks can enter but from which they cannot exit, and a terminus-specific site called MatS governs the structure and dynamic properties of a 1 Mb terminus domain. In contrast, Noc sites that prevent septum closure on DNA are specifically absent from the terminus region. Chi sites that stimulate homologous recombination are over-represented in the orientation that favours the repair of DNA double-stand breaks occurring at replication forks, and Kops sites, recognised by the septum protein FtsK, are oriented to direct the segregation of chromosomes into two daughter cells. In eukaryotes, although replication termination and initiation regions are broader than in prokaryotes, replication and transcription directions are also biased towards co-directionality [45]. In archaea, replication-biased genome organisation has also been reported [46].
Conclusion At least in the long-term – if not within a few generations under laboratory conditions – it is clear that replication initiation from locations that have not been selected during evolution, and cell cycle-independent initiation, are bound to decrease fitness. The provocative work by Hawkins et al. raises several interesting questions. Either an archaeal version of RDR can replicate chromosomes efficiently and even improve growth rate, or dormant or newly integrated origins are responsible for the viability of H. volcanii deleted of its main replication origins. If confirmed, any of these two hypotheses would be highly interesting. According to the former, H. volcanii would be the first example of a free-living organism able to duplicate its chromosome from recombination intermediates only. And the latter hypothesis proposes that unidentified sequences could turn into active replication origins if needed.
Acknowledgements We are very grateful to Dr. Linda Sperling for helpful reading of the manuscript. We thank Dr. Thorsten Allers for helpful discussion. This work was supported by grant 621-2010-5551 from the Swedish Research Council to R.B., and ANR 11-BSV-006-01 from the Agence Nationale de la Recherche to B.M.
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