Archaeal DNA replication and repair: new genetic, biophysical and molecular tools for discovering and characterizing enzymes, pathways and mechanisms

Archaeal DNA replication and repair: new genetic, biophysical and molecular tools for discovering... Abstract DNA replication and repair are essential biological processes needed for the survival of all organisms. Although these processes are fundamentally conserved in the three domains, archaea, bacteria and eukarya, the proteins and complexes involved differ. The genetic and biophysical tools developed for archaea in the last several years have accelerated the study of DNA replication and repair in this domain. In this review, the current knowledge of DNA replication and repair processes in archaea will be summarized, with emphasis on the contribution of genetics and other recently developed biophysical and molecular tools, including capillary gel electrophoresis, next-generation sequencing and single-molecule approaches. How these new tools will continue to drive archaeal DNA replication and repair research will also be discussed. archaea, capillary gel electrophoresis, DNA repair, DNA replication, genetics, next-generation sequencing INTRODUCTION DNA replication plays an essential role in all life forms. The process is required for the propagation and evolution of genetic information. The mechanism of DNA replication guarantees the duplication and transfer of genetic information during cell division. Although the process needs to be precise, the fidelity of the replication process is not absolute and other cellular and environmental insults result in DNA damage and alterations in the DNA sequence. Consequently, another process, DNA repair, is also required to ensure the integrity of chromosomal DNA. In all organisms, chromosomal DNA replication is a complex event that involves many factors to ensure the accurate and timely duplication of genetic information. The mechanism of DNA replication is fundamentally conserved in all life forms (reviewed in Kelman and Kelman 2014; Yao and O’Donnell 2016). The process is thought to start at a specific sequence called an origin of replication, at which origin-binding proteins (OBP) bind and locally unwind duplex DNA. Additional proteins interact with the OBP-DNA complex and are responsible for the assembly of the DNA helicase around the DNA. Once assembled, the helicase unwinds the duplex and forms the initial replication bubble. The exposed single-stranded (ss) DNA at the replication bubble is coated with ssDNA-binding protein (SSB). DNA primase, DNA polymerases (Pol) and the rest of the replication machinery are recruited to the SSB-ssDNA complex to initiate bidirectional DNA synthesis. Due to the antiparallel nature of duplex DNA, and the unidirectionality of DNA polymerases, one strand of the chromosome is synthesized continuously (leading strand) while the other is copied discontinuously (lagging strand) as a series of Okazaki fragments (Fig. 1). Figure 1. View largeDownload slide New tools to study archaeal DNA replication and repair. A model for the archaeal replication fork. The leading and lagging strands are shown as well as the proteins involved in Okazaki fragment maturation, and ribonucleotide excision repair. The tools described in the review used to study these processes are shown: single-molecule analysis (pink), next-generation sequencing (orange), genetics (green) and capillary electrophoresis (blue). Figure 1. View largeDownload slide New tools to study archaeal DNA replication and repair. A model for the archaeal replication fork. The leading and lagging strands are shown as well as the proteins involved in Okazaki fragment maturation, and ribonucleotide excision repair. The tools described in the review used to study these processes are shown: single-molecule analysis (pink), next-generation sequencing (orange), genetics (green) and capillary electrophoresis (blue). As cells grow and divide, they are continuously exposed to DNA-damaging agents that arise endogenously by cellular metabolism, or from external environmental factors. The exposure to DNA-damaging agents leads to the formation of a variety of DNA lesions that are mutagenic and/or cytotoxic to the cell, and can halt DNA replication and cause cellular apoptosis. Due to the toxic nature of DNA lesions, organisms have evolved several DNA repair pathways including direct damage reversal, base excision repair (BER), nucleotide excision repair (NER), non-homologous end joining (NHEJ) and homologous recombination (HR). The extreme environments in which many archaeal species thrive, including high temperature, high salinity or low pH, facilitate DNA damage and lesion formation, and thus archaea likely require robust DNA repair mechanisms for genome maintenance. For many years, the study of archaeal DNA replication and repair was hindered by a lack of genetic and other biophysical tools. Therefore, most of the information regarding these processes was derived from biochemical, structural and limited cellular studies. However, in the last decade new tools have been developed in several archaeal species including genetic tools, such as the knockin and knockout of genes, and biophysical and molecular tools, such as capillary gel electrophoresis (CE), single-molecule analysis of individual proteins and processes, and next-generation sequencing (NGS) (Fig. 1). These assays more accurately reflect biological processes in the cell and accelerate understanding of enzymes and pathways. In this review, the impact of recently developed tools on the study of the replication and repair processes in archaea will be summarized. We also speculate how these new tools will drive future archaeal DNA replication and repair research. Readers are referred to several past reviews for previously summarized results (Kelman and White 2005; Barry and Bell 2006; White 2011; Grasso and Tell 2014; Kelman and Kelman 2014). GENETIC TOOLS Genetic tools allow for the in vivo manipulation of genes within an organism, including the introduction, modification or deletion of a particular gene of interest. Importantly, genetic tools allow one to understand the importance, or lack thereof, of a particular gene. Genetic tool systems have long been established in eukaryotic and bacterial organisms, including the routinely used yeast Saccharomyces cerevisiae and the Escherichia coli bacterium, as well as in a variety of higher organisms (Doudna and Charpentier 2014; Duina et al.2014; Blount 2015). In the last several years, robust genetic tools have also been developed for several archaeal species from Euryarchaeota and Crenarchaeota phyla (Atomi, Imanaka and Fukui 2012). Species include Methanosarcina acetivorans (Kohler and Metcalf 2012), Haloferax volcanii (Leigh et al.2011), Pyrococcus furiosus (Lipscomb et al.2011), Thermococcus kodakarensis (Hileman and Santangelo 2012; Farkas et al.2013) and Sulfolobus solfataricus (Wagner et al.2012). As the replication machinery of crenarchaea is different from all other archaeal branches, we will concentrate on the tools developed in euryarchaea and their impact on the fields of DNA replication and repair. –) Gene knockout When genetic tools became available for archaea, one of the first questions asked was: Which replication and repair proteins are essential for viability? To answer this question, two main genetic approaches were used. One approach is the direct deletion of a specific gene or region of the chromosome (Fig. 2), while the other uses random integration of transposons to disrupt gene function to identify essential genes (Sarmiento, Mrazek and Whitman 2013). Most results were as expected; proteins that were shown to be essential in bacteria and/or eukarya were also essential in archaea, and those that were not essential in the other domains were not essential in archaea. For example, the polymerase accessory proteins, replication factor C (RFC) and proliferating cell nuclear antigen (PCNA) are essential in both eukarya and archaea (Haracska et al.2001; Kuba et al.2012; Pan et al.2013; Sarmiento, Mrazek and Whitman 2013), while Flap endonuclease 1 (Fen1), an enzyme that removes 5΄ overhangs generated during Okazaki fragment maturation and DNA repair, is dispensable for viability in both (Reagan et al.1995; Burkhart et al.2017). However, there were also unexpected results; several of those will be described here as examples. Figure 2. View largeDownload slide A genetic knockout system for T. kodakarensis. A target gene (yellow) is deleted from a recipient strain via two homologous recombination events using donor DNA containing selectable marker (SM), and CSM (counter selectable marker), for use as positive and negative selection. This approach results in the construction of markerless deletion strains (see Hileman and Santangelo 2012 for details). Figure 2. View largeDownload slide A genetic knockout system for T. kodakarensis. A target gene (yellow) is deleted from a recipient strain via two homologous recombination events using donor DNA containing selectable marker (SM), and CSM (counter selectable marker), for use as positive and negative selection. This approach results in the construction of markerless deletion strains (see Hileman and Santangelo 2012 for details). +) Genetic knockouts of a conserved archaeal origin of replication and origin initiator proteins One of the most exciting and unexpected observations made using knockout experiments in archaea is the observation that several archaeal species do not require a traditional conserved origin of replication. It is well established that organisms require an origin of replication to initiate chromosomal replication and to provide a point of regulation for the replication process. Importantly, while bacterial organisms contain a single conserved origin of replication, higher eukaryotes contain many less-defined origins (Kornberg and Baker 1992; Boulikas 1996; DePamphilis 1996; O’Donnell, Langston and Stillman 2013). It has been proposed that archaea use conserved origins, similar to bacteria, due to bioinformatic studies that identified a conserved archaeal sequence containing known origin of replication motifs (Norais et al.2007). Interestingly, gene knockout studies have shown that in some archaeal species these origins are dispensable for cell viability and can be readily deleted in laboratory strains [(Hawkins et al.2013; Gehring et al.2017) summarized in (Kelman and Kelman 2018)]. A study demonstrating that the origin can be deleted led to even more surprising results. Under normal laboratory growth conditions, T. kodakarensis does not utilize the predicted conserved origin of replication to initiate DNA synthesis (Gehring et al.2017). Furthermore, a secondary origin of replication was not identified, suggesting random initiation of DNA replication across the chromosome. This interesting, dogma-shattering observation questions the necessity of the origin of replication in archaea, and will require further investigation. It is important to reevaluate under which conditions the conserved origin of replication is used, why it is retained in the genome and how DNA replication is initiated on a chromosome in strains lacking a conserved origin (Hawkins et al.2013). Archaea contain homologs of several subunits of the eukaryotic origin recognition complex (Orc), in particular Orc1 and the replication initiation protein Cdc6 (referred to as Orc1/Cdc6 in archaea). These archaeal homologs were shown to bind to the archaeal origin of replication and were suggested to initiate the replication process by assembling the helicase (Matsunaga et al.2001; Costa, Hood and Berger 2013). Similar to the observation that the origin of replication is dispensable in some archaeal species, it was shown that the Orc1/Cdc6 gene can also be deleted in those species (Gehring et al.2017). In bacteria and eukarya, the OBPs and initiation proteins participate in regulating the initiation process. Further study is needed to determine which proteins and processes regulate the initiation of replication in the absence of Orc1/Cdc6. +) Genetic knockouts of archaeal DNA polymerases Most archaeal linages, except for crenarchaea, contain two different polymerases, a family B DNA polymerase, PolB, and a family D DNA polymerase, PolD (Cann et al.1998; Ishino et al.1998). The three replicative polymerases in eukarya, Polα, Polε and Polδ, all belong to family B, and crenarchaea contain only PolB homologs. Therefore, it was proposed that members of the PolB family are responsible for chromosomal replication in all of archaea (Grabowski and Kelman 2003; Johansson and Dixon 2013). However, in many archaeal species, the genes encoding the two archaeal-specific subunits of PolD are located in close proximity to the origin of replication and are often in an operon with other replication enzymes (for example, see Kelman 2000). The proximity of proteins involved in chromosomal replication to the origin of replication is common in bacteria and archaea (Kornberg and Baker 1992). In addition, the small exonuclease subunit of PolD shares amino acid sequence similarity with several of the small non-catalytic subunits of eukaryotic Polα, Polδ and Polε (Aravind and Koonin 1998). It was therefore proposed that PolD might also function at the archaeal replication fork, or that both PolB and PolD may be present at the fork. This would be similar to the situation in eukarya in which Polε replicates the leading strand, while Polδ copies the lagging strand (reviewed in Walsh and Eckert 2014). Early genetic studies supported the idea that both PolB and PolD may be present at the fork, as studies with Halobacterium indicated that both PolB and PolD may be essential for cell viability (Berquist, DasSarma and DasSarma 2007). However, studies in other archaeal species showed that PolB was dispensable for cell viability, while PolD was essential (Čuboňová et al.2013; Sarmiento, Mrazek and Whitman 2013). This result was unexpected and changes our understanding of chromosomal replication in at least some archaeal species. These data suggest that PolD (in the absence of PolB) could carry out both leading and lagging strand synthesis. It is possible, however, that when both polymerases are present in the cell, PolB, with its strand displacement activity (Greenough, Kelman and Gardner 2015) replicates the lagging strand, while PolD replicates the leading strand. +) Genetic knockouts help define the roles of archaeal Cdc45, GINS and MCM in DNA replication In eukarya, the heterohexameric minichromosome maintenance (MCM) helicase, the major replicative helicase, is not active on its own, but is activated by association with two accessory factors, the tetrameric GINS complex and the Cdc45 protein, forming the CMG (Cdc45, MCM, GINS) complex (reviewed in Onesti and MacNeill 2013; Li and O’Donnell 2018). In contrast to eukarya, the archaeal MCM helicase is active on its own in vitro without the requirement for additional factors (Kelman, Lee and Hurwitz 1999; Chong et al.2000 summarized in Costa and Onesti 2009; Sakakibara, Kelman and Kelman 2009). Homologs of the eukayotic GINS complex and Cdc45 have been identified in the archaeal genomes (Makarova et al.2005; Marinsek et al.2006; Makarova, Koonin and Kelman 2012; Oyama et al.2016 reviewed in Bell 2011). While the eukaryotic GINS complex is a ring-shaped heterotetramer of four related but different polypeptides, most archaeal GINS are tetramers compromised of two different proteins (MacNeill 2010). While the eukaryotic Cdc45 is essential for cell viability (Onesti and MacNeill 2013), in some archaeal species the Cdc45 homolog can be readily deleted with no effect on cell growth under normal growth conditions (Burkhart et al.2017 and Stuart MacNeill, personal communication). In contrast, the MCM and GINS proteins are essential in archaea (Kelman and Kelman 2014 and references therein). Nevertheless, it is possible that archaeal MCM, Cdc45 and GINS form a replicative helicase complex in vivo, as they are shown to form a stable complex in vivo (Marinsek et al.2006; Nagata et al.2017). +) Discovery of the archaeal primase using genetic knockouts In most archaeal genomes, a homolog of the bacterial primase, DnaG, has been identified. It was originally thought that this protein was the archaeal primase. However, studies have identified a two-subunit complex in archaea that is similar to the two-subunit eukaryotic primase (Bocquier et al.2001). Genetic studies showed that this two-subunit eukaryotic-like primase is the archaeal primase, not DnaG. While the gene encoding DnaG can be readily deleted, the genes encoding the eukaryotic-like primase cannot (Le Breton et al.2007). The archaeal primase is a two-subunit complex with similarities to the two-subunit eukayotic primase; a small subunit, PriS, contains the catalytic activity, and a large subunit, PriL, regulates primase activity (Bocquier et al.2001; Chemnitz Galal et al.2012 reviewed in Lao-Sirieix, Pellegrini and Bell 2005). Both subunits were shown to be essential for cell viability (Sarmiento, Mrazek and Whitman 2013). +) Genetic knockouts of archaeal DNA repair genes The use of genetic knockout techniques has been used to examine which archaeal DNA repair genes are essential, those that are non-essential and those in which knockout leads to phenotypic changes. Early experiments in H. volcanii lead to successful knockout of radA, which catalyzes strand exchange during HR (Woods and Dyall-Smith 1997). However, the ΔradA strain had severe HR defects and increased sensitivity to DNA-damaging agents. In T. kodakarensis, attempts to knock out several genes encoding HR proteins, including radA, rad50 and mre11, were unsuccessful, suggesting HR may be essential for T. kodakarensis survival, and not H. volcanii (Fujikane et al.2010). On the other hand, in T. kodakarensis, the genes encoding for xeroderma pigmentosum type B and D (XPB and XPD) helicase proteins can be readily deleted, and the knockout strains showed little or no sensitivity when challenged with UV radiation (Fujikane et al.2010). It is well established that mutations in XPB and XPD proteins in mammalians leads to xeroderma pigmentosum, a genetic disorder which renders an individual unable to repair DNA damage caused by UV-light exposure (reviewed in Lehmann, McGibbon and Stefanini 2011). The role XPB and XPD play in archaeal DNA repair is still unclear. Several other DNA repair genes have successfully been knocked out in T. kodakarensis, including Hjm, a RecQ like helicase, Hjc, a structure specific endonuclease, Hef, a helicase/nuclease whose role is proposed to help stalled replication forks, Fen1, a 5’ flap endonuclease, and RNaseH2, an endonuclease that cleaves at rNMP:dNMP junctions (Fujikane et al.2010; Burkhart et al.2017). +) Uncovering the role of multiple gene copies using genetic knockouts There are examples of genes that are present in a single copy in certain archaeal species, but present in multiple copies in other species. Gene knockouts were used to determine which gene copies are essential and which, if any, are dispensable. A few examples are described below. The vast majority of the known archaeal genomes encode for a single homolog of the MCM helicase, forming homohexamers (Sakakibara, Kelman and Kelman 2009). The genomes of several archaeal species, however, contain multiple MCM homologs. The presence of multiple MCM proteins in a single archaeon led to the proposal that in those organisms the MCM complex is similar to the eukaryotic helicase, where multiple subunits form a heterohexamer complex at the replication fork (Sakakibara, Kelman and Kelman 2009 and references therein). Using gene knockout experiments, it was found that in archaeal species that contain multiple MCM homologs, only one MCM homolog was essential for cell viability (for example, Ishino et al.2011; Pan et al.2011), suggesting they are similar to archaea containing a single MCM homolog with a homohexameric MCM helicase active at the replication fork. Another example is the polymerase processivity factor, PCNA. To date, all archaeal species encode for a single PCNA homolog forming a homotrimeric ring. There are two exceptions. One is organisms belonging to the crenarchaea in which three PCNA homologs are present, forming a heterotrimeric PCNA (Daimon et al.2002; Dionne et al.2003), and the other exception is the genome of T. kodakarensis that encodes for two PCNA homologs (Pan, Kelman and Kelman 2011). It is not clear why two PCNA proteins are present in the T. kodakarensis genome, or whether they have redundant functions. Although both PCNA proteins form similar homotrimeric rings (Ladner et al.2011) and can stimulate the activity of several enzymes (Ladner et al.2011; Kuba et al.2012; Li et al.2014), only one is essential for viability (Kuba et al.2012; Pan et al.2013). These results show that T. kodakarensis is similar to all other archaea species that contain only one PCNA protein. Non-essential gene copies, as in the case of MCM and PCNA in T. kodakarensis, are typically the result of gene duplication by viral integration (Fukui et al.2005). Importantly, all virally integrated regions can be removed from T. kodakarensis, without substantial effects on cell growth (Tagashira et al.2013). –) In vivo protein tagging The use of in vivo protein tagging allows for the incorporation of an affinity or detection tag at the genomic level to capture or visualize the resulting protein. The creation of genetic tools in archaeal systems has enabled archaeal in vivo protein tagging, which has been utilized to discover new archaeal DNA replication factors (Fig. 3). Figure 3. View largeDownload slide In vivo protein tagging. (A) Strains are constructed that encode for His-tagged replication and repair proteins. Stable in vivo complexes formed with the His-tagged protein are purified from clarified cell lysates. For example, His-tagged PCNA (black) forms a stable complex with Fen1 (purple), PolB (orange) and DNA ligase (pink). Co-purified proteins are identified by mass spectrometry. (B) Thermococcus kodakarensis PCNA is a central DNA replication enzyme and 12 proteins were co-purified with a tagged version of PCNA (Li et al.2010). (C) Thermococcales inhibitor of PCNA (TIP) was co-purified with PCNA and the proteins form a complex in vitro (PDB ID: 5DA7) (Altieri et al.2016). Orange, TIP; blue, PCNA. Figure 3. View largeDownload slide In vivo protein tagging. (A) Strains are constructed that encode for His-tagged replication and repair proteins. Stable in vivo complexes formed with the His-tagged protein are purified from clarified cell lysates. For example, His-tagged PCNA (black) forms a stable complex with Fen1 (purple), PolB (orange) and DNA ligase (pink). Co-purified proteins are identified by mass spectrometry. (B) Thermococcus kodakarensis PCNA is a central DNA replication enzyme and 12 proteins were co-purified with a tagged version of PCNA (Li et al.2010). (C) Thermococcales inhibitor of PCNA (TIP) was co-purified with PCNA and the proteins form a complex in vitro (PDB ID: 5DA7) (Altieri et al.2016). Orange, TIP; blue, PCNA. It is well established that proteins involved in the same pathway often form a stable complex. This is particularly true for the replisome complex, which is responsible for the duplication of chromosomal DNA (Yao and O’Donnell 2016). Although many archaeal replication and repair proteins and complexes have been identified based on their similarity to a bacterial or eukaryal counterpart, it is likely that some factors required for archaeal replication and regulation have yet to be identified (Kelman and Kelman 2014). Several known replication proteins were in vivo tagged with a 6 His-tag using a knockin approach (Li et al.2010) (Fig. 3). The insertion of the His-tag did not change the genomic context, including regulatory sequences, or position in an operon. Following purification on a Ni2+ column, the proteins that co-purified with the tagged protein were identified using mass spectrometry. Some of these interactions support previously reported biochemical and genetic studies. For example, it was found that the archaeal DnaG homolog does not interact with any known replication factors (Li et al.2010) but rather with exosome proteins, as was previously reported (Evguenieva-Hackenberg et al.2003), thus providing further evidence that DnaG is not the archaeal primase. Another example is the observation that PolB does not interact with many replisome proteins but PolD does. This study also identified a number of potentially new replication factors. Two examples are described below. +) Using in vivo protein tagging to discover the archaeal Cdc45 homolog As described above, the CMG complex in eukarya forms the replicative helicase required for the duplication of chromosomal DNA (Onesti and MacNeill 2013; Li and O’Donnell 2018). For many years, the components of the MCM and the GINS complexes were identified in archaea by similarity to eukaryotic homologus. However, no clear homolog of the Cdc45 protein could be identified by amino acid sequence comparison. The first suggestion that archaea might contain a Cdc45 homolog came from in vivo tagging experiments (Li et al.2010). A protein with homology to the bacterial RecJ nuclease co-eluted with the GINS complex. Furthermore, in silico analysis and structural prediction suggested that the protein, referred to as GAN (GINS-associated nuclease), shares similarity with RecJ and the eukaryotic Cdc45 protein (Makarova, Koonin and Kelman 2012; Oyama et al.2016). It was subsequently shown that the archaeal and eukaryal proteins have similar three-dimensional structures (Oyama et al.2016). The availability of genetic tools accelerated the discovery of GAN, underscoring the power and importance of in vivo tagging. +) Using in vivo protein tagging to uncover an archaeal DNA replication regulator protein PCNA plays an essential role in both DNA replication and repair (Moldovan, Pfander and Jentsch 2007). In eukarya, a number of mechanisms regulate the activity of PCNA during replication and repair, including post-translational modification and association with regulatory proteins (Vivona and Kelman 2003; López de Saro 2009; Dieckman, Freudenthal and Washington 2012). Until the development and utilization of archaeal in vivo tagging, no regulator of the archaeal PCNA had been reported. However, when the PCNA protein was tagged, a small protein co-eluted with it on Ni2+ column. The protein, referred to as TIP (Thermococcales inhibitor of PCNA), forms a complex with PCNA in vivo (Li et al.2010) and in vitro (Li et al.2014; Altieri et al.2016), and binding of TIP to PCNA inhibits PCNA activities (Li et al.2014) (Fig. 3). TIP is currently the only known protein that may regulate DNA replication, and possibly also DNA repair, in archaea. The emergence of archaeal protein tagging facilitated the discovery of this small protein. MOLECULAR AND BIOPHYSICAL TOOLS New molecular and biophysical tools are needed to discover new archaeal DNA replication and repair enzymes, to understand their function, and to explore cellular pathways and mechanisms. Sensitive assays will provide a clear picture of enzyme activities and interactions. Because enzymes do not function in isolation, but rather as complexes, assays that monitor coupled reactions will also be critical for understanding pathways occurring in the cell. To address these needs, various approaches to study single enzymes, protein complexes and in vivo processes using CE, NGS and single-molecule analysis have been developed and will be described here. –) Capillary gel electrophoresis CE is a sensitive high-throughput, high-resolution system for nucleic acid analysis, and serves as an alternative to gel-based electrophoresis analysis (Greenough et al.2016). In CE, fluorescently labeled nucleic acids are separated by size and charge, and detected by laser excitation. Sample loading and data acquisition are automated and rapid, allowing 96 samples to be analyzed in under an hour. Multiplexed substrates are designed to simultaneously analyze multiple substrates, products and/or reaction intermediates in a single reaction, reducing analysis time and costs associated with enzyme characterization, and facilitate the study of complex pathways, such as DNA replication or repair (Fig. 4). Figure 4. View largeDownload slide RNaseH2 analysis by capillary gel electrophoresis (CE). (A) For detection by CE, DNA substrates are labeled with fluorophores. To monitor RNaseH2 activity, DNA (50 nt) containing a single rG is labeled with MAX (green) and FAM (blue). RNaseH2 nicks 5΄ to the rG generating a 21 nt FAM and 29 nt MAX labeled product that are separated and detected by CE (Heider et al.2017). (B) RNaseH2 substrate (50 nt) and product (21 and 29 nt) peaks generated by CE are shown at 0, 0.002 and 0.03 s. Figure 4. View largeDownload slide RNaseH2 analysis by capillary gel electrophoresis (CE). (A) For detection by CE, DNA substrates are labeled with fluorophores. To monitor RNaseH2 activity, DNA (50 nt) containing a single rG is labeled with MAX (green) and FAM (blue). RNaseH2 nicks 5΄ to the rG generating a 21 nt FAM and 29 nt MAX labeled product that are separated and detected by CE (Heider et al.2017). (B) RNaseH2 substrate (50 nt) and product (21 and 29 nt) peaks generated by CE are shown at 0, 0.002 and 0.03 s. +) Analyzing Okazaki fragment maturation by CE Lagging strand DNA replication utilizes Okazaki fragments, which start with short RNA primers that are extended by a replicative DNA polymerase (Fig. 1). Importantly, the RNA primers must then be removed and replaced with DNA by a process known as Okazaki fragment maturation. CE analysis was used to examine the Okazaki fragment maturation process in Thermococcus species 9°N using a dual-color fluorescence assay (Greenough, Kelman and Gardner 2015). The process of joining Okazaki fragments includes the removal of RNA primers at the 5΄ end of each Okazaki fragment, gap filling and ligating the resultant DNA nicks to form a continuous lagging strand. The CE study showed that PolB extends the primer and undergoes strand displacement synthesis to displace the RNA primer, creating a flap structure. Fen1 cleaves the flap and the gap is filled by PolB to generate a nick. Finally, DNA ligase seals the nick to produce a continuous lagging strand. Due to the dual-labeling assay design, both colored reaction substrates, intermediates and products can be quantified in a single experiment. This allows a detailed understanding of the Okazaki fragment maturation process, and coordination amongst enzymes. Interestingly, in CE experiments lacking PolB, PolD left four base pair (bp) gaps between Okazaki fragments on the lagging strand, and maturation was extremely inefficient (Greenough, Kelman and Gardner 2015). Because genetic data (described above) suggest that cells can survive in the presence of only PolD, it is not clear how DNA synthesis proceeds on the lagging strand in the absence of PolB, and which polymerase is involved in filling the 4 bp gaps between Okazaki fragments, along with the removal of RNA primers from the lagging strand. +) Analyzing the kinetic properties of PolD using CE As discussed above, PolD is proposed to be the major leading and lagging strand replicative DNA polymerase in archaea. CE was utilized to study the DNA replication kinetic properties of PolD from Thermococcus species 9°N and to compare them with the properties of other known replicative polymerases (Schermerhorn and Gardner 2015). Using a 5' fluorophore labeled primer/template, rapid analysis of correct, incorrect and ribonucleotide incorporation kinetics, exonuclease kinetics and pyrophosphorolysis were obtained. These studies suggest that PolD nucleotide selectivity is a product of tight binding of correct nucleotide versus incorrect nucleotide. A mechanism within the PolD active site is present to prevent ribonucleotide binding and incorporation, and forward DNA synthesis is favored over exonuclease and pyrophosphorolysis. Similar kinetic properties are associated with many replicative DNA polymerases and support the notion that PolD is the replicative enzyme in archaea (Fiala 2004; Schermerhorn and Gardner 2015). +) Characterizing the archaeal ribonucleotide excision repair pathway using CE Studies performed in eukarya and bacteria report that ribonucleotides are commonly incorporated by replicative DNA polymerases during DNA replication, leading to genome instability (Cerritelli and Crouch 2016). The presence of a ribonucleotide excision repair (RER) pathway, initiated by the enzyme RNaseH2, removes ribonucleotides from the genome (Sparks et al.2012). CE was utilized to characterize the RER pathway in T. kodakarensis (Heider et al.2017). Using a dual-color fluorescent substrate containing a single embedded ribonucleotide, the initiation, intermediates and final repair products of archaeal RER were observed (Fig. 4). These experiments report that RNaseH2 is exclusively responsible for initiating RER repair in T. kodakarensis; that PolB, not PolD, extends the nick, displacing a downstream strand cleaved by Fen1, and DNA ligase completes the repair process. This study highlights the importance of RNaseH2 and suggests a role for PolB, Fen1 and DNA ligase in archaeal DNA damage repair. –) Next-generation sequencing The emergence of NGS technologies within the past decade has enabled cost-effective, rapid sequencing of genomes from all three domains of life. Importantly, due to their small genome size (2–4 Mb), NGS of archaeal species is especially cheap and fast. Such sequencing technologies include Illumina (Fig. 5) and Pacific Bioscience (PacBio) single-molecule real time. An overview of these sequencing platforms can be found in Goodwin, McPherson and McCombie (2016). NGS technologies have been utilized in the archaeal DNA replication and repair field in several different ways, including validation of genetic knockouts, methylation analysis and transcriptome analysis. Some examples are described below. Figure 5. View largeDownload slide Analysis of gene knockouts using next-generation sequencing (NGS). (A) Genomic DNA is sequenced on an NGS platform. Sequencing analysis allows for visualization of mapped sequencing reads. (B) Genomic knockouts of T. kodakarensis GAN, RNaseH2 and Fen1 were confirmed by loss of sequencing reads across each gene, using Illumina sequencing (Burkhart et al.2017). Figure 5. View largeDownload slide Analysis of gene knockouts using next-generation sequencing (NGS). (A) Genomic DNA is sequenced on an NGS platform. Sequencing analysis allows for visualization of mapped sequencing reads. (B) Genomic knockouts of T. kodakarensis GAN, RNaseH2 and Fen1 were confirmed by loss of sequencing reads across each gene, using Illumina sequencing (Burkhart et al.2017). +) Validation of archaeal genomic knockouts using NGS As discussed above, advancements in genetic tools for archaeal species have allowed the knockout of specific DNA replication and repair genes. Importantly, many archaeal species harbor multiple copies of their chromosome, making it difficult to knock out genes of interest and to validate that successful knockout occurred in all copies (Hildenbrand et al.2011; Spaans, van der Oost and Kengen 2015). Typically, labs validate genomic knockouts by PCR amplification of the genomic region in which the desired knockout gene was located, by Southern blot analysis or by antibody detection of the gene product using Western blotting. While these validation methods allow rapid and cheap ways to confirm knockouts, there is the underlying concern that loss of PCR product or antibody detection is due to experimental error rather than gene knockout. Also, due to sensitivity of the assays, such validation methods make it difficult to confirm knockout has occurred in all chromosomal copies. NGS has provided an avenue to validate successful gene knockout. By sequencing the genome of knockout strains, one can map the sequencing reads across the desired knockout region; complete knockout will result in loss of sequencing coverage in that genomic region (Fig. 5). Both Illumina and PacBio sequencing have been used to validate gene knockouts in archaeal organisms. Illumina sequencing was used to validate the deletion of the three origins of replication on the main chromosome of H. volcanii (Hawkins et al.2013). Furthermore, sequencing confirmed that the origin(s) did not translocate to different regions of the chromosome. The ability of H. volcanii to replicate without an origin of replication has changed our understanding of archaeal growth and survival, and suggests other mechanisms must be at play to initiate chromosomal replication (described above). NGS has been used in other archaeal species to validate successful knockout of DNA replication and repair proteins. NGS studies with T. kodakarensis, an archaeal organism that harbors 10–20 copies of its genome, were used to confirm knockouts of Fen1, GAN and RNaseH2 (Burkhart et al.2017). These studies suggested that Fen1, GAN and RNaseH2 provide redundant pathways for 5΄-3΄ RNA primer removal on the lagging strand during Okazaki fragment maturation. This conclusion was the result of the inability to knockout both GAN and Fen1 or GAN and RNaseH2, while the individual knockout of each gene was successful (Burkhart et al.2017) (Fig. 5). The adoption of NGS technologies by the archaeal community to validate genomic knockouts will drive our understanding of essential DNA replication and repair proteins, and help map coordination and/or redundancy amongst important replication and repair players. Another genomic method, transposon-sequencing (Tn-seq), has been used to probe in vivo protein function by inserting transposons into random regions of the genome thereby disrupting the gene (van Opijnen, Bodi and Camilli 2009). Non-essential genes can be disrupted by transposon insertion, while essential genes will not tolerate insertions. NGS is a key for mapping genome-wide transposon insertion sites and determining essential genes. The method was used to identify essential DNA replication and repair genes in Methanococcus maripaludis. Essential genes include those that encode PolD, DNA primase, MCM1, replicaiton protein A (RPA), the archaeal SSB, DNA ligase, PCNA, RFC, topoisomerase and GINS (Sarmiento, Mrazek and Whitman 2013). PolB, on the other hand, was not essential (discussed above). +) Uncovering archaeal DNA methylation patterns using NGS DNA methylation occurs in many archaeal organisms as the product of DNA methyltransferases (MTase). MTases are often a component of restriction-modification defense systems in prokarya, in which an MTase encoded by the host methylates the host genomic DNA, while a restriction endonuclease degrades invading exogenous, unmethylated DNA, for example invading viral DNA (Wilson 1991). MTases may also play a role in genome regulation, so understanding genomic methylation patterns may provide insight into the diversity, regulatory, and defense roles of DNA methylation systems in archaea (Moore, Le and Fan 2013). PacBio sequencing identifies both base sequence and DNA modifications to uncover patterns of methylation in a genome (Rhoads and Au 2015), and is particularly useful for identifying archaeal methylomes due to their small genome size (Blow et al.2016). A complete catalog of known archaeal genome methylation sites can be found at the Restriction Enzyme Database (http://rebase.neb.com). +) Using NGS to explore the archaeal transcriptome and inform DNA replication and repair mechanisms Aside from sequencing of genomic DNA, NGS RNA-seq technology can also be used to sequence cellular RNA levels, and provide information about an organism's transcriptome. Transcriptome analyses of T. kodakarensis and Thermococcus onnurineus NA1 identified transcriptionally active genes, including the multitude of DNA replication and repair genes (Jager et al.2014; Cho et al.2017). In addition, RNA-seq identified regulatory mRNA elements, including primary transciption start sites (TSS) upstream of genes, a consensus sequence for transctription initiation and 5΄-untranslated regions (5΄-UTRs). RNA-seq in T. kodakarensis also uncovered small non-coding RNAs, and primary internal transcriptional start sites (iTSS) and their promoters that occur within an annotated gene. For example, an iTSS was located within the MCM3 gene, and transcriptional initiation at this iTSS suggests a truncated version of MCM3. This truncated version is theorized to play a role in MCM3 maturation. Furthermore, new NGS methods, such as Cappable-seq, enable high-resolution mapping of TSS and iTSS (Ettwiller et al.2016). The use of NGS RNA-seq can provide an understanding of the expression of important DNA repair and replication enzymes in response to cellular stresses and how archaeal organisms regulate these transcripts. +) Using NGS to reveal the archaeal chromatin structure Archaea encode histones that organize DNA into higher order chromatin structures (Sandman and Reeve 2001; Ammar et al.2012). NGS methods have accelerated understanding of archaeal chromatin complex size, location along the chromosome and the role of chromatin positioning in gene regulation. Micrococcal nuclease sequencing (MNase-seq) mapped chromatin complex size and position along the T. kodakarensis chromosome and showed specific organization along the genome (Maruyama et al.2013). –) Single-molecule analysis A variety of assays have been established that allow for single-molecule visualization of DNA replication in real time, including a bead-tether assay and fluorescence visualization techniques. These assays have been used to obtain kinetics parameter of replication and repair enzymes (Tanner and van Oijen 2010; van Oijen and Loparo 2010). However, these studies have been performed at 37°C to study mesophilic enzymes, limiting the ability to study enzymes from thermophilic archaea. To overcome this limitation, a single-molecule bead-tether assay was developed to study enzymes from thermophilic archaeons at high temperature (up to 65°C) (Fig. 6). This technique relies upon the assembly of a DNA construct which mimics an in vivo replication fork, and is composed of λ-DNA (48.5 kb), and four oligonucleotides that are ligated together (Fig. 6A). Importantly, one oligo contains a 5΄-biotin tag (red dot), while another is covalently linked to a magnetic reflective bead (gray sphere) (Fig. 6A). Furthermore, this technique relies upon the construction of a thermostable flow cell that can be heated to 65°C by way of a power source connected to an aluminum block on the outside of the flow cell. The DNA construct is delivered to the flow cell and is anchored by way of a biotin–streptavidin interaction. Processive thermostable enzymes, such as a DNA polymerase or helicase (shown in blue in Fig. 6A), are delivered to the flow cell, and assemble at the synthetic replication fork. Delivery of necessary reagants, such as ATP or dNTPs, to the flow cell initiate enzyme activity on single DNA constructs. The processive activity of an enzyme is visualized by the movement of the magnetic reflective bead tethered to the DNA construct. The movement of the beads is converted to trajectories that can be analyzed to determine rates and distances of processive enzyme activity (Fig. 6A). The use of this technique to examine the kinetics of DNA unwinding of MCM helicases from two archaeal species is discussed below. Figure 6. View largeDownload slide Analysis of MCM processivity using a high-temperature single-molecule bead tether assay. (A) A flow-cell and 5΄-biotin (red circle) labeled λ-DNA construct are used to carry out a high-temperature single-molecule bead tether assay to study DNA unwinding by archaeal MCM helicases. Bead trajectories are analyzed to obtain the rate and distance of DNA unwinding (Schermerhorn et al.2016). (B) The rate and distance of DNA unwinding from 50 events for three MCM helicases, MCM from M. thermautotrophicus (Mth MCM), and MCM2 and MCM3 from Thermococcus species 9°N (9°N MCM2 and 9°N MCM3, respectively), are displayed (Schermerhorn et al.2016) Make sure 2016 is connected to reference down below. Figure 6. View largeDownload slide Analysis of MCM processivity using a high-temperature single-molecule bead tether assay. (A) A flow-cell and 5΄-biotin (red circle) labeled λ-DNA construct are used to carry out a high-temperature single-molecule bead tether assay to study DNA unwinding by archaeal MCM helicases. Bead trajectories are analyzed to obtain the rate and distance of DNA unwinding (Schermerhorn et al.2016). (B) The rate and distance of DNA unwinding from 50 events for three MCM helicases, MCM from M. thermautotrophicus (Mth MCM), and MCM2 and MCM3 from Thermococcus species 9°N (9°N MCM2 and 9°N MCM3, respectively), are displayed (Schermerhorn et al.2016) Make sure 2016 is connected to reference down below. +) Using single-molecule analysis to obtain MCM helicase DNA unwinding parameters In eukarya, MCM is not active and needs to associate with GINS and Cdc45 to form the active CMG complex (described in detail above). Archaeal genomes encode for CMG components and thus it is possible that an archaeal CMG complex is needed for processive DNA unwinding. To test this hypothesis, the high-temperature single-molecule bead tether assay examined the DNA helicase unwinding kinetics of MCM helicases from two thermophilic archaea, Thermococcus sp. 9°N, which harbors multiple MCM copies, and Methanothermobacter thermautotrophicus, which harbors a single MCM (Fig. 5) (Schermerhorn et al.2016). This study revealed that the archaeal MCM helicases alone have robust DNA unwinding activity, unwinding stretches of 14 kb in length at a rate of up to 150 bp/s, and do not require additional factors for processivity. It is important to note that the single-molecule approach allows one to obtain kinetic parameters of processivity and unwinding rate of individual events, and therefore provides a distribution of rates and distances (Fig. 6B). This is in contrast to bulk kinetic methods which provide average rates. This study establishes a system to study long-range DNA synthesis and unwinding kinetics by DNA replication proteins from hyperthermophiles, and will aid in our understanding of the coordination between replisome components during processive DNA unwinding and synthesis. HOW CAN NEW TOOLS DRIVE ARCHAEAL DNA REPLICATION AND REPAIR RESEARCH? The implementation of new genetic, biophysical and molecular tools to study archaeal DNA replication and repair has enabled scientists to begin answering long-standing biological questions, including the necessity of an archaeal CMG complex, how Okazaki fragments are processed and if archaea possess DNA methylation. Importantly, we have just started to understand the complex nature of archaeal DNA replication and repair. In future work, we envision that tools described above will drive further understanding of these processes. Below we highlight a few remaining questions, and speculate how these tools can aid in answering them. +) How is archaeal DNA replication initiated in organisms lacking an origin of replication? The observation that the conserved origin of replication and the initiator proteins Orc1 and Cdc6 can be deleted without major effect on certain archaeal species growth changes our view of the origin of replication in archaea and its role in replication initiation (Kelman and Kelman 2018). Furthermore, such evidence suggests other mechanisms for replication initiation must be at play. One hypothesis is the emergence of dormant, non-conserved origins, which would facilitate bidirectional DNA synthesis. NGS technologies have routinely been used to track DNA replication initiation and termination sites in eukarya and bacteria, and would help uncover dormant origins in archaea (Marchal et al.2017). A second hypothesis is the utilization of HR to allow origin-independent replication initiation. Genetic knockouts of HR components in organisms other than H. volcanii and T. kodakarensis would help to facilitate our understanding and the necessity of HR during DNA replication, while NGS technologies could answer where HR initiates in the genome. It is possible that different archaeal organisms utilize different mechanisms to initiate DNA replication and the NGS approach may help to shed light on these processes. +) What regulates archaeal DNA replication? In bacteria and eukarya, the DNA replication process is highly regulated and many regulatory mechanisms have been identified (Skarstad and Katayama 2013; Parker, Botchan and Berger 2017). To date, only a single archaeal replication factor TIP (described above) has been identified as a potential replication regulator. The use of in vivo protein tagging, pull-downs and genetic knockouts can aid in the discovery of additional factors that regulate the replication process. +) What DNA repair mechanisms protect extremophiles? Due to the extreme environments in which many archaea thrive, and the DNA-damaging nature of these environments, robust mechanisms must exist to maintain the genome. We currently lack a complete understanding of what DNA repair pathways are present in archaea. Genetic knockouts provide an avenue to uncover the role of potential repair proteins, and to assess if they are essential, while in vivo protein tagging may help elucidate repair complexes and pathways. Furthermore, CE analysis can be used to study the substrate specificities and activities of individual DNA repair enzymes, as well as complexes. +) What mechanisms for lesion bypass exist in archaea? All eukaryotic, bacterial and crenarchaeal organisms possess family Y DNA lesion bypass polymerases (PolY), whose role is to perform DNA replication across bulky DNA lesions and prevent stalled replication (Broyde et al.2008). Euryarchaeal organisms lack PolY, and therefore how these organisms handle bulky lesions is a mystery. CE can be utilized to examine the ability of PolB and PolD to synthesize through bulky lesions, and assess how other replication factors may assist in the process. +) How is leading and lagging strand DNA synthesis coordinated? There is evidence to suggest that leading and lagging strand DNA synthesis is well coordinated, allowing the replisome to move along the DNA at a defined rate. Furthermore, coordination between DNA unwinding by the helicase and DNA synthesis by the polymerase must also occur to prevent exposure of ssDNA. How, and if, these events are coordinated in archaeal DNA replication is still unknown. The high-temperature single-molecule assay provides a system to enable the examination of these events, and would help to uncover which factors are needed to couple DNA unwinding and synthesis, and leading and lagging strand replication. 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Construction and analysis of a recombination-deficient (radA) mutant of Haloferax volcanii . Mol Microbiol 1997 ; 23 : 791 – 7 . Yao NY , O’Donnell ME . Evolution of replication machines . Crit Rev Biochem Mol 2016 ; 51 : 135 – 49 . © FEMS 2018. This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/4.0/), which permits non-commercial re-use, distribution, and reproduction in any medium, provided the original work is properly cited. For commercial re-use, please contact journals.permissions@oup.com http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png FEMS Microbiology Reviews Oxford University Press

Archaeal DNA replication and repair: new genetic, biophysical and molecular tools for discovering and characterizing enzymes, pathways and mechanisms

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Abstract

Abstract DNA replication and repair are essential biological processes needed for the survival of all organisms. Although these processes are fundamentally conserved in the three domains, archaea, bacteria and eukarya, the proteins and complexes involved differ. The genetic and biophysical tools developed for archaea in the last several years have accelerated the study of DNA replication and repair in this domain. In this review, the current knowledge of DNA replication and repair processes in archaea will be summarized, with emphasis on the contribution of genetics and other recently developed biophysical and molecular tools, including capillary gel electrophoresis, next-generation sequencing and single-molecule approaches. How these new tools will continue to drive archaeal DNA replication and repair research will also be discussed. archaea, capillary gel electrophoresis, DNA repair, DNA replication, genetics, next-generation sequencing INTRODUCTION DNA replication plays an essential role in all life forms. The process is required for the propagation and evolution of genetic information. The mechanism of DNA replication guarantees the duplication and transfer of genetic information during cell division. Although the process needs to be precise, the fidelity of the replication process is not absolute and other cellular and environmental insults result in DNA damage and alterations in the DNA sequence. Consequently, another process, DNA repair, is also required to ensure the integrity of chromosomal DNA. In all organisms, chromosomal DNA replication is a complex event that involves many factors to ensure the accurate and timely duplication of genetic information. The mechanism of DNA replication is fundamentally conserved in all life forms (reviewed in Kelman and Kelman 2014; Yao and O’Donnell 2016). The process is thought to start at a specific sequence called an origin of replication, at which origin-binding proteins (OBP) bind and locally unwind duplex DNA. Additional proteins interact with the OBP-DNA complex and are responsible for the assembly of the DNA helicase around the DNA. Once assembled, the helicase unwinds the duplex and forms the initial replication bubble. The exposed single-stranded (ss) DNA at the replication bubble is coated with ssDNA-binding protein (SSB). DNA primase, DNA polymerases (Pol) and the rest of the replication machinery are recruited to the SSB-ssDNA complex to initiate bidirectional DNA synthesis. Due to the antiparallel nature of duplex DNA, and the unidirectionality of DNA polymerases, one strand of the chromosome is synthesized continuously (leading strand) while the other is copied discontinuously (lagging strand) as a series of Okazaki fragments (Fig. 1). Figure 1. View largeDownload slide New tools to study archaeal DNA replication and repair. A model for the archaeal replication fork. The leading and lagging strands are shown as well as the proteins involved in Okazaki fragment maturation, and ribonucleotide excision repair. The tools described in the review used to study these processes are shown: single-molecule analysis (pink), next-generation sequencing (orange), genetics (green) and capillary electrophoresis (blue). Figure 1. View largeDownload slide New tools to study archaeal DNA replication and repair. A model for the archaeal replication fork. The leading and lagging strands are shown as well as the proteins involved in Okazaki fragment maturation, and ribonucleotide excision repair. The tools described in the review used to study these processes are shown: single-molecule analysis (pink), next-generation sequencing (orange), genetics (green) and capillary electrophoresis (blue). As cells grow and divide, they are continuously exposed to DNA-damaging agents that arise endogenously by cellular metabolism, or from external environmental factors. The exposure to DNA-damaging agents leads to the formation of a variety of DNA lesions that are mutagenic and/or cytotoxic to the cell, and can halt DNA replication and cause cellular apoptosis. Due to the toxic nature of DNA lesions, organisms have evolved several DNA repair pathways including direct damage reversal, base excision repair (BER), nucleotide excision repair (NER), non-homologous end joining (NHEJ) and homologous recombination (HR). The extreme environments in which many archaeal species thrive, including high temperature, high salinity or low pH, facilitate DNA damage and lesion formation, and thus archaea likely require robust DNA repair mechanisms for genome maintenance. For many years, the study of archaeal DNA replication and repair was hindered by a lack of genetic and other biophysical tools. Therefore, most of the information regarding these processes was derived from biochemical, structural and limited cellular studies. However, in the last decade new tools have been developed in several archaeal species including genetic tools, such as the knockin and knockout of genes, and biophysical and molecular tools, such as capillary gel electrophoresis (CE), single-molecule analysis of individual proteins and processes, and next-generation sequencing (NGS) (Fig. 1). These assays more accurately reflect biological processes in the cell and accelerate understanding of enzymes and pathways. In this review, the impact of recently developed tools on the study of the replication and repair processes in archaea will be summarized. We also speculate how these new tools will drive future archaeal DNA replication and repair research. Readers are referred to several past reviews for previously summarized results (Kelman and White 2005; Barry and Bell 2006; White 2011; Grasso and Tell 2014; Kelman and Kelman 2014). GENETIC TOOLS Genetic tools allow for the in vivo manipulation of genes within an organism, including the introduction, modification or deletion of a particular gene of interest. Importantly, genetic tools allow one to understand the importance, or lack thereof, of a particular gene. Genetic tool systems have long been established in eukaryotic and bacterial organisms, including the routinely used yeast Saccharomyces cerevisiae and the Escherichia coli bacterium, as well as in a variety of higher organisms (Doudna and Charpentier 2014; Duina et al.2014; Blount 2015). In the last several years, robust genetic tools have also been developed for several archaeal species from Euryarchaeota and Crenarchaeota phyla (Atomi, Imanaka and Fukui 2012). Species include Methanosarcina acetivorans (Kohler and Metcalf 2012), Haloferax volcanii (Leigh et al.2011), Pyrococcus furiosus (Lipscomb et al.2011), Thermococcus kodakarensis (Hileman and Santangelo 2012; Farkas et al.2013) and Sulfolobus solfataricus (Wagner et al.2012). As the replication machinery of crenarchaea is different from all other archaeal branches, we will concentrate on the tools developed in euryarchaea and their impact on the fields of DNA replication and repair. –) Gene knockout When genetic tools became available for archaea, one of the first questions asked was: Which replication and repair proteins are essential for viability? To answer this question, two main genetic approaches were used. One approach is the direct deletion of a specific gene or region of the chromosome (Fig. 2), while the other uses random integration of transposons to disrupt gene function to identify essential genes (Sarmiento, Mrazek and Whitman 2013). Most results were as expected; proteins that were shown to be essential in bacteria and/or eukarya were also essential in archaea, and those that were not essential in the other domains were not essential in archaea. For example, the polymerase accessory proteins, replication factor C (RFC) and proliferating cell nuclear antigen (PCNA) are essential in both eukarya and archaea (Haracska et al.2001; Kuba et al.2012; Pan et al.2013; Sarmiento, Mrazek and Whitman 2013), while Flap endonuclease 1 (Fen1), an enzyme that removes 5΄ overhangs generated during Okazaki fragment maturation and DNA repair, is dispensable for viability in both (Reagan et al.1995; Burkhart et al.2017). However, there were also unexpected results; several of those will be described here as examples. Figure 2. View largeDownload slide A genetic knockout system for T. kodakarensis. A target gene (yellow) is deleted from a recipient strain via two homologous recombination events using donor DNA containing selectable marker (SM), and CSM (counter selectable marker), for use as positive and negative selection. This approach results in the construction of markerless deletion strains (see Hileman and Santangelo 2012 for details). Figure 2. View largeDownload slide A genetic knockout system for T. kodakarensis. A target gene (yellow) is deleted from a recipient strain via two homologous recombination events using donor DNA containing selectable marker (SM), and CSM (counter selectable marker), for use as positive and negative selection. This approach results in the construction of markerless deletion strains (see Hileman and Santangelo 2012 for details). +) Genetic knockouts of a conserved archaeal origin of replication and origin initiator proteins One of the most exciting and unexpected observations made using knockout experiments in archaea is the observation that several archaeal species do not require a traditional conserved origin of replication. It is well established that organisms require an origin of replication to initiate chromosomal replication and to provide a point of regulation for the replication process. Importantly, while bacterial organisms contain a single conserved origin of replication, higher eukaryotes contain many less-defined origins (Kornberg and Baker 1992; Boulikas 1996; DePamphilis 1996; O’Donnell, Langston and Stillman 2013). It has been proposed that archaea use conserved origins, similar to bacteria, due to bioinformatic studies that identified a conserved archaeal sequence containing known origin of replication motifs (Norais et al.2007). Interestingly, gene knockout studies have shown that in some archaeal species these origins are dispensable for cell viability and can be readily deleted in laboratory strains [(Hawkins et al.2013; Gehring et al.2017) summarized in (Kelman and Kelman 2018)]. A study demonstrating that the origin can be deleted led to even more surprising results. Under normal laboratory growth conditions, T. kodakarensis does not utilize the predicted conserved origin of replication to initiate DNA synthesis (Gehring et al.2017). Furthermore, a secondary origin of replication was not identified, suggesting random initiation of DNA replication across the chromosome. This interesting, dogma-shattering observation questions the necessity of the origin of replication in archaea, and will require further investigation. It is important to reevaluate under which conditions the conserved origin of replication is used, why it is retained in the genome and how DNA replication is initiated on a chromosome in strains lacking a conserved origin (Hawkins et al.2013). Archaea contain homologs of several subunits of the eukaryotic origin recognition complex (Orc), in particular Orc1 and the replication initiation protein Cdc6 (referred to as Orc1/Cdc6 in archaea). These archaeal homologs were shown to bind to the archaeal origin of replication and were suggested to initiate the replication process by assembling the helicase (Matsunaga et al.2001; Costa, Hood and Berger 2013). Similar to the observation that the origin of replication is dispensable in some archaeal species, it was shown that the Orc1/Cdc6 gene can also be deleted in those species (Gehring et al.2017). In bacteria and eukarya, the OBPs and initiation proteins participate in regulating the initiation process. Further study is needed to determine which proteins and processes regulate the initiation of replication in the absence of Orc1/Cdc6. +) Genetic knockouts of archaeal DNA polymerases Most archaeal linages, except for crenarchaea, contain two different polymerases, a family B DNA polymerase, PolB, and a family D DNA polymerase, PolD (Cann et al.1998; Ishino et al.1998). The three replicative polymerases in eukarya, Polα, Polε and Polδ, all belong to family B, and crenarchaea contain only PolB homologs. Therefore, it was proposed that members of the PolB family are responsible for chromosomal replication in all of archaea (Grabowski and Kelman 2003; Johansson and Dixon 2013). However, in many archaeal species, the genes encoding the two archaeal-specific subunits of PolD are located in close proximity to the origin of replication and are often in an operon with other replication enzymes (for example, see Kelman 2000). The proximity of proteins involved in chromosomal replication to the origin of replication is common in bacteria and archaea (Kornberg and Baker 1992). In addition, the small exonuclease subunit of PolD shares amino acid sequence similarity with several of the small non-catalytic subunits of eukaryotic Polα, Polδ and Polε (Aravind and Koonin 1998). It was therefore proposed that PolD might also function at the archaeal replication fork, or that both PolB and PolD may be present at the fork. This would be similar to the situation in eukarya in which Polε replicates the leading strand, while Polδ copies the lagging strand (reviewed in Walsh and Eckert 2014). Early genetic studies supported the idea that both PolB and PolD may be present at the fork, as studies with Halobacterium indicated that both PolB and PolD may be essential for cell viability (Berquist, DasSarma and DasSarma 2007). However, studies in other archaeal species showed that PolB was dispensable for cell viability, while PolD was essential (Čuboňová et al.2013; Sarmiento, Mrazek and Whitman 2013). This result was unexpected and changes our understanding of chromosomal replication in at least some archaeal species. These data suggest that PolD (in the absence of PolB) could carry out both leading and lagging strand synthesis. It is possible, however, that when both polymerases are present in the cell, PolB, with its strand displacement activity (Greenough, Kelman and Gardner 2015) replicates the lagging strand, while PolD replicates the leading strand. +) Genetic knockouts help define the roles of archaeal Cdc45, GINS and MCM in DNA replication In eukarya, the heterohexameric minichromosome maintenance (MCM) helicase, the major replicative helicase, is not active on its own, but is activated by association with two accessory factors, the tetrameric GINS complex and the Cdc45 protein, forming the CMG (Cdc45, MCM, GINS) complex (reviewed in Onesti and MacNeill 2013; Li and O’Donnell 2018). In contrast to eukarya, the archaeal MCM helicase is active on its own in vitro without the requirement for additional factors (Kelman, Lee and Hurwitz 1999; Chong et al.2000 summarized in Costa and Onesti 2009; Sakakibara, Kelman and Kelman 2009). Homologs of the eukayotic GINS complex and Cdc45 have been identified in the archaeal genomes (Makarova et al.2005; Marinsek et al.2006; Makarova, Koonin and Kelman 2012; Oyama et al.2016 reviewed in Bell 2011). While the eukaryotic GINS complex is a ring-shaped heterotetramer of four related but different polypeptides, most archaeal GINS are tetramers compromised of two different proteins (MacNeill 2010). While the eukaryotic Cdc45 is essential for cell viability (Onesti and MacNeill 2013), in some archaeal species the Cdc45 homolog can be readily deleted with no effect on cell growth under normal growth conditions (Burkhart et al.2017 and Stuart MacNeill, personal communication). In contrast, the MCM and GINS proteins are essential in archaea (Kelman and Kelman 2014 and references therein). Nevertheless, it is possible that archaeal MCM, Cdc45 and GINS form a replicative helicase complex in vivo, as they are shown to form a stable complex in vivo (Marinsek et al.2006; Nagata et al.2017). +) Discovery of the archaeal primase using genetic knockouts In most archaeal genomes, a homolog of the bacterial primase, DnaG, has been identified. It was originally thought that this protein was the archaeal primase. However, studies have identified a two-subunit complex in archaea that is similar to the two-subunit eukaryotic primase (Bocquier et al.2001). Genetic studies showed that this two-subunit eukaryotic-like primase is the archaeal primase, not DnaG. While the gene encoding DnaG can be readily deleted, the genes encoding the eukaryotic-like primase cannot (Le Breton et al.2007). The archaeal primase is a two-subunit complex with similarities to the two-subunit eukayotic primase; a small subunit, PriS, contains the catalytic activity, and a large subunit, PriL, regulates primase activity (Bocquier et al.2001; Chemnitz Galal et al.2012 reviewed in Lao-Sirieix, Pellegrini and Bell 2005). Both subunits were shown to be essential for cell viability (Sarmiento, Mrazek and Whitman 2013). +) Genetic knockouts of archaeal DNA repair genes The use of genetic knockout techniques has been used to examine which archaeal DNA repair genes are essential, those that are non-essential and those in which knockout leads to phenotypic changes. Early experiments in H. volcanii lead to successful knockout of radA, which catalyzes strand exchange during HR (Woods and Dyall-Smith 1997). However, the ΔradA strain had severe HR defects and increased sensitivity to DNA-damaging agents. In T. kodakarensis, attempts to knock out several genes encoding HR proteins, including radA, rad50 and mre11, were unsuccessful, suggesting HR may be essential for T. kodakarensis survival, and not H. volcanii (Fujikane et al.2010). On the other hand, in T. kodakarensis, the genes encoding for xeroderma pigmentosum type B and D (XPB and XPD) helicase proteins can be readily deleted, and the knockout strains showed little or no sensitivity when challenged with UV radiation (Fujikane et al.2010). It is well established that mutations in XPB and XPD proteins in mammalians leads to xeroderma pigmentosum, a genetic disorder which renders an individual unable to repair DNA damage caused by UV-light exposure (reviewed in Lehmann, McGibbon and Stefanini 2011). The role XPB and XPD play in archaeal DNA repair is still unclear. Several other DNA repair genes have successfully been knocked out in T. kodakarensis, including Hjm, a RecQ like helicase, Hjc, a structure specific endonuclease, Hef, a helicase/nuclease whose role is proposed to help stalled replication forks, Fen1, a 5’ flap endonuclease, and RNaseH2, an endonuclease that cleaves at rNMP:dNMP junctions (Fujikane et al.2010; Burkhart et al.2017). +) Uncovering the role of multiple gene copies using genetic knockouts There are examples of genes that are present in a single copy in certain archaeal species, but present in multiple copies in other species. Gene knockouts were used to determine which gene copies are essential and which, if any, are dispensable. A few examples are described below. The vast majority of the known archaeal genomes encode for a single homolog of the MCM helicase, forming homohexamers (Sakakibara, Kelman and Kelman 2009). The genomes of several archaeal species, however, contain multiple MCM homologs. The presence of multiple MCM proteins in a single archaeon led to the proposal that in those organisms the MCM complex is similar to the eukaryotic helicase, where multiple subunits form a heterohexamer complex at the replication fork (Sakakibara, Kelman and Kelman 2009 and references therein). Using gene knockout experiments, it was found that in archaeal species that contain multiple MCM homologs, only one MCM homolog was essential for cell viability (for example, Ishino et al.2011; Pan et al.2011), suggesting they are similar to archaea containing a single MCM homolog with a homohexameric MCM helicase active at the replication fork. Another example is the polymerase processivity factor, PCNA. To date, all archaeal species encode for a single PCNA homolog forming a homotrimeric ring. There are two exceptions. One is organisms belonging to the crenarchaea in which three PCNA homologs are present, forming a heterotrimeric PCNA (Daimon et al.2002; Dionne et al.2003), and the other exception is the genome of T. kodakarensis that encodes for two PCNA homologs (Pan, Kelman and Kelman 2011). It is not clear why two PCNA proteins are present in the T. kodakarensis genome, or whether they have redundant functions. Although both PCNA proteins form similar homotrimeric rings (Ladner et al.2011) and can stimulate the activity of several enzymes (Ladner et al.2011; Kuba et al.2012; Li et al.2014), only one is essential for viability (Kuba et al.2012; Pan et al.2013). These results show that T. kodakarensis is similar to all other archaea species that contain only one PCNA protein. Non-essential gene copies, as in the case of MCM and PCNA in T. kodakarensis, are typically the result of gene duplication by viral integration (Fukui et al.2005). Importantly, all virally integrated regions can be removed from T. kodakarensis, without substantial effects on cell growth (Tagashira et al.2013). –) In vivo protein tagging The use of in vivo protein tagging allows for the incorporation of an affinity or detection tag at the genomic level to capture or visualize the resulting protein. The creation of genetic tools in archaeal systems has enabled archaeal in vivo protein tagging, which has been utilized to discover new archaeal DNA replication factors (Fig. 3). Figure 3. View largeDownload slide In vivo protein tagging. (A) Strains are constructed that encode for His-tagged replication and repair proteins. Stable in vivo complexes formed with the His-tagged protein are purified from clarified cell lysates. For example, His-tagged PCNA (black) forms a stable complex with Fen1 (purple), PolB (orange) and DNA ligase (pink). Co-purified proteins are identified by mass spectrometry. (B) Thermococcus kodakarensis PCNA is a central DNA replication enzyme and 12 proteins were co-purified with a tagged version of PCNA (Li et al.2010). (C) Thermococcales inhibitor of PCNA (TIP) was co-purified with PCNA and the proteins form a complex in vitro (PDB ID: 5DA7) (Altieri et al.2016). Orange, TIP; blue, PCNA. Figure 3. View largeDownload slide In vivo protein tagging. (A) Strains are constructed that encode for His-tagged replication and repair proteins. Stable in vivo complexes formed with the His-tagged protein are purified from clarified cell lysates. For example, His-tagged PCNA (black) forms a stable complex with Fen1 (purple), PolB (orange) and DNA ligase (pink). Co-purified proteins are identified by mass spectrometry. (B) Thermococcus kodakarensis PCNA is a central DNA replication enzyme and 12 proteins were co-purified with a tagged version of PCNA (Li et al.2010). (C) Thermococcales inhibitor of PCNA (TIP) was co-purified with PCNA and the proteins form a complex in vitro (PDB ID: 5DA7) (Altieri et al.2016). Orange, TIP; blue, PCNA. It is well established that proteins involved in the same pathway often form a stable complex. This is particularly true for the replisome complex, which is responsible for the duplication of chromosomal DNA (Yao and O’Donnell 2016). Although many archaeal replication and repair proteins and complexes have been identified based on their similarity to a bacterial or eukaryal counterpart, it is likely that some factors required for archaeal replication and regulation have yet to be identified (Kelman and Kelman 2014). Several known replication proteins were in vivo tagged with a 6 His-tag using a knockin approach (Li et al.2010) (Fig. 3). The insertion of the His-tag did not change the genomic context, including regulatory sequences, or position in an operon. Following purification on a Ni2+ column, the proteins that co-purified with the tagged protein were identified using mass spectrometry. Some of these interactions support previously reported biochemical and genetic studies. For example, it was found that the archaeal DnaG homolog does not interact with any known replication factors (Li et al.2010) but rather with exosome proteins, as was previously reported (Evguenieva-Hackenberg et al.2003), thus providing further evidence that DnaG is not the archaeal primase. Another example is the observation that PolB does not interact with many replisome proteins but PolD does. This study also identified a number of potentially new replication factors. Two examples are described below. +) Using in vivo protein tagging to discover the archaeal Cdc45 homolog As described above, the CMG complex in eukarya forms the replicative helicase required for the duplication of chromosomal DNA (Onesti and MacNeill 2013; Li and O’Donnell 2018). For many years, the components of the MCM and the GINS complexes were identified in archaea by similarity to eukaryotic homologus. However, no clear homolog of the Cdc45 protein could be identified by amino acid sequence comparison. The first suggestion that archaea might contain a Cdc45 homolog came from in vivo tagging experiments (Li et al.2010). A protein with homology to the bacterial RecJ nuclease co-eluted with the GINS complex. Furthermore, in silico analysis and structural prediction suggested that the protein, referred to as GAN (GINS-associated nuclease), shares similarity with RecJ and the eukaryotic Cdc45 protein (Makarova, Koonin and Kelman 2012; Oyama et al.2016). It was subsequently shown that the archaeal and eukaryal proteins have similar three-dimensional structures (Oyama et al.2016). The availability of genetic tools accelerated the discovery of GAN, underscoring the power and importance of in vivo tagging. +) Using in vivo protein tagging to uncover an archaeal DNA replication regulator protein PCNA plays an essential role in both DNA replication and repair (Moldovan, Pfander and Jentsch 2007). In eukarya, a number of mechanisms regulate the activity of PCNA during replication and repair, including post-translational modification and association with regulatory proteins (Vivona and Kelman 2003; López de Saro 2009; Dieckman, Freudenthal and Washington 2012). Until the development and utilization of archaeal in vivo tagging, no regulator of the archaeal PCNA had been reported. However, when the PCNA protein was tagged, a small protein co-eluted with it on Ni2+ column. The protein, referred to as TIP (Thermococcales inhibitor of PCNA), forms a complex with PCNA in vivo (Li et al.2010) and in vitro (Li et al.2014; Altieri et al.2016), and binding of TIP to PCNA inhibits PCNA activities (Li et al.2014) (Fig. 3). TIP is currently the only known protein that may regulate DNA replication, and possibly also DNA repair, in archaea. The emergence of archaeal protein tagging facilitated the discovery of this small protein. MOLECULAR AND BIOPHYSICAL TOOLS New molecular and biophysical tools are needed to discover new archaeal DNA replication and repair enzymes, to understand their function, and to explore cellular pathways and mechanisms. Sensitive assays will provide a clear picture of enzyme activities and interactions. Because enzymes do not function in isolation, but rather as complexes, assays that monitor coupled reactions will also be critical for understanding pathways occurring in the cell. To address these needs, various approaches to study single enzymes, protein complexes and in vivo processes using CE, NGS and single-molecule analysis have been developed and will be described here. –) Capillary gel electrophoresis CE is a sensitive high-throughput, high-resolution system for nucleic acid analysis, and serves as an alternative to gel-based electrophoresis analysis (Greenough et al.2016). In CE, fluorescently labeled nucleic acids are separated by size and charge, and detected by laser excitation. Sample loading and data acquisition are automated and rapid, allowing 96 samples to be analyzed in under an hour. Multiplexed substrates are designed to simultaneously analyze multiple substrates, products and/or reaction intermediates in a single reaction, reducing analysis time and costs associated with enzyme characterization, and facilitate the study of complex pathways, such as DNA replication or repair (Fig. 4). Figure 4. View largeDownload slide RNaseH2 analysis by capillary gel electrophoresis (CE). (A) For detection by CE, DNA substrates are labeled with fluorophores. To monitor RNaseH2 activity, DNA (50 nt) containing a single rG is labeled with MAX (green) and FAM (blue). RNaseH2 nicks 5΄ to the rG generating a 21 nt FAM and 29 nt MAX labeled product that are separated and detected by CE (Heider et al.2017). (B) RNaseH2 substrate (50 nt) and product (21 and 29 nt) peaks generated by CE are shown at 0, 0.002 and 0.03 s. Figure 4. View largeDownload slide RNaseH2 analysis by capillary gel electrophoresis (CE). (A) For detection by CE, DNA substrates are labeled with fluorophores. To monitor RNaseH2 activity, DNA (50 nt) containing a single rG is labeled with MAX (green) and FAM (blue). RNaseH2 nicks 5΄ to the rG generating a 21 nt FAM and 29 nt MAX labeled product that are separated and detected by CE (Heider et al.2017). (B) RNaseH2 substrate (50 nt) and product (21 and 29 nt) peaks generated by CE are shown at 0, 0.002 and 0.03 s. +) Analyzing Okazaki fragment maturation by CE Lagging strand DNA replication utilizes Okazaki fragments, which start with short RNA primers that are extended by a replicative DNA polymerase (Fig. 1). Importantly, the RNA primers must then be removed and replaced with DNA by a process known as Okazaki fragment maturation. CE analysis was used to examine the Okazaki fragment maturation process in Thermococcus species 9°N using a dual-color fluorescence assay (Greenough, Kelman and Gardner 2015). The process of joining Okazaki fragments includes the removal of RNA primers at the 5΄ end of each Okazaki fragment, gap filling and ligating the resultant DNA nicks to form a continuous lagging strand. The CE study showed that PolB extends the primer and undergoes strand displacement synthesis to displace the RNA primer, creating a flap structure. Fen1 cleaves the flap and the gap is filled by PolB to generate a nick. Finally, DNA ligase seals the nick to produce a continuous lagging strand. Due to the dual-labeling assay design, both colored reaction substrates, intermediates and products can be quantified in a single experiment. This allows a detailed understanding of the Okazaki fragment maturation process, and coordination amongst enzymes. Interestingly, in CE experiments lacking PolB, PolD left four base pair (bp) gaps between Okazaki fragments on the lagging strand, and maturation was extremely inefficient (Greenough, Kelman and Gardner 2015). Because genetic data (described above) suggest that cells can survive in the presence of only PolD, it is not clear how DNA synthesis proceeds on the lagging strand in the absence of PolB, and which polymerase is involved in filling the 4 bp gaps between Okazaki fragments, along with the removal of RNA primers from the lagging strand. +) Analyzing the kinetic properties of PolD using CE As discussed above, PolD is proposed to be the major leading and lagging strand replicative DNA polymerase in archaea. CE was utilized to study the DNA replication kinetic properties of PolD from Thermococcus species 9°N and to compare them with the properties of other known replicative polymerases (Schermerhorn and Gardner 2015). Using a 5' fluorophore labeled primer/template, rapid analysis of correct, incorrect and ribonucleotide incorporation kinetics, exonuclease kinetics and pyrophosphorolysis were obtained. These studies suggest that PolD nucleotide selectivity is a product of tight binding of correct nucleotide versus incorrect nucleotide. A mechanism within the PolD active site is present to prevent ribonucleotide binding and incorporation, and forward DNA synthesis is favored over exonuclease and pyrophosphorolysis. Similar kinetic properties are associated with many replicative DNA polymerases and support the notion that PolD is the replicative enzyme in archaea (Fiala 2004; Schermerhorn and Gardner 2015). +) Characterizing the archaeal ribonucleotide excision repair pathway using CE Studies performed in eukarya and bacteria report that ribonucleotides are commonly incorporated by replicative DNA polymerases during DNA replication, leading to genome instability (Cerritelli and Crouch 2016). The presence of a ribonucleotide excision repair (RER) pathway, initiated by the enzyme RNaseH2, removes ribonucleotides from the genome (Sparks et al.2012). CE was utilized to characterize the RER pathway in T. kodakarensis (Heider et al.2017). Using a dual-color fluorescent substrate containing a single embedded ribonucleotide, the initiation, intermediates and final repair products of archaeal RER were observed (Fig. 4). These experiments report that RNaseH2 is exclusively responsible for initiating RER repair in T. kodakarensis; that PolB, not PolD, extends the nick, displacing a downstream strand cleaved by Fen1, and DNA ligase completes the repair process. This study highlights the importance of RNaseH2 and suggests a role for PolB, Fen1 and DNA ligase in archaeal DNA damage repair. –) Next-generation sequencing The emergence of NGS technologies within the past decade has enabled cost-effective, rapid sequencing of genomes from all three domains of life. Importantly, due to their small genome size (2–4 Mb), NGS of archaeal species is especially cheap and fast. Such sequencing technologies include Illumina (Fig. 5) and Pacific Bioscience (PacBio) single-molecule real time. An overview of these sequencing platforms can be found in Goodwin, McPherson and McCombie (2016). NGS technologies have been utilized in the archaeal DNA replication and repair field in several different ways, including validation of genetic knockouts, methylation analysis and transcriptome analysis. Some examples are described below. Figure 5. View largeDownload slide Analysis of gene knockouts using next-generation sequencing (NGS). (A) Genomic DNA is sequenced on an NGS platform. Sequencing analysis allows for visualization of mapped sequencing reads. (B) Genomic knockouts of T. kodakarensis GAN, RNaseH2 and Fen1 were confirmed by loss of sequencing reads across each gene, using Illumina sequencing (Burkhart et al.2017). Figure 5. View largeDownload slide Analysis of gene knockouts using next-generation sequencing (NGS). (A) Genomic DNA is sequenced on an NGS platform. Sequencing analysis allows for visualization of mapped sequencing reads. (B) Genomic knockouts of T. kodakarensis GAN, RNaseH2 and Fen1 were confirmed by loss of sequencing reads across each gene, using Illumina sequencing (Burkhart et al.2017). +) Validation of archaeal genomic knockouts using NGS As discussed above, advancements in genetic tools for archaeal species have allowed the knockout of specific DNA replication and repair genes. Importantly, many archaeal species harbor multiple copies of their chromosome, making it difficult to knock out genes of interest and to validate that successful knockout occurred in all copies (Hildenbrand et al.2011; Spaans, van der Oost and Kengen 2015). Typically, labs validate genomic knockouts by PCR amplification of the genomic region in which the desired knockout gene was located, by Southern blot analysis or by antibody detection of the gene product using Western blotting. While these validation methods allow rapid and cheap ways to confirm knockouts, there is the underlying concern that loss of PCR product or antibody detection is due to experimental error rather than gene knockout. Also, due to sensitivity of the assays, such validation methods make it difficult to confirm knockout has occurred in all chromosomal copies. NGS has provided an avenue to validate successful gene knockout. By sequencing the genome of knockout strains, one can map the sequencing reads across the desired knockout region; complete knockout will result in loss of sequencing coverage in that genomic region (Fig. 5). Both Illumina and PacBio sequencing have been used to validate gene knockouts in archaeal organisms. Illumina sequencing was used to validate the deletion of the three origins of replication on the main chromosome of H. volcanii (Hawkins et al.2013). Furthermore, sequencing confirmed that the origin(s) did not translocate to different regions of the chromosome. The ability of H. volcanii to replicate without an origin of replication has changed our understanding of archaeal growth and survival, and suggests other mechanisms must be at play to initiate chromosomal replication (described above). NGS has been used in other archaeal species to validate successful knockout of DNA replication and repair proteins. NGS studies with T. kodakarensis, an archaeal organism that harbors 10–20 copies of its genome, were used to confirm knockouts of Fen1, GAN and RNaseH2 (Burkhart et al.2017). These studies suggested that Fen1, GAN and RNaseH2 provide redundant pathways for 5΄-3΄ RNA primer removal on the lagging strand during Okazaki fragment maturation. This conclusion was the result of the inability to knockout both GAN and Fen1 or GAN and RNaseH2, while the individual knockout of each gene was successful (Burkhart et al.2017) (Fig. 5). The adoption of NGS technologies by the archaeal community to validate genomic knockouts will drive our understanding of essential DNA replication and repair proteins, and help map coordination and/or redundancy amongst important replication and repair players. Another genomic method, transposon-sequencing (Tn-seq), has been used to probe in vivo protein function by inserting transposons into random regions of the genome thereby disrupting the gene (van Opijnen, Bodi and Camilli 2009). Non-essential genes can be disrupted by transposon insertion, while essential genes will not tolerate insertions. NGS is a key for mapping genome-wide transposon insertion sites and determining essential genes. The method was used to identify essential DNA replication and repair genes in Methanococcus maripaludis. Essential genes include those that encode PolD, DNA primase, MCM1, replicaiton protein A (RPA), the archaeal SSB, DNA ligase, PCNA, RFC, topoisomerase and GINS (Sarmiento, Mrazek and Whitman 2013). PolB, on the other hand, was not essential (discussed above). +) Uncovering archaeal DNA methylation patterns using NGS DNA methylation occurs in many archaeal organisms as the product of DNA methyltransferases (MTase). MTases are often a component of restriction-modification defense systems in prokarya, in which an MTase encoded by the host methylates the host genomic DNA, while a restriction endonuclease degrades invading exogenous, unmethylated DNA, for example invading viral DNA (Wilson 1991). MTases may also play a role in genome regulation, so understanding genomic methylation patterns may provide insight into the diversity, regulatory, and defense roles of DNA methylation systems in archaea (Moore, Le and Fan 2013). PacBio sequencing identifies both base sequence and DNA modifications to uncover patterns of methylation in a genome (Rhoads and Au 2015), and is particularly useful for identifying archaeal methylomes due to their small genome size (Blow et al.2016). A complete catalog of known archaeal genome methylation sites can be found at the Restriction Enzyme Database (http://rebase.neb.com). +) Using NGS to explore the archaeal transcriptome and inform DNA replication and repair mechanisms Aside from sequencing of genomic DNA, NGS RNA-seq technology can also be used to sequence cellular RNA levels, and provide information about an organism's transcriptome. Transcriptome analyses of T. kodakarensis and Thermococcus onnurineus NA1 identified transcriptionally active genes, including the multitude of DNA replication and repair genes (Jager et al.2014; Cho et al.2017). In addition, RNA-seq identified regulatory mRNA elements, including primary transciption start sites (TSS) upstream of genes, a consensus sequence for transctription initiation and 5΄-untranslated regions (5΄-UTRs). RNA-seq in T. kodakarensis also uncovered small non-coding RNAs, and primary internal transcriptional start sites (iTSS) and their promoters that occur within an annotated gene. For example, an iTSS was located within the MCM3 gene, and transcriptional initiation at this iTSS suggests a truncated version of MCM3. This truncated version is theorized to play a role in MCM3 maturation. Furthermore, new NGS methods, such as Cappable-seq, enable high-resolution mapping of TSS and iTSS (Ettwiller et al.2016). The use of NGS RNA-seq can provide an understanding of the expression of important DNA repair and replication enzymes in response to cellular stresses and how archaeal organisms regulate these transcripts. +) Using NGS to reveal the archaeal chromatin structure Archaea encode histones that organize DNA into higher order chromatin structures (Sandman and Reeve 2001; Ammar et al.2012). NGS methods have accelerated understanding of archaeal chromatin complex size, location along the chromosome and the role of chromatin positioning in gene regulation. Micrococcal nuclease sequencing (MNase-seq) mapped chromatin complex size and position along the T. kodakarensis chromosome and showed specific organization along the genome (Maruyama et al.2013). –) Single-molecule analysis A variety of assays have been established that allow for single-molecule visualization of DNA replication in real time, including a bead-tether assay and fluorescence visualization techniques. These assays have been used to obtain kinetics parameter of replication and repair enzymes (Tanner and van Oijen 2010; van Oijen and Loparo 2010). However, these studies have been performed at 37°C to study mesophilic enzymes, limiting the ability to study enzymes from thermophilic archaea. To overcome this limitation, a single-molecule bead-tether assay was developed to study enzymes from thermophilic archaeons at high temperature (up to 65°C) (Fig. 6). This technique relies upon the assembly of a DNA construct which mimics an in vivo replication fork, and is composed of λ-DNA (48.5 kb), and four oligonucleotides that are ligated together (Fig. 6A). Importantly, one oligo contains a 5΄-biotin tag (red dot), while another is covalently linked to a magnetic reflective bead (gray sphere) (Fig. 6A). Furthermore, this technique relies upon the construction of a thermostable flow cell that can be heated to 65°C by way of a power source connected to an aluminum block on the outside of the flow cell. The DNA construct is delivered to the flow cell and is anchored by way of a biotin–streptavidin interaction. Processive thermostable enzymes, such as a DNA polymerase or helicase (shown in blue in Fig. 6A), are delivered to the flow cell, and assemble at the synthetic replication fork. Delivery of necessary reagants, such as ATP or dNTPs, to the flow cell initiate enzyme activity on single DNA constructs. The processive activity of an enzyme is visualized by the movement of the magnetic reflective bead tethered to the DNA construct. The movement of the beads is converted to trajectories that can be analyzed to determine rates and distances of processive enzyme activity (Fig. 6A). The use of this technique to examine the kinetics of DNA unwinding of MCM helicases from two archaeal species is discussed below. Figure 6. View largeDownload slide Analysis of MCM processivity using a high-temperature single-molecule bead tether assay. (A) A flow-cell and 5΄-biotin (red circle) labeled λ-DNA construct are used to carry out a high-temperature single-molecule bead tether assay to study DNA unwinding by archaeal MCM helicases. Bead trajectories are analyzed to obtain the rate and distance of DNA unwinding (Schermerhorn et al.2016). (B) The rate and distance of DNA unwinding from 50 events for three MCM helicases, MCM from M. thermautotrophicus (Mth MCM), and MCM2 and MCM3 from Thermococcus species 9°N (9°N MCM2 and 9°N MCM3, respectively), are displayed (Schermerhorn et al.2016) Make sure 2016 is connected to reference down below. Figure 6. View largeDownload slide Analysis of MCM processivity using a high-temperature single-molecule bead tether assay. (A) A flow-cell and 5΄-biotin (red circle) labeled λ-DNA construct are used to carry out a high-temperature single-molecule bead tether assay to study DNA unwinding by archaeal MCM helicases. Bead trajectories are analyzed to obtain the rate and distance of DNA unwinding (Schermerhorn et al.2016). (B) The rate and distance of DNA unwinding from 50 events for three MCM helicases, MCM from M. thermautotrophicus (Mth MCM), and MCM2 and MCM3 from Thermococcus species 9°N (9°N MCM2 and 9°N MCM3, respectively), are displayed (Schermerhorn et al.2016) Make sure 2016 is connected to reference down below. +) Using single-molecule analysis to obtain MCM helicase DNA unwinding parameters In eukarya, MCM is not active and needs to associate with GINS and Cdc45 to form the active CMG complex (described in detail above). Archaeal genomes encode for CMG components and thus it is possible that an archaeal CMG complex is needed for processive DNA unwinding. To test this hypothesis, the high-temperature single-molecule bead tether assay examined the DNA helicase unwinding kinetics of MCM helicases from two thermophilic archaea, Thermococcus sp. 9°N, which harbors multiple MCM copies, and Methanothermobacter thermautotrophicus, which harbors a single MCM (Fig. 5) (Schermerhorn et al.2016). This study revealed that the archaeal MCM helicases alone have robust DNA unwinding activity, unwinding stretches of 14 kb in length at a rate of up to 150 bp/s, and do not require additional factors for processivity. It is important to note that the single-molecule approach allows one to obtain kinetic parameters of processivity and unwinding rate of individual events, and therefore provides a distribution of rates and distances (Fig. 6B). This is in contrast to bulk kinetic methods which provide average rates. This study establishes a system to study long-range DNA synthesis and unwinding kinetics by DNA replication proteins from hyperthermophiles, and will aid in our understanding of the coordination between replisome components during processive DNA unwinding and synthesis. HOW CAN NEW TOOLS DRIVE ARCHAEAL DNA REPLICATION AND REPAIR RESEARCH? The implementation of new genetic, biophysical and molecular tools to study archaeal DNA replication and repair has enabled scientists to begin answering long-standing biological questions, including the necessity of an archaeal CMG complex, how Okazaki fragments are processed and if archaea possess DNA methylation. Importantly, we have just started to understand the complex nature of archaeal DNA replication and repair. In future work, we envision that tools described above will drive further understanding of these processes. Below we highlight a few remaining questions, and speculate how these tools can aid in answering them. +) How is archaeal DNA replication initiated in organisms lacking an origin of replication? The observation that the conserved origin of replication and the initiator proteins Orc1 and Cdc6 can be deleted without major effect on certain archaeal species growth changes our view of the origin of replication in archaea and its role in replication initiation (Kelman and Kelman 2018). Furthermore, such evidence suggests other mechanisms for replication initiation must be at play. One hypothesis is the emergence of dormant, non-conserved origins, which would facilitate bidirectional DNA synthesis. NGS technologies have routinely been used to track DNA replication initiation and termination sites in eukarya and bacteria, and would help uncover dormant origins in archaea (Marchal et al.2017). A second hypothesis is the utilization of HR to allow origin-independent replication initiation. Genetic knockouts of HR components in organisms other than H. volcanii and T. kodakarensis would help to facilitate our understanding and the necessity of HR during DNA replication, while NGS technologies could answer where HR initiates in the genome. It is possible that different archaeal organisms utilize different mechanisms to initiate DNA replication and the NGS approach may help to shed light on these processes. +) What regulates archaeal DNA replication? In bacteria and eukarya, the DNA replication process is highly regulated and many regulatory mechanisms have been identified (Skarstad and Katayama 2013; Parker, Botchan and Berger 2017). To date, only a single archaeal replication factor TIP (described above) has been identified as a potential replication regulator. The use of in vivo protein tagging, pull-downs and genetic knockouts can aid in the discovery of additional factors that regulate the replication process. +) What DNA repair mechanisms protect extremophiles? Due to the extreme environments in which many archaea thrive, and the DNA-damaging nature of these environments, robust mechanisms must exist to maintain the genome. We currently lack a complete understanding of what DNA repair pathways are present in archaea. Genetic knockouts provide an avenue to uncover the role of potential repair proteins, and to assess if they are essential, while in vivo protein tagging may help elucidate repair complexes and pathways. Furthermore, CE analysis can be used to study the substrate specificities and activities of individual DNA repair enzymes, as well as complexes. +) What mechanisms for lesion bypass exist in archaea? All eukaryotic, bacterial and crenarchaeal organisms possess family Y DNA lesion bypass polymerases (PolY), whose role is to perform DNA replication across bulky DNA lesions and prevent stalled replication (Broyde et al.2008). Euryarchaeal organisms lack PolY, and therefore how these organisms handle bulky lesions is a mystery. CE can be utilized to examine the ability of PolB and PolD to synthesize through bulky lesions, and assess how other replication factors may assist in the process. +) How is leading and lagging strand DNA synthesis coordinated? There is evidence to suggest that leading and lagging strand DNA synthesis is well coordinated, allowing the replisome to move along the DNA at a defined rate. Furthermore, coordination between DNA unwinding by the helicase and DNA synthesis by the polymerase must also occur to prevent exposure of ssDNA. How, and if, these events are coordinated in archaeal DNA replication is still unknown. The high-temperature single-molecule assay provides a system to enable the examination of these events, and would help to uncover which factors are needed to couple DNA unwinding and synthesis, and leading and lagging strand replication. 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FEMS Microbiology ReviewsOxford University Press

Published: Apr 18, 2018

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