Abstract There has long been a fascination in the DNA repair pathways of archaea, for two main reasons. Firstly, many archaea inhabit extreme environments where the rate of physical damage to DNA is accelerated. These archaea might reasonably be expected to have particularly robust or novel DNA repair pathways to cope with this. Secondly, the archaea have long been understood to be a lineage distinct from the bacteria, and to share a close relationship with the eukarya, particularly in their information processing systems. Recent discoveries suggest the eukarya arose from within the archaeal domain, and in particular from lineages related to the TACK superphylum and Lokiarchaea. Thus, archaeal DNA repair proteins and pathways can represent a useful model system. This review focuses on recent advances in our understanding of archaeal DNA repair processes including base excision repair, nucleotide excision repair, mismatch repair and double-strand break repair. These advances are discussed in the context of the emerging picture of the evolution and relationship of the three domains of life. Archaea, DNA repair, homologous recombination INTRODUCTION Although double-stranded DNA is a stable, chemically inert molecule, damage to DNA is largely unavoidable, and can have serious consequences for a cell, including mutation and death. While some level of mutation is acceptable, and indeed constitutes the raw material for evolution, high mutational load is incompatible with life. Efficient repair of DNA damage is therefore essential for all forms of life. The archaea are no exception, and indeed they often inhabit challenging environments and are thus exposed to extremes of temperature, salinity, pressure or pH. Archaea would thus be expected to have particularly robust DNA repair pathways, and they do, but we do not yet understand them very well. As has been noted in previous reviews of the topic, there are many enigmas in the field of archaeal DNA repair (Grogan 1998, 2015; White 2003; Rouillon and White 2011). Some of these are gradually being resolved whilst others remain stubbornly opaque. In this review, we focus on recent research that illuminates aspects of the four universal DNA repair pathways: base excision repair (BER), nucleotide excision repair (NER), mismatch repair (MMR) and homologous recombination/double-strand break repair (HR/DSBR) (Fig. 1). Figure 1. View largeDownload slide Schematic of DNA damage causes, consequences and repair pathways. Further details are found in the main text. Figure 1. View largeDownload slide Schematic of DNA damage causes, consequences and repair pathways. Further details are found in the main text. The last few years have seen rapid advances in several areas. Genomics has given us vast new datasets and unveiled a diverse array of new archaeal species that are shaking our view of the tree of life (Adam et al.2017; Spang, Caceres and Ettema 2017). Genetic systems are being developed for key model organisms such as the Halophiles, Methanogens, Sulfolobales and Thermococcales that allow the increasingly sophisticated study of archaeal gene function (Farkas, Picking and Santangelo 2013). Biochemical and structural studies are revealing mechanistic detail on individual DNA repair proteins and pathways. Used in combination, these approaches can lead to swift and significant advances in understanding. A good example is the discovery of a non-canonical MMR pathway, based on the EndoMS nuclease, by the Ishino lab (Ishino et al.2016). This advance, described in detail below, has the potential to answer one of the major outstanding questions of the archaeal DNA repair field. This is a field in transition. Much of the early work on DNA replication and repair in the archaea arose from a desire to study simpler model systems of eukaryal (ultimately, human) processes. This approach led to many notable successes. However, as the need for model systems has faded, there is a growing realisation that the archaea are not a niche player in the biosphere but rather a major, significant component that deserves study in their own right. Their cellular and molecular biology is often distinct from those of the bacteria and eukarya, and this is certainly true for their DNA repair pathways. DNA repair and the origin of the eukarya Although still not universally agreed, the recent discovery of new archaeal lineages known collectively as the ‘ASGARD’ archaea, which includes the species Lokiarchaeota and Thorarchaeota, has caused a reassessment of the relationship between the archaeal and eukaryal domains (reviewed in Eme et al.2017). The large number of gene families previously thought to be specific to the eukarya that are found in ASGARD genomes has led to the suggestion that Eukarya arose from an archaeal species related to the ASGARD archaea. Other experts however disagree with this interpretation of the data (Da Cunha et al.2017). What can the distribution of DNA repair genes across the archaea add to this hot topic (Fig. 2)? If we take the example of the XPF nuclease, it comes in two ‘flavours’ in archaea. The short version consists only of a nuclease domain, which interacts with PCNA, and is found only in the TACK superphylum (Rouillon and White 2011). The long version has a nuclease fused to a helicase domain matching eukaryal XPF. This is present predominantly in the Euryarchaea, but also in the ASGARD archaea. Similarly, a eukaryal-type replication protein A (RPA, a single-stranded DNA-binding protein) is present in most archaea with the exception of the Crenarchaea and Thermoplasma, which have a short version (Rouillon and White 2011). Focussing on the two examples of ASGARD archaea in Fig. 2, it is apparent that Lokiarchaea and Thorarchaea have the complement of eukaryal-type repair proteins one would expect for an ancestor of the eukarya. This includes copies of the bacterial-type MMR proteins MutS and MutL, which are also present throughout the eukaryal lineage. Intriguingly, the ASGARD archaea have also picked up the bacterial UvrABC NER system. Overall, the distribution pattern of DNA repair genes in the archaea, and the ASGARD lineage in particular, is consistent with the hypothesis that the latter gave rise to the eukaryal domain of life. Figure 2. View largeDownload slide Distribution of DNA repair genes in the archaea. Genus names on the left are organised as members of the TACK superphylum and Euryarchaea. For each genus, a shaded box indicates the presence of the relevant gene. Bacterial genes probably acquired by lateral gene transfer are shown in green, others in blue. Accession numbers are shown in Table S1 (Supporting Information). Figure 2. View largeDownload slide Distribution of DNA repair genes in the archaea. Genus names on the left are organised as members of the TACK superphylum and Euryarchaea. For each genus, a shaded box indicates the presence of the relevant gene. Bacterial genes probably acquired by lateral gene transfer are shown in green, others in blue. Accession numbers are shown in Table S1 (Supporting Information). MISMATCH REPAIR The canonical MutL-MutS pathway MMR is the process by which bases incorporated in error by the DNA replication machinery are detected and corrected. The MutL-MutS MMR pathway first characterised in Escherichia coli is present in most bacteria (with the notable exception of the Actinobacteria) and in the eukarya, but is the exception rather than the rule in the archaea (Kelman and White 2005). Most archaea lack plausible MutS and MutL homologues, and those that have them tend to be temperature mesophiles such as halophiles and methanogens that most likely captured these genes by lateral gene transfer from bacteria (Fig. 2). The mode of inheritance of a bacterial-type MMR pathway from bacteria to the eukarya is a matter of conjecture. One possibility is that endosymbiotic event that led to the evolution of the mitochondrion from an Alpha-proteobacterium allowed the bacterial genes for MMR to become established in the early eukaryal genome. An alternative possibility is that the eukarya inherited the bacterial MMR machinery via their archaeal lineage. It is notable that the ASGARD archaea including Lokiarchaeum and Thorarchaeum, which have been proposed as the most closely related extant archaea to the progenitor of the eukarya (Eme et al.2017), possess clear MutS and MutL homologues. The emerging role of EndoMS The lack of canonical MMR in most archaea is not reflected in high mutation rates (Grogan 2004), and deletion of MutS-MutL in Halobacterium salinarum did not give rise to a hypermutation phenotype (Busch and DiRuggiero 2010). These observations suggest that alternative pathways exist to detect and remove mismatches post DNA replication. To search for this pathway, Ishino et al. (2016) devised a functional screen for enzymes capable of cleaving DNA mismatches in Pyrococcus furiosus. This resulted in the identification of an enzyme, which was named EndoMS for endonuclease mismatch specific, capable of cleaving a range of mismatched DNAs by the introduction of staggered cleavages in both strands of the DNA, leaving 5 nt 5΄-overhangs (Ishino et al.2016). EndoMS had originally been identified in the Myllykallio lab and named NucS, based on its activity against single-stranded DNA (Ren et al.2009). The structure of NucS revealed a dimeric, two-domain organisation, and the enzyme was shown to form a physical interaction with the sliding clamp PCNA (proliferating cell nuclear antigen) (Ren et al.2009). As the enzyme has a much higher specificity for mismatches than for branched or ssDNA, the nomenclature ‘EndoMS’ will be used henceforth. The recent DNA:protein co-crystal structure reveals that EndoMS wraps around mismatched DNA substrates, flipping out two bases and cleaving the DNA backbone in a manner reminiscent of type II restriction enzymes (Nakae et al.2016) (Fig. 3). The enzyme is active against G-T, G-G, T-T, T-C and A-G mismatches, but not against C-C, A-C or A-A mismatches in vitro (Ishino et al.2016), which is consistent with higher binding affinities for substrates with a mismatched G or T (Nakae et al.2016). Figure 3. View largeDownload slide Structure of the EndoMS dimer bound to DNA (Nakae et al.2016). EndoMS subunits are shown in cyan and green, with the N-terminal dimerisation domain at the top and the C-terminal nuclease domains at the bottom. The two catalytic sites are indicated by the green spheres that denote the active site magnesium ions. The DNA duplex (blue) is distorted by EndoMS binding and two bases are flipped out. Figure 3. View largeDownload slide Structure of the EndoMS dimer bound to DNA (Nakae et al.2016). EndoMS subunits are shown in cyan and green, with the N-terminal dimerisation domain at the top and the C-terminal nuclease domains at the bottom. The two catalytic sites are indicated by the green spheres that denote the active site magnesium ions. The DNA duplex (blue) is distorted by EndoMS binding and two bases are flipped out. EndoMS has a complex distribution in the archaea (Fig. 2), with examples in the halophiles, various thermophiles from the crenarchaeal and euryarchaeal phyla, and Thorarchaeum from the ASGARD phylum. EndoMS is also present in some bacterial genomes, particularly the phylum Actinobacteria where MutS-MutL is generally absent. A screen for mutation avoidance genes showed that deletion of the gene encoding EndoMS in Mycobacterium smegmatis resulted in a hypermutation phenotype, increasing background mutation rate by about 100-fold (Castaneda-Garcia et al.2017). The higher rates of mutation were due to elevated levels of transitions (A:T to G:C or G:C to A:T), which is a hallmark of an MMR defect, and similar effects were observed when EndoMS was deleted in Streptomyces coelicolor. Mycobacterial EndoMS has no nuclease activity when presented with mismatched DNA substrates in vitro, suggesting that further components in this non-canonical MMR pathway remain to be identified (Castaneda-Garcia et al.2017). Taken together, the studies in archaea and bacteria make a compelling case that EndoMS participates in an MMR pathway. However, many important aspects of this pathway remain to be elucidated. The generation of double-strand breaks (DSB) by P. furiosus EndoMS is suggestive of an MMR process that functions via HR/DSBR (Ishino et al.2016). This has the advantage that there is no need to identify nascent DNA strands to pinpoint the mismatched base, as both will be resected during DSBR. The observation that EndoMS is sometimes found in an operon with the RadA recombinase lends further support to this hypothesis (Ren et al.2009). However, generation of a DSB each time a mismatch is detected seems a risky strategy, unless HR is very efficient. This is probably the case in many of the Euryarchaea, which are highly polyploid. It is much less obvious for the Crenarchaea, which have a eukaryal-like cell cycle with monoploid and diploid stages (Lundgren and Bernander 2007). Clearly, dissection and reconstitution of the pathway using genetic and biochemical techniques is a pressing priority. The interaction of archaeal EndoMS with the sliding clamp PCNA may provide a means to locate EndoMS at the replication fork to interrogate newly synthesised DNA, and could give the opportunity for co-location of a variety of DNA manipulation enzymes on the PCNA toolbelt (Beattie and Bell 2011). In this regard, it will be interesting to see whether the bacterial EndoMS protein requires an interaction with the bacterial sliding clamp for activity. NUCLEOTIDE EXCISION REPAIR NER is a pathway that removes bulky, helix-distorting lesions such as photoproducts from DNA (Fig. 1). Because it does not rely on direct detection of the lesion, but rather the resultant structural perturbation, it can repair many different types of DNA damage. The NER pathway in bacteria is catalysed by the UvrABC proteins, where UvrA is involved in damage recognition, UvrB is the helicase that opens the dsDNA and UvrC the nuclease that cuts on both sides of the lesion. In eukarya, an analogous and more complex pathway exists, which involves damage recognition by XPC-hr23b, DNA opening by transcription factor IIH (TFIIH), subsequent binding of the XPA and RPA proteins, resulting in recruitment of the nucleases XPF-ERCC1 and XPG to cut on either side of the lesion. Archaea have a diverse and, frankly, confusing range of NER proteins encoded in their genomes (Fig. 2) (Rouillon and White 2011). In archaea that have co-opted the bacterial NER genes encoding UvrABC, the bacterial system seems to be dominant for NER. For example, the NER patch repair size of 10–11 bp for Methanothermobacter thermautotrophicus is consistent with UvrABC function (Ogrunc et al.1998). Likewise, deletion of the genes for UvrA, UvrB or UvrC in Halobacterium NRC-1 resulted in a severe UV sensitivity despite the fact that this organism has also homologues of the eukaryal-type NER proteins XPF, XPB and XPD (Crowley et al.2006). Furthermore, there are no recognisable orthologues of the damage recognition proteins XPC and XPA in archaea. The SSB protein, which can melt damaged DNA specifically (Cubeddu and White 2005) and can bind quickly and cooperatively on ssDNA (Morten et al.2015), could conceivably carry out this function. Since most archaea have at least some eukaryal type NER genes, the question of their function is pertinent. Genetic studies of the putative archaeal NER pathway have been limited. Deletion of the XPD and XPB genes in Thermococcus kodakaraensis resulted in only very mild repair phenotypes (Fujikane et al.2010). In contrast, deletion of the XPF homologue Hef in this organism resulted in a marked sensitivity to the crosslinking agent mitomycin C (MMC), methylmethanesulfonate (MMS) and gamma radiation, suggesting an important role for Hef in multiple repair pathways including crosslink repair and replication restart (Fujikane et al.2010). This is consistent with the known roles of the eukaryal XPF and Mus81 proteins, which share a common ancestor with Hef (Rouillon and White 2011). Both the helicase and nuclease activities of Hef were shown to be important, suggesting that Hef needs to unwind and cleave DNA during repair (Fujikane et al.2010). In the crenarchaeon Sulfolobus islandicus, deletion of the XPD, XPB and Bax1 genes has been reported with no resulting phenotype (She et al.2009). Although these results should be viewed as preliminary until published in more detail in a peer reviewed journal, they are consistent with the work in T. kodakaraensis. Overall then, genetic studies have shown that putative NER proteins are not essential, but have not progressed our understanding of the archaeal NER pathway very far. This has led Grogan to speculate that there is no NER pathway per se in archaea lacking UvrABC—raising the possibility that bulky NER-type lesions, which would represent a barrier to the replication fork, are removed by pathways that restart stalled forks (Grogan 2015). Although this is an interesting hypothesis, it does beg the question: Why do most archaea have XPB and XPD genes? After all, they must be doing something. XPD helicase Although we still have a rather limited understanding of archaeal NER, study of the XPD and XPB helicases has nonetheless been quite revealing. XPD is a 5΄ to 3΄ helicase with an essential iron-sulfur cluster (Rudolf et al.2006). In eukarya, XPD exists in the 10-subunit transcription factor TFIIH, along with the XPB helicase. TFIIH is involved in both NER, where DNA around a lesion is unwound, and transcription initiation, where RNA polymerase II promoters are unwound. XPD is essential for DNA unwinding in NER, but its activity is not required in transcription (Kuper et al.2014). Until recently, TFIIH was difficult to study at a structural level and the archaeal XPD, which is a monomer, was thus an attractive model system. Three groups independently reported the structure of archaeal XPD (Fan et al.2008; Liu et al.2008; Wolski et al.2008), revealing a four-domain organisation with two motor domains, an Arch and FeS domain (Fig. 4). The mutations that cause the genetic condition xeroderma pigmentosum in humans, which arises from defective NER, could be mapped onto the archaeal XPD structures. The residues targeted by mutation are highly conserved, and cluster in areas involved in the catalytic mechanism of the archaeal enzyme—a striking example of conservation of function spanning the archaeal and eukaryal domains (Liu et al.2008). Figure 4. View largeDownload slide Structure of XPD from T. acidophilum (Constantinescu-Aruxandei et al.2016). Motor domain 1 (MD1) is pink, motor domain 2 (MD2) green, the FeS domain yellow and the Arch domain teal. The covalently bound 5 nt of DNA is shown in blue. The interface between the Arch and FeS domains that must open is indicated, and the central pore through which DNA must pass is labelled. Figure 4. View largeDownload slide Structure of XPD from T. acidophilum (Constantinescu-Aruxandei et al.2016). Motor domain 1 (MD1) is pink, motor domain 2 (MD2) green, the FeS domain yellow and the Arch domain teal. The covalently bound 5 nt of DNA is shown in blue. The interface between the Arch and FeS domains that must open is indicated, and the central pore through which DNA must pass is labelled. In eukaryal NER, XPD has been shown to ‘proofread’ for the presence of a DNA lesion in the translocated strand as a mechanism to increase the specificity of the NER reaction (Mathieu et al.2013). A lesion recognition pocket, close to the FeS cluster and immediately adjacent to the pore through which XPD pulls ssDNA, was identified. Two amino acids, Tyr-192 and Arg-196, were identified as an important part of this pocket, and mutations at these positions reduced DNA repair in a eukaryal system (Mathieu et al.2013). The authors went on to make the same changes in XPD from the archaeon Ferroplasma acidophilum (FacXPD), which correspond to residues Tyr-171 and Lys-175. This enzyme had been shown previously to stall at CPD lesions on the translocated strand (Mathieu, Kaczmarek and Naegeli 2010). They found that mutation of these residues did indeed abrogate the ability of FacXPD to stall at a CPD lesion, although helicase activity was unaffected (Mathieu et al.2013). However, XPD from S. acidocaldarius is not stalled by CPD or extrahelical fluorescein adducts in model substrates (Rudolf et al.2010). This may point to differences in the functions of XPD in the Euryarchaea and Crenarchaea. Both SacXPD and FacXPD display only modest (∼2-fold) increases in binding affinity for damaged versus undamaged DNA (Rudolf et al.2010; Ghoneim and Spies 2014), suggesting that damage recognition, when it occurs, could be subtle. This picture is further complicated by the observation from atomic force microscopy studies that TacXPD binds to extrahelical fluorescein lesions in the translocated strand, but CPDs in the displaced strand (Buechner et al.2014). Furthermore, single molecule studies of FacXPD revealed the unexpected ability of the enzyme to bypass a bound single-strand DNA-binding protein without either protein dissociating from the nucleic acid—a phenomenon that is still not fully understood (Honda et al.2009). Clearly, further work in this area would be desirable to improve our understanding of damage recognition by the XPD helicase. Further studies of archaeal XPDs have revealed mechanistic insights into DNA binding and associated conformational changes. The Kisker lab succeeded in co-crystallising TacXPD with a short piece of ssDNA, demonstrating unequivocally the polarity of unwinding by the enzyme (Kuper et al.2012). The DNA was bound by motor domain 2, and the authors predicted that, since XPD can unwind bubble structures (Rudolf et al.2010) and eukaryal NER functions on DNA without ends, full engagement with DNA would require the opening of the interface between the Arch and FeS domains to allow DNA passage. This hypothesis was confirmed by the observation of transient opening of the interface in a single molecule study by the Spies lab (Ghoneim and Spies 2014) and recently nailed down by a study which covalently closed the interface with a crosslinker (Constantinescu-Aruxandei et al.2016). Constantinescu and co-workers demonstrated that TacXPD can still bind DNA with high affinity when the interface between the Arch and FeS domain is covalently closed, but cannot function as a helicase. They proposed a two-stage binding mechanism for XPD, with ssDNA initially bound tightly by motor domain 2, followed by transient opening of the Arch domain to allow passage through the central pore (Constantinescu-Aruxandei et al.2016). This mechanism is likely to hold true for eukaryal XPD in the context of TFIIH. XPB helicase (or not?) XPB has historically been considered to be a 3΄ to 5΄ DNA helicase; however, the evidence supporting this assignment is rather thin. Helicase activity was ascribed to XPB from Archaeoglobus fulgidus (Fan et al.2006), but was not detected in either XPB protein from S. solfataricus (Richards et al. 2008a). The structure of AfuXPB revealed an unusual conformation, with the motor domains rotated away from the canonical structure by 170°. The structure revealed two accessory domains: damage recognition domain (DRD) and thumb (Fan et al.2006). The White lab reported that XPB is often found in an operon with a protein they named Bax1, and that the two proteins from a 1:1 complex (Richards et al. 2008). Subsequently, Bax1 was shown to be a nuclease (Roth et al.2009), and a detailed study revealed that XPB and Bax1 function in concert to extend bubble structures and cleave DNA (Rouillon and White 2010). The thumb domain was shown to be essential for DNA unwinding by XPB, and the DRD was shown to be essential for the function of the XPB-Bax1 complex, as no unwinding or nuclease activity was observed when it was deleted (Rouillon and White 2010). In the past few years, evidence from studies of eukaryal TFIIH has accumulated that supports a role for XPB as a dsDNA translocase rather than a helicase. In this model, XPB binds dsDNA and catalyses opening of a DNA bubble downstream of the binding site in an ATP-dependent reaction (He et al.2016). Recent cryo-EM studies of the structural biology of transcription initiation appear to place this model beyond doubt (Schilbach et al.2017), at least for transcription and most likely for NER too. The work on archaeal XPB is largely consistent with a function as a dsDNA translocase rather than a helicase. The XPB-Bax1 complex could thus function as a stripped-down version of the eukaryal NER apparatus by binding at the site of helix-destabilising lesions, opening a bubble through XPB's ATP-dependent translocase activity and cleavage at the lesion by Bax1. Such a mechanism is still largely speculative however, requiring further study. Transcription coupled repair Transcription coupled repair (TCR) differs from global genome repair (GGR, described above) in being initiated by stalling of RNA polymerase on the transcribed strand of genes. A coupling factor (Mfd in bacteria; CS-B/RAD26 in eukarya) is then recruited to the stalled complex and in turn recruits the NER machinery to repair the damage. This alternative NER pathway typically has faster kinetics than GGR, meaning that DNA lesions in transcribed strands are repaired more quickly than those in non-transcribed ones. RNA polymerase from the archaeon T. kodakarensis has been shown to stall when encountering a variety of DNA lesions in template strands during transcription, suggesting that stalled RNA polymerase molecules are a common sensor for DNA damage in all domains of life (Gehring and Santangelo 2017). Accelerated TCR has been observed in the halophiles and shown to be dependent on UvrA in H. salinarum (Stantial et al.2016). This suggests a mechanism similar to that in bacteria, although there is no clear Mfd orthologue in archaea. On the other hand, two independent studies have demonstrated that TCR is not faster than GGR in S. solfataricus (Dorazi et al.2007; Romano et al.2007). A comparison of the rates of repair of transcribed and non-transcribed strands in S. solfataricus, E. coli and Saccharomyces cerevisiae suggests that the archaeon has a significantly faster rate of GGR, which may explain the apparent lack of accelerated TCR (Dorazi et al.2007). At any rate, the identification of coupling factors in archaea that link stalled RNA polymerase to the NER pathways is an important area for further study. DNA TRANSFER SYSTEMS Two independent studies of the transcriptional response to UV radiation in the Sulfolobales highlighted the upregulation of an operon comprised of five genes of unknown function (Fröls et al.2007; Götz et al.2007). UV treatment was also observed to result in significant and reversible cell aggregation in S. acidocaldarius. Subsequent analysis revealed that the operon encoded genes specific for a type IV pilus structure, leading the renaming of the operon as the ups operon (for UV-inducible pili operon of Sulfolobus) (Fröls et al.2008). These findings led to the hypothesis that the Ups system represents a machinery for the exchange of DNA following DNA damage (Fröls et al.2008). Pili were shown to mediate species-specific aggregates and to support large increases in the rate of HR, providing a survival advantage in following DNA damage (Ajon et al.2011). Downstream of the ups operon in S. acidocaldarius are four conserved genes predicted to encode a ParB-like nuclease, a glycosyl transferase, an EndoIII-like nuclease and a helicase. Deletion of these genes did not abrogate UV-induced aggregation, but did result in a significant decrease in survival following UV irradiation, suggesting that this operon likely plays a role in DNA repair that is related in some way to the UV-inducible DNA transfer pathway (van Wolferen, Ma and Albers 2015). A further twist to the story came with the identification of the Ced (crenarchaeal system for exchange of DNA) system for DNA import. The UV-inducible ced genes encode CedA, an integral membrane protein, and CedB, a membrane-bound hexameric DNA translocase related to HerA (van Wolferen et al.2016). CedA and CedB are thought to assemble to form a machine for the import of DNA following Ups-mediated cell aggregation, thus enhancing recombination and DNA repair. This fascinating system seems to be unique to the Crenarchaea—no other examples of a DNA import (rather than export) machinery is currently known in the prokaryotes (van Wolferen et al.2016). BASE EXCISION REPAIR AND ALTERNATIVE EXCISION REPAIR Damage to individual bases, caused for example by hydrolytic deamination, oxidation or methylation, is the most common and unavoidable type of DNA damage. Therefore, it is perhaps not surprising that the DNA repair pathway responsible for detection and correction of these lesions, BER, is ubiquitous and fundamentally conserved across all domains of life. The canonical BER pathway is initiated when a glycosylase specific for a particular damaged base detects the lesion, usually by base flipping, and cleaves the glycosidic bond, generating an abasic (AP) site. The AP site in DNA is detected by AP endonuclease, which cleaves the phosphodiester backbone on the 5΄ side of the lesion, allowing DNA polymerase to initiate repair synthesis. Depending on circumstances, BER is completed by flap displacement and subsequent removal by the Fen1 nuclease (long patch repair), or by removal of the abasic nucleotide by RP lyase (short patch repair)—with both pathways resulting in nicked DNA that can be ligated (reviewed in Grasso and Tell 2014). The alternative excision repair (AER) pathway is imitated by an endonuclease (rather than a glycosylase), which nicks the DNA backbone next to a DNA lesion (reviewed in Yasui 2013). Since rates of hydrolytic deamination increase with growth temperature, this type of damage is particularly problematic for thermophiles and hyperthermophiles. Deamination of uracil, guanine and adenine, which give rise to uracil, xanthine and hypoxanthine, respectively, is a particular problem as they have the potential to result in altered base pairing and hence mutation if not repaired quickly. Endonuclease V (EndoV) is a nuclease found in all domains of life that cuts at the 3΄ side of hypoxanthine residues in DNA, initiating the AER pathway. Many archaeal genomes, including the majority of the thermophiles, possess a gene encoding EndoV (Kiyonari et al.2014). Biochemical studies revealed that the EndoV enzyme from A. fulgidus and P. furiosus has the canonical specificity for inosine (Liu et al.2000; Kiyonari et al.2014). In contrast, EndoV from Ferroplasma acidarmanus has a broader specificity for deaminated bases (Kanugula et al.2005). Recently, a second nuclease, endonuclease Q (EndoQ), has been identified in P. furiosus which cleaves the DNA backbone on the 5΄ side of deaminated bases or abasic sites (Shiraishi et al.2015). In contrast to EndoV, the EndoQ enzyme has a narrow distribution in the archaea (Shiraishi et al.2015). EndoQ forms a physical and functional interaction with the sliding clamp PCNA (Shiraishi et al.2016), which may help direct the enzyme to the replication fork, increase the efficiency of the enzyme and allow coordinated repair with Fen1, DNA polymerase and DNA ligase, which are all PCNA-interacting enzymes (Fig. 5). PCNA is an important partner for many other DNA repair enzymes, including AP endonuclease (Kiyonari et al.2009), uracil DNA glycosylase (Kiyonari et al.2008), the XPF nuclease (Roberts, Bell and White 2003) and the more recently characterised Nre protein (Giroux and MacNeill 2016), which has a clear though as yet undefined role in DNA repair. In the future, we can expect that further BER enzymes, particularly nucleases, will be discovered in the distinct archaeal lineages. Orthologues of EndoQ outwith the Thermococcales, for example, seem very likely to exist but are as yet unidentified. Figure 5. View largeDownload slide EndoQ pathway for alternative excision repair of deaminated DNA. 1. EndoQ detects deaminated base, cleaving DNA backbone on 5΄ side. 2. DNA polymerase extends the 3΄ end of DNA, displacing a DNA flap including the lesion. 3. Fen1 removes the 5΄ flap, leaving nicked DNA that is ligated by DNA ligase (4). The process may be coordinated by PCNA, which interacts with each of the enzymes. Similar pathways may pertain for other glycosylases and DNA repair nucleases that interact with PCNA. It is not yet clear whether this ‘molecular toolbelt’ view of PCNA reflects reality, as protein partners will associate and dissociate in dynamic equilibrium. Figure 5. View largeDownload slide EndoQ pathway for alternative excision repair of deaminated DNA. 1. EndoQ detects deaminated base, cleaving DNA backbone on 5΄ side. 2. DNA polymerase extends the 3΄ end of DNA, displacing a DNA flap including the lesion. 3. Fen1 removes the 5΄ flap, leaving nicked DNA that is ligated by DNA ligase (4). The process may be coordinated by PCNA, which interacts with each of the enzymes. Similar pathways may pertain for other glycosylases and DNA repair nucleases that interact with PCNA. It is not yet clear whether this ‘molecular toolbelt’ view of PCNA reflects reality, as protein partners will associate and dissociate in dynamic equilibrium. DSB REPAIR Pathways of DSBR DSBs are arguably the most lethal form of DNA damage that cells can incur. DSBs have the potential to block essential processes such as transcription, DNA replication and cell division. Because both strands of the DNA duplex are broken, the inaccurate repair of DSBs can lead not just to mutations but also to genome rearrangements. The most accurate form of DSBR, which largely avoids such collateral damage, is HR. However, this is a complex and energetically demanding process and for this reason, simpler but less accurate pathways of DSBR operate alongside HR (Fig. 6). Figure 6. View largeDownload slide Pathways of DSB repair. Double-strand DNA breaks are repaired by non-homologous end joining (NHEJ), microhomology-mediated end joining (MMEJ) or homologous recombination (HR). Figure 6. View largeDownload slide Pathways of DSB repair. Double-strand DNA breaks are repaired by non-homologous end joining (NHEJ), microhomology-mediated end joining (MMEJ) or homologous recombination (HR). Non-homologous end joining (NHEJ) is initiated by binding of the Ku protein complex, which acts as a scaffold to recruit nucleases, polymerases and ligases that process and repair the DSB (Fig. 5). It is a rapid and versatile pathway of repair, which can accommodate DNA ends with a variety of lesions that would otherwise be refractory to ligation. Although it is error-prone, NHEJ is commonly used in eukaryotic cells, in particular higher eukaryotes that are quiescent in the G1 phase of the cell cycle, because it does not depend on the presence of a homologous DNA duplex. However, NHEJ in archaea is rare because it requires the Ku protein and this is found in only a small number of species. In fact, a complete NHEJ complex, comprising Ku, polymerase, phosphoesterase and ligase, has only been found in Methanocella paludicola (Bartlett, Brissett and Doherty 2013). Crystal structures of these archaeal enzymes have demonstrated a conservation with their bacterial NHEJ counterparts (Bartlett et al.2016). Microhomology-mediated end joining (MMEJ) is a primitive method of DSBR that does not require the Ku complex. Instead, DSBs are resected by exonucleases to expose short single-stranded tracts of homology that anneal with each other. Trimming of the resulting flaps is followed by DNA synthesis and ligation (Fig. 6); like NHEJ, this method of DSBR can result in deletions. MMEJ has been observed in Haloferax volcanii (Delmas et al.2009; Stachler et al.2017) and S. islandicus (Zhang and Whitaker 2018), but the enzymatic basis is currently unknown. HR is the only error-free method of DSBR, because it uses a second copy of DNA as a template (Fig. 6). As suggested by its name, HR requires the intact template to be homologous to the broken DNA duplex, but genetic studies in S. acidocaldarius have found that archaea might utilise shorter tracts of homology than bacteria or eukaryotes (Grogan and Stengel 2008; Rockwood, Mao and Grogan 2013). There are three steps to HR. (1) Pre-synapsis: The DSB is resected by exonucleases to generate 3΄ single-stranded DNA tails that are bound by the RecA-family recombinase, which in archaea is RadA. (2) Synapsis: The nucleoprotein filament formed by RadA engages in a homology search with an intact duplex, whereupon it catalyses strand exchange to form a displacement loop (D-loop); the 3΄ end in the D-loop is used to prime DNA synthesis. (3) Post-synapsis: At this point the invading strand may be displaced by a helicase, and the newly synthesised section of DNA will allow it to reanneal with the other end of the DSB. This method of HR results exclusively in non-crossovers. Alternatively, capture of the second DSB end by the D-loop will result in the formation of a double Holliday junction structure. This is resolved by structure-specific endonucleases to yield either crossover or non-crossover products, depending on the orientation of the cuts. HR is the best-studied pathway of DSBR in archaea (White 2011). In addition to its primary role in DNA repair (Fujikane et al.2010; Liang et al.2013; Zhang et al.2013), HR is used to promote genetic diversity following DNA transfer between Sulfolobus species (van Wolferen et al.2013, 2016; van Wolferen, Ma and Albers 2015) and between Haloferax species (Naor et al.2012, 2016). HR is also used to restart DNA replication at stalled forks, which arise at DNA damage or protein roadblocks. This ability to initiate DNA replication using the invading 3΄ end of a D-loop is harnessed in strains of H. volcanii and T. kodakarensis that are deleted for replication origins. In originless mutants, HR is essential because it is used constitutively to initiate all DNA replication (Hawkins et al.2013; Gehring et al.2017). HR pre-synapsis—Mre11-Rad50 and NurA-HerA DSBs must be processed by exonucleases to generate the 3΄ single-stranded DNA tails that form nucleoprotein filaments with RadA. In E. coli, this resection is carried out by RecBCD helicase/exonuclease. In eukaryotes, the Mre11 and Rad50 proteins form a complex that initiates resection by limited 3΄ to 5΄ degradation, followed by extensive resection by 5΄ to 3΄ exonucleases. Mre11 and Rad50 are conserved in archaea and structural studies have shown that they form a complex with DNA binding, unwinding and resection activities (Deshpande et al.2014; Sung et al.2014; Liu et al.2016). In S. acidocaldarius, the Mre11-Rad50 complex undergoes post-translational methylation in response to γ-irradiation (Kish et al.2016), and in H. volcanii the Mre11-Rad50 complex acts in both the repair of DSBs and the compaction of the nucleoid after DNA damage (Delmas et al.2009; Delmas, Duggin and Allers 2013). In many archaeal species, the genes for Mre11 and Rad50 are found in an operon with those for the hexameric HerA helicase and the NurA nuclease, and the NurA-HerA complex has recently been the subject of much exciting research. Structural studies have revealed that NurA forms a toroidal dimer with a narrow central channel that can accommodate the two strands of an unwound duplex (Blackwood et al.2012; Byrne et al.2014). In complex with a HerA hexamer, the NurA dimer generates a continuous channel, indicating that HerA-driven translocation propels the DNA duplex through the NurA nuclease ring, where it is unwound and degraded (Fig. 7) (Rzechorzek et al.2014; Ahdash et al.2017). The nuclease activity of NurA is modulated by HerA, and was found to be essential for cell viability in S. islandicus (De Falco et al.2015; Huang et al.2015). Bacterial homologues of NurA-HerA have been identified in Deinococcus radiodurans, and play a role in HR (Cheng et al.2015a,b). Figure 7. View largeDownload slide NurA-HerA complex. Model for how HerA and NurA might process DNA ends. Double-stranded DNA is channelled through HerA helicase and unwound by the ploughshare motif in NurA. Both DNA strands are degraded by the NurA nuclease. From Rzechorzek et al. (2014). Figure 7. View largeDownload slide NurA-HerA complex. Model for how HerA and NurA might process DNA ends. Double-stranded DNA is channelled through HerA helicase and unwound by the ploughshare motif in NurA. Both DNA strands are degraded by the NurA nuclease. From Rzechorzek et al. (2014). HR synapsis—SSB, RadA and its paralogues The 3΄ single-stranded tail is bound by the RecA-family recombinase RadA (Morrical 2015). RadA polymerisation is driven by the insertion of an invariant phenylalanine in the N-terminal domain into a binding pocket of an adjacent monomer (Fig. 8). The DNA in this nucleoprotein filament is stretched ∼1.5× in length, which facilitates the search for homologous sequences and the strand exchange process (Fig. 6). To form the nucleoprotein filament, RadA must first displace single-stranded DNA-binding protein (SSB), a ubiquitous protein with an oligonucleotide-binding (OB) fold, a twisted β-barrel with a binding site that accommodates four nucleotides of ssDNA (Lin et al.2008). The SSBs found in Euryarchaea are similar to the heterotrimeric eukaryotic RPA, which forms a heterotrimer, whereas the SSBs in Crenarchaea are more akin to the homotetrameric bacterial SSB; both the euryarchaeal RPA and crenarchaeal SSBs show a greater variety of architectures than their eukaryotic or bacterial counterparts. The S. solfataricus SSB has been shown to interact with RadA and inhibit its single-stranded DNA-dependent ATPase activity (Rolfsmeier and Haseltine 2010). In order to stimulate strand exchange and overcome inhibition by SSB, the Rad54 protein of S. solfataricus can interact with RadA and remodel the topology of the homologous duplex DNA (Haseltine and Kowalczykowski 2009). Figure 8. View largeDownload slide RadA recombinase. Rad:DNA nucleoprotein filament formation by insertion of phenylalanine into binding pocket of an adjacent RadA monomer. From Wardell et al. (2017). Figure 8. View largeDownload slide RadA recombinase. Rad:DNA nucleoprotein filament formation by insertion of phenylalanine into binding pocket of an adjacent RadA monomer. From Wardell et al. (2017). The role of displacing SSB from single-stranded DNA and loading RadA more commonly falls to RadA paralogues (Lin et al.2006) and in this capacity, they are known as recombination mediators. RadB is found only in Euryarchaea, it interacts with RadA (Patoli et al.2017) and functions as a recombination mediator in H. volcanii, where it has been proposed to induce a conformational change in RadA and thereby promote its polymerisation on DNA (Wardell et al.2017). Similarly in S. solfataricus, the RadA paralogue SsoRal1 enhances RadA binding of single-stranded DNA and stabilises the nucleoprotein filament (Graham, Rolfsmeier and Haseltine 2013). By contrast, the S. solfataricus paralogue Sso2452 and the Sulfolobus tokodaii paralogue stRadC2 have been found to inhibit strand exchange and D-loop formation by RadA (McRobbie et al.2009; Wang et al.2012). An in vivo study of two RadA paralogues in S. islandicus, RadC1 and RadC2, has shown that both are involved in DNA repair but the effect on HR has yet to be determined (Liang et al.2013). HR post-synapsis—Hel308, Hef and Hjc Once a D-loop is formed it can be used to prime DNA synthesis; the nascent 3΄ end may then be unwound to reanneal with the other side of the DSB. This is known as synthesis-dependent strand annealing and yields only non-crossover products. In archaea, the enzyme responsible for unwinding the invading strand is likely to be Hel308, a Ski2-family helicase found in archaea and metazoans but not in bacteria or yeast (Woodman and Bolt 2009). Hel308 is essential for cell viability in S. tokodaii (Hong et al.2012; Song, Ni and Shen 2016) but not in H. volcanii (T. Allers, unpublished). It interacts with RPA (Woodman, Brammer and Bolt 2011), and structural studies have shown that when Hel308 is bound to a 3΄ single-strand tailed partial duplex (Fig. 9), the helicase domains encircle single-stranded DNA in a ‘ratchet’ for directional translocation (Richards et al. 2008). It has recently been found that DNA binding and unwinding by Hel308 requires a distinctive winged helix domain (Northall et al.2017). Taken together, these studies suggest that Hel308 controls HR at the D-loop step and assists in the restart of stalled DNA replication forks (Northall et al.2016). Figure 9. View largeDownload slide Hel308 helicase. DNA duplex is unwound into single strands by Hel308. From Richards et al. (2008b). Figure 9. View largeDownload slide Hel308 helicase. DNA duplex is unwound into single strands by Hel308. From Richards et al. (2008b). Instead of being unwound, the D-loop may capture the second end of the DSB and thereby form a four-way Holliday junction structure. An enzyme that most likely mediates this transition in Euryarchaea is Hef (Lestini, Delpech and Myllykallio 2015). A member of the XPF/MUS81 family of structure-specific endonucleases, Hef, comprises two distinct domains: an N-terminal domain of the DEAH helicase family and a C-terminal domain of the XPF endonuclease family, it acts on nicked, flapped and forked DNA (Komori et al.2004). Hef forms specific localisation foci in vivo in response to replication fork arrest (Lestini et al.2013), and has been shown to interact with several DNA repair and replication proteins, including RecJ-like exonucleases and the PCNA sliding clamp of the DNA replication apparatus (Ishino et al.2014; Rohleder et al.2016; Nagata et al.2017). In H. volcanii, Hef is essential for cell viability when the Holliday junction resolvase Hjc is absent, and both the helicase and nuclease activities of Hef are indispensable (Lestini, Duan and Allers 2010). It has been proposed that Hef and Hjc provide alternative means to restart stalled DNA replication forks by processing Holliday junctions. In contrast to Hef, Hjc has only nuclease activity and is specific for four-way DNA structures (Komori et al.1999). Enzymes of this class are known as Holliday junction resolvases and are capable of generating crossover products (Fig. 6). A second Holliday junction resolvase, Hje, is present in Sulfolobales and a genetic study of Hjc and Hje in S. islandicus found that while deletion of either hje or hjc had no effect on cell viability, deletion of both hje and hjc is lethal (Huang et al.2015). This parallels the redundancy between Hjc and Hef in H. volcanii (Lestini, Duan and Allers 2010). Hjc has been observed to interact with many DNA repair proteins such as the RadA paralogue RadC2 (Wang et al.2012), the Hel308 helicase (Hong et al.2012) and a novel ATPase from S. islandicus termed SisPINA (Zhai et al.2017); the latter forms hexameric rings, similar to the bacterial Holliday junction migration helicase RuvB. Another novel protein that has been reported to bind to Holliday junctions is the phMutS5 MMR enzyme from P. horikoshii (Ohshita et al.2017), but unlike eukaryotic MutS homologues that act in HR, phMutS5 showed no nuclease activity on branched DNA. Applications of DSBR HR is not only an error-free method of DSBR but also a cornerstone of archaeal genetics (Leigh et al.2011; Farkas, Picking and Santangelo 2013). The ability to target a specific gene for deletion or mutation, using plasmid constructs with flanking regions of homology, relies on HR (Fig. 10). Refinements of these methods have enabled the high-throughput generation and screening of targeted mutants in P. furiosus (Farkas et al.2012), S. islandicus (Zhang et al.2013) and H. volcanii (Kiljunen et al.2014); the latter is notable for using a transposon insertion library to carry out saturation mutagenesis, which facilitates the identification of non-essential genes in any specific pathway. Figure 10. View largeDownload slide Typical strategy for gene deletion in archaea using HR. A plasmid with flanking homology is used to delete and replace a target gene with a selectable marker (Δ). A second marker for uracil biosynthesis (URA) is used for selection and counterselection (using 5-FOA) of cells that have undergone HR as indicated. Figure 10. View largeDownload slide Typical strategy for gene deletion in archaea using HR. A plasmid with flanking homology is used to delete and replace a target gene with a selectable marker (Δ). A second marker for uracil biosynthesis (URA) is used for selection and counterselection (using 5-FOA) of cells that have undergone HR as indicated. Other pathways of DSBR have been harnessed in genetic manipulation and genome engineering. MMEJ has recently been used for a high-throughput method for targeted gene inactivation in S. islandicus, in one case the minimal size of microhomology for marker replacement was as few as 10 bp (Zhang and Whitaker 2018). In Methanosarcina acetivorans, a system of CRISPR-Cas9-mediated genome engineering has been developed and it was found that co-expression the NHEJ machinery from M. paludicola allowed efficient genome editing without the need for a repair template (Nayak and Metcalf 2017). The enzymes involved in DSBR have also found applications in vitro. For example, the thermostable RadA recombinase from P. woesei enhances the specificity of simplex and multiplex PCR assays (Stefanska et al.2016). Similarly, the Hel308 helicase from T. gammatolerans has found a new lease of life as a motor protein for nanopore sequencing. Owing to its ability to unwind duplex DNA and ratchet the single stranded DNA through the nanopore in a stepwise manner, Hel308 significantly improves the accuracy of single-molecule sequencing (Craig et al.2015, 2017; Derrington et al.2015). CONCLUDING REMARKS Research into DNA repair in the archaea has flourished since the turn of the millennium, driven largely by the availability of genome sequences. However, the emerging picture fits with neither of the preconceptions that were held 20 years ago. Archaea are neither ‘odd’ bacteria, a view held by detractors of the third domain of life, nor are they ‘mini-eukaryotes’ as proposed by those who believed they would serve as simplified models for human cells. Instead, archaea have proved to be every bit as unique and diverse as bacteria and eukaryotes, and the archaeal systems for DNA repair reflect this distinctive status. Genomic surveys have revealed a patchwork of bacterial and eukaryotic repair enzymes, alongside proteins that are unique to archaea, but laboratory studies have shown that these enzymes do not necessarily behave in the same way as their bacterial or eukaryotic counterparts. Nevertheless, one aspect of the field has not changed in 20 years—archaea and their systems for DNA repair continue to serve as a window into our evolutionary past. SUPPLEMENTARY DATA Supplementary data are available at FEMSRE online. FUNDING MFW is the recipient of a Royal Society Wolfson Research Merit Award. Work in TA's laboratory is funded by the Biotechnology and Biological Sciences Research Council (BBSRC). Conflicts of interest. None declared. REFERENCES Adam PS , Borrel G , Brochier-Armanet C et al. The growing tree of Archaea: new perspectives on their diversity, evolution and ecology . ISME J 2017 ; 11 : 2407 – 25 . Ahdash Z , Lau AM , Byrne RT et al. Mechanistic insight into the assembly of the HerA-NurA helicase-nuclease DNA end resection complex . Nucleic Acids Res 2017 ; 45 : 12025 – 38 . Ajon M , Frols S , van Wolferen M et al. UV-inducible DNA exchange in hyperthermophilic archaea mediated by type IV pili . Mol Microbiol 2011 ; 82 : 807 – 17 . 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FEMS Microbiology Reviews – Oxford University Press
Published: May 5, 2018
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