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Critical Reviews in Biochemistry and Molecular Biology

ISSN: 1040-9238 (Print) 1549-7798 (Online) Journal homepage: http://www.tandfonline.com/loi/ibmg20

Mechanisms and regulation of DNA replicationinitiation in eukaryotes

Matthew W. Parker, Michael R. Botchan & James M. Berger

To cite this article: Matthew W. Parker, Michael R. Botchan & James M. Berger (2017)Mechanisms and regulation of DNA replication initiation in eukaryotes, Critical Reviews inBiochemistry and Molecular Biology, 52:2, 107-144, DOI: 10.1080/10409238.2016.1274717

To link to this article: https://doi.org/10.1080/10409238.2016.1274717

Published online: 17 Jan 2017.

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Mechanisms and regulation of DNA replication initiation in eukaryotes

Matthew W. Parkera, Michael R. Botchanb and James M. Bergera

aDepartment of Biophysics and Biophysical Chemistry, Johns Hopkins University School of Medicine, Baltimore, MD, USA; bDepartmentof Molecular and Cell Biology, University of California Berkeley, Berkeley, CA, USA

ABSTRACTCellular DNA replication is initiated through the action of multiprotein complexes that recognizereplication start sites in the chromosome (termed origins) and facilitate duplex DNA meltingwithin these regions. In a typical cell cycle, initiation occurs only once per origin and each roundof replication is tightly coupled to cell division. To avoid aberrant origin firing and re-replication,eukaryotes tightly regulate two events in the initiation process: loading of the replicative helicase,MCM2-7, onto chromatin by the origin recognition complex (ORC), and subsequent activation ofthe helicase by its incorporation into a complex known as the CMG. Recent work has begun toreveal the details of an orchestrated and sequential exchange of initiation factors on DNA thatgive rise to a replication-competent complex, the replisome. Here, we review the molecularmechanisms that underpin eukaryotic DNA replication initiation – from selecting replication startsites to replicative helicase loading and activation – and describe how these events are often dis-tinctly regulated across different eukaryotic model organisms.

ARTICLE HISTORYReceived 17 October 2016Revised 14 December 2016Accepted 16 December 2016

KEYWORDSDNA replication; ORC; Cdc6;initiator; Cdt1; MCM2-7;CMG; helicase

Introduction

The success of biological organisms depends on thefaithful transmission of genetic information from parentto progeny. All life-forms store their genetic content inthe form of nucleic acids, and the replication and dis-semination of this information forms the fundamentalbasis of inheritance. In cells, the process of replicationinvolves two primary tasks: (1) the separation of duplexDNA into two single-stranded templates and (2) semi-conservative replication of each strand. These eventsare coupled with cell division to produce progenywith essentially identical copies of the parent’s geneticinformation. Through all cellular lineages, a conserveddivision of labor has been applied to the process ofDNA replication, such that separable tasks (e.g. startsite selection, duplex unwinding, DNA synthesis) areallocated to different, albeit sometimes overlapping,factors. Although this basic framework is conservedthroughout the bacterial, archaeal, and eukaryotickingdoms, there has been significant evolutionary diver-sification of the molecules that complete each task, tothe point where it is now clear that aspects of the repli-cative machinery emerged twice, independently, duringcellular evolution (Edgell & Doolittle, 1997; Leipe et al.,1999). Replication initiation in eukaryl species has

become particularly elaborated by disparate forms ofregulation to meet the specific demands of multicellu-larity, development and large genome size.

Replication is started by a trans-acting “initiator” fac-tor that directs, in both space and time, loading of thereplicative machinery onto particular genomic lociknown as origins. In general, the number of originsscales with genome size, thereby ensuring that chromo-some duplication can be carried out on a physiologic-ally manageable timescale (Gilbert, 2004). Bacteria, aswell as certain archaea, frequently initiate replicationwith a single chromosomal start site (Costa et al., 2013;Wu et al., 2014b). Conversely, some archaeal chromo-somes possess multiple origins (Wu et al., 2014b), as doall eukaryotic genomes (the 12 Mbp Saccharomycescerevisiae genome contains around 400 origins and the3 Gbp human genome �30,000–50,000 origins(Leonard & Mechali, 2013)). The large number of originsand the need to coordinate initiation across these sitesrepresents a fundamental challenge to DNA replicationin eukaryotes; other factors, such as the use of multiplelinear chromosomes, as opposed to a single circularchromosome, add additional replicative complexity.Replication is particularly problematic in multicellulareukaryotes, where the process of development can alter

CONTACT James M. Berger jmberger@jhmi.edu Department of Biophysics and Biophysical Chemistry, Johns Hopkins University School of Medicine,Baltimore, MD, USA; Michael R. Botchan mbotchan@berkeley.edu Department of Molecular and Cell Biology, University of California Berkeley,Berkeley, CA, USA� 2017 Informa UK Limited, trading as Taylor & Francis Group

CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY, 2017VOL. 52, NO. 2, 107–144http://dx.doi.org/10.1080/10409238.2016.1274717

replication timing and frequency, and in the context ofcellular differentiation, which changes the chromatinlandscape and requires the coordinated transmission ofepigenetic marks. Despite these and other challenges,more than 15 trillion repetitive rounds of DNA replica-tion and cell division are successfully executed on thedevelopmental path of a fertilized human embryo tothe adult human body (Bianconi et al., 2013). Althoughour understanding of the replicative process is far fromcomplete, we are beginning to understand how eukar-yotes utilize a variety of sequential and redundant regu-latory mechanisms to achieve this biological feat.

The eukaryotic replisome is built from the regulatedand stepwise assembly of multiple intermediary replica-tion factor complexes. In S. cerevisiae, 42 individual pro-teins are sufficient to fully reconstitute DNA replicationin vitro, and since many of these proteins functionwithin large macromolecular assemblies, fewer than 15pre-assembled replication factors are required (Yeeleset al., 2015). In short, replication initiation entails foursteps (Figure 1): (1) demarcation of start sites by theOrigin Recognition Complex (ORC) and the Cdc6 heli-case-loader; (2) reiterative loading of an inactive form ofthe replicative helicase, MCM2–7, by ORC·Cdc6 and theCdt1 chaperone to form the pre-replication complex(pre-RC); (3) helicase activation by the formation of theCdc45·MCM2–7·GINS (CMG) complex (the pre-initiationcomplex, pre-IC) and (4) generation of a bidirectionalreplication fork that depends on prior origin melting by

the MCM2–7 complex and on the tethering of DNA pol-ymerases and additional accessory factors to the repli-cative helicase. Here, we review the molecularmechanisms underpinning eukaryotic replication initi-ation, from origin specification to helicase activation.This review, although focused on eukaryotic mecha-nisms, will, as needed, refer studies of the archaeal sys-tem to fill critical gaps in knowledge. Given theextensive number of publications in the field, we apolo-gize to those colleagues whose work is not referenceddue to space limitations.

Origins of replication

Origins of replication are chromosomal regions thatrecruit replication initiators for facilitating assembly ofthe replication machinery (Francois Jacob, 1963). Thedefining features of eukaryotic origins are complicatedand continuously evolving; for more thorough coveragewe refer the reader to a number of excellent reviews onthe topic (Creager et al., 2015; Leonard & Mechali, 2013;MacAlpine & Bell, 2005). Here, we briefly discuss themost salient features of origins in S. cerevisiae, S. pombeand metazoans, with a particular focus on details pertin-ent to later topics of discussion.

S. cerevisiae origins of replication

Saccharomyces cerevisiae origins of replication were ini-tially identified as chromosomal regions capable of

Figure 1. Mechanistic outline of DNA replication initiation in eukarya. During the G1 phase of the cell cycle, an origin-boundORC·Cdc6 complex together with Cdt1 facilitates the sequential recruitment and loading of two MCM2–7 complexes into a stabledouble hexamer that encircles duplex DNA (pre-RC). At the onset of S phase, the helicase is activated, leading to origin unwind-ing. The recruitment of other initiation factors (Cdc45 and GINS, the pre-IC) and double-hexamer dissolution activate the helicaseto drive fork progression as a single-stranded DNA-bound Cdc45·MCM2–7·GINS (CMG) complex. A color version of this figure isavailable at www.tandfonline.com/ibmg.

108 M. W. PARKER ET AL.

conferring replicative properties to exogenous plas-mids (Stinchcomb et al., 1979). These AT-rich, autono-mous replication sequences (ARSs) function asreplication start sites (Brewer & Fangman, 1987;Huberman et al., 1987,1988) and serve to recruitthe eukaryotic initiator, ORC (Bell & Stillman, 1992).The ARS contains a number of essential elements, themost important of which is the 11-basepair “A ele-ment”, which constitutes the ARS consensus sequence(ACS) and represents the primary site of initiator bind-ing (Marahrens & Stillman, 1992; Rao et al., 1994; Rao& Stillman, 1995; Theis & Newlon, 1994). Notably,among the different model eukaryotic systems usedfor studying replication, only S. cerevisiae appears toutilize a specific consensus sequence (Figure 2).

Although there are over 12,000 ACS-like sequencesin the yeast genome, only about 400 facilitate replica-tion initiation (Nieduszynski et al., 2006; Wyrick et al.,

2001; Xu et al., 2006). This low usage of possibleS. cerevisiae origins (<5%) derives in part from an add-itional level of origin specification that is imposed bythe local chromatin structure. Saccharomyces cerevisiaeorigins, like those of other eukaryotes, are maintainedas nucleosome-free regions (NFRs) (Berbenetz et al.,2010; Eaton et al., 2010). However, surrounding an ARS,ORC-dependent nucleosome phasing directly affectsthe efficiency of origin usage (Berbenetz et al., 2010;Lipford & Bell, 2001; Simpson, 1990; Thoma et al., 1984),and the eviction of ORC results in nucleosomeencroachment into ARS regions (Eaton et al., 2010;Thoma et al., 1984). Mechanistically, NFRs allow ORCaccess to DNA, while phased nucleosomes provide add-itional favorable sites for binding ORC (Hizume et al.,2013; Muller et al., 2010). Thus, S. cerevisiae origins aredefined by ORC binding to both an ACS and specificchromatin features (Hoggard et al., 2013; Leonard &Mechali, 2013).

S. pombe origins of replication

Genome-wide studies demonstrate that the S. pombegenome contains around 400 origin sequences that aregenerally nucleosome-free (Givens et al., 2012; Xu et al.,2012), AT-rich, and around 1 kilobase long (Dai et al.,2005; Heichinger et al., 2006; Segurado et al., 2003).These findings are consistent with biochemical studiesshowing that origin usage in fission yeast depends onclustered stretches of adenine and thymine (Clyne &Kelly, 1995; Dai et al., 2005; Kim et al., 2001; Okunoet al., 1999). Origin selection in S. pombe is facilitated bya species-specific insertion in the Orc4 subunit of ORCthat encodes a DNA-binding element that specificallyrecognizes the minor groove of AT-rich sequences(Figure 2) (Chuang & Kelly, 1999; Moon et al., 1999) .

Metazoan origins of replication

Unlike budding yeast ORC, which shows a degree ofsequence-specificity, metazoan ORC binds DNA promis-cuously (Remus et al., 2004; Vashee et al., 2003). Thisbehavior is consistent with the observation that meta-zoan replication initiates from diverse sequences(Heinzel et al., 1991; Hyrien & Mechali, 1993; Mechali &Kearsey, 1984). Despite origin sequence variability, thegenome-wide analysis of replication start sites hasrevealed some common patterns in metazoan origins.As with budding and fission yeast, metazoan ORC bindsto NFRs in the genome (Eaton et al., 2011; Karnani et al.,2010; MacAlpine et al., 2010), which in turn favorablycontributes to assembly of the replication machinery(Lubelsky et al., 2011). Interestingly, G-rich sequences

Figure 2. Molecular details of eukaryotic origins and mecha-nisms of ORC binding. (gray) Saccharomyces cerevisiae originsare distinctive among eukaryotes for conforming to a consen-sus sequence, the ACS. ScORC can bind the ACS directly andspecifically, although interactions between the Orc1-BAHdomain and nucleosomes can also modulate ORC origin selec-tion. (PURPLE) Although they do not possess a strict consen-sus sequence, S. pombe origins are AT-rich. SpORC specificallybinds such sites using a domain insertion unique to SpOrc4that encodes a DNA-binding AT-hook motif. (green) MetazoanORC can be targeted to chromosomes through a variety ofmechanisms, including the Orc1 BAH domain, the DNA-bind-ing TFIIB domain of Orc6, and through interactions with chro-matin-associated factors. A majority of metazoan origins arealso predicted to contain G-quadruplex secondary structureelements, but how this feature affects ORC binding is cur-rently unclear. A color version of this figure is available atwww.tandfonline.com/ibmg.

CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY 109

and CpG islands are highly enriched in metazoan ori-gins (Cadoret et al., 2008; Delgado et al., 1998; Prioleau,2009; Sequeira-Mendes et al., 2009) and have been pro-posed to serve two purposes: NFR maintenance(Huppert & Balasubramanian, 2007; Fenouil et al., 2012;Wong & Huppert, 2009) and the favoring of G-quadra-plex formation (Cayrou et al., 2011, 2012, 2015; Valtonet al., 2014). Preliminary analyses suggest that ORC maypreferentially associate with G-rich elements (Hoshinaet al., 2013; Zellner et al., 2007), indicating that a con-served structural feature in DNA, rather than a specificconsensus sequence, may aid ORC binding in metazo-ans. As in S. cerevisiae, metazoan origin selection byORC is further fine-tuned by direct interactions withnucleosomes and chromatin-associated factors. Forexample, the N-terminal bromo-adjacent homology(BAH) domain in Orc1 directly interacts with histones todirect origin usage (Figure 2) (Kuo et al., 2012; Mulleret al., 2010; Noguchi et al., 2006).

Although ORC can be targeted to specific genomicloci, origins cannot be strictly defined by the position ofORC binding on chromatin. Indeed, once loaded ontoDNA by ORC, the MCM2–7 helicase (which eventuallynucleates replisome assembly following duplex melting)appears to lack positional restraints and is free to eitherdiffuse away from ORC (Evrin et al., 2009; Remus et al.,2009) or be forcibly displaced by other chromatin-local-ized cellular processes (such as the transcriptionalmachinery) (Edwards et al., 2002; Gros et al., 2015;Powell et al., 2015; Ritzi et al., 1998). Thus, origins ineukaryotes must be defined flexibly, as the site of initi-ator binding does not always reflect the site of replica-tion initiation.

Overall, both yeast and metazoa contain conservedsequence elements at origins that are known or pro-posed to guide ORC binding. In yeast, these elementsare encoded within the DNA primary sequence and inmetazoa potentially by a propensity to form distinctivesecondary structures. Chromatin context plays an add-itional critical, but poorly understood role in originusage in all eukaryotes. Understanding how these cis-and trans-acting origin elements interface is an import-ant and active area of future research.

The origin recognition complex (ORC)

Eukaryotic origins direct the recruitment of the originrecognition complex (ORC), a conserved heterohexa-meric protein assembly identified for its ability to specif-ically recognize the double-stranded form of the yeastACS (Bell & Stillman, 1992). Upon recruitment tochromosomal replication start sites, ORC binds an add-itional factor, Cdc6, as a necessary prerequisite to

helicase loading. Despite its centrality to ORC function,the mechanism of DNA binding by the initiator haslong remained ambiguous. However, structural studiesfrom archaea and eukaryotes have revealed a conservedmechanism for the association of ORC with DNA thatinforms not only our understanding of how theORC·Cdc6 initiator stably binds replication start sites,but also how ORC mediates downstream helicase load-ing events (Bleichert et al., 2015; Dueber et al., 2007;Gaudier et al., 2007; Sun et al., 2013).

The origin recognition complex and Cdc6

ORC was first identified by fractionation of ARS-bindingproteins in budding yeast (Bell & Stillman, 1992).Although many other ARS-binding factors had beenidentified previously (Buchman et al., 1988; Diffley &Stillman, 1988; Jazwinski & Edelman, 1982; Shore et al.,1987; Sweder et al., 1988), ORC proved uniquely able tobind the ACS (Diffley & Cocker, 1992; Li & Herskowitz,1993; Marahrens & Stillman, 1992,), and temperaturesensitive mutants exhibited cell-cycle arrest at a stageconsistent with a role in the early aspects of DNA repli-cation (Bell et al., 1993; Foss et al., 1993; Micklem et al.,1993). Following the discovery of ORC in budding yeast,the broad eukaryotic conservation of ORC was demon-strated with the identification of orthologs in S. pombe(Grallert & Nurse, 1996; Muzi-Falconi & Kelly, 1995),Drosophila melanogaster (Gossen et al., 1995; Landiset al., 1997), Xenopus laevis (Carpenter et al., 1996;Romanowski et al., 1996b) and humans (Gavin et al.,1995). The mechanism of ORC function has now beeninvestigated across multiple model organisms, revealingthat the core components, subunit organization andfunction of ORC are broadly conserved (although notuniversally, particularly in protozoa (El-Sayed et al.,2005)). Despite this conservation, there do exist certainspecies-specific alterations that appear to have gener-ated notable functional differences.

Saccharomyces cerevisiae ORC is a roughly 400 kDaassembly composed of six proteins (Orc1–6) named indescending order of molecular weight (Bell et al., 1993).Orc1–5 exhibit relatively good conservation across spe-cies and are members of the ATPases associated withdiverse cellular activities (AAAþ) superfamily of pro-teins; Orc1–5 also contain a C-terminal winged-helix(WH) domain (Bell et al., 1995; Liu et al., 2000; Loo et al.,1995; Muzi-Falconi & Kelly, 1995). Overall, the Orc1–5AAAþ and WH domains, which account for a bit overhalf the total mass of ORC, display an average of�45–50% similarity and 25–30% identity between yeastand human orthologs (Speck et al., 2005; Tugal et al.,1998). A majority of AAAþ family members bind and

110 M. W. PARKER ET AL.

hydrolyze ATP through the formation of a composite,bipartite active site that is generated between neigh-boring AAAþ protomers upon subunit oligomerization(Guenther et al., 1997; Lenzen et al., 1998; Putnam et al.,2001). AAAþ proteins, like the broad P-loop family ofNTPases to which they belong (Iyer et al., 2004), containthe so-called “Walker A” and “Walker B” signaturesequence motifs that contribute to nucleotide bindingand hydrolysis, respectively. A third motif, generally crit-ical to AAAþ ATPase function, is the arginine finger,which is presented in trans to the neighboring subunitto reconstitute an active ATPase site (Neuwald et al.,1999; Zhang et al., 2000). While many of the structuralelements important for ATP binding are conserved inOrc1, Orc4 and Orc5, only Orc1 has been found to pos-sess ATPase activity (Bleichert et al., 2015; Klemm et al.,1997; Makise et al., 2003; Ranjan & Gossen, 2006;Siddiqui & Stillman, 2007); Orc2 and Orc3 retain degen-erate AAAþ scaffolds that lack functional active sitemotifs altogether (Bleichert et al., 2015; Clarey et al.,2006; Speck et al., 2005). Intriguingly, although manyAAAþ family members function as toroidal hexamericassemblies (reviewed in (Hanson & Whiteheart, 2005)),ORC retains only five proteins with AAAþ domains.Orc6 lacks a AAAþ domain (Balasov et al., 2007;Bleichert et al., 2013; Chesnokov et al., 2003) and its pri-mary sequence is only weakly conserved between yeastand human, making it the least conserved ORC subunit(Dhar & Dutta, 2000). Nonetheless, certain elements ofOrc6 are conserved across species, including an N-ter-minal TFIIB-like domain and a short conserved region atthe extreme C-terminus of the protein (Bleichert et al.,2013; Liu et al., 2011).

Cdc6 forms a complex with ORC at origins and isrequired for initiator function. Cdc6 was first identifiedin S. cerevisiae mutant screens (Hartwell, 1976) and waslater found to have a role in replication initiation(Palmer et al., 1990), with functional requirements priorto S-phase (Hogan & Koshland, 1992; Kelly et al., 1993;Zwerschke et al., 1994). A genetic interaction betweenORC and Cdc6 suggested a coordinated activity forthese factors (Liang et al., 1995), with supporting datademonstrating an ORC-dependent recruitment of Cdc6to origins that relies on a direct interaction between theproteins (Cocker et al., 1996; Coleman et al., 1996;Grallert & Nurse, 1996; Kong et al., 2003; Leatherwoodet al., 1996; Liang et al., 1995; Santocanale & Diffley,1996). Importantly, analysis of the Cdc6 primarysequence reveals conserved nucleotide binding andhydrolysis motifs (Lisziewicz et al., 1988; Zhou et al.,1989; Zwerschke et al., 1994), as well as close homologyto ORC’s AAAþ subunits, particularly Orc1 (Bell et al.,1995; Quintana et al., 1997). Cdc6 associates with

chromatin-bound ORC in an ATP-dependent manner(Coster et al., 2014; Evrin et al., 2013; Kang et al., 2014;Kneissl et al., 2003; Perkins & Diffley, 1998; Ticau et al.,2015), an interaction that activates Cdc6’s ATPase activ-ity and is consistent with the reconstitution of canonicalAAAþ interactions (Randell et al., 2006; Speck &Stillman, 2007). Thus, Cdc6 recruitment provides a sixthAAAþ subunit to the initiator complex overall.

Structure of ORC and ORC·Cdc6

Orc/Cdc6 homologs have been identified in all eukary-otic and archaeal species analyzed (Aves et al., 2012). Inmany archaea, the initiation factors are geneticallystreamlined such that certain species possess only a sin-gle Orc/Cdc6 gene (Barry & Bell, 2006) and the relativesimplicity of the archaeal system has been exploited tohelp understand the structure and function of eukary-otic Orc homologs. A conserved three-domain architec-ture is observed for Orc and Cdc6 homologs, with twodomains at the N-terminus forming the central AAAþmodule and a third at the C-terminus comprising aloosely-tethered WH domain (Figure 3(A)) (Bleichertet al., 2015; Dueber et al., 2007; Liu et al., 2000; Gaudieret al., 2007; Singleton et al., 2004). The structure of arch-aeal Orc in complex with DNA has shown that both theWH and AAAþ domains contact DNA (Dueber et al.,2007; Gaudier et al., 2007), and biochemical studieshave demonstrated that contacts mediated by theAAAþ domain contribute to origin specificity (Dueberet al., 2011). Surprisingly, structures of Orc/DNA com-plexes have not revealed evidence for the formation ofan ATPase-competent Orc dimer (Dueber et al., 2007;Gaudier et al., 2007), despite the existence of a con-served arginine finger in these proteins, which wouldotherwise suggest that oligomerization might occur.The mechanism that promotes formation of a catalytic-ally active ATPase in the archaeal system is unclear.

Saccharomyces cerevisiae Cdc6 forms a stable com-plex with ORC in the presence of DNA and ATP(Mizushima et al., 2000; Seki & Diffley, 2000; Speck et al.,2005; Randell et al., 2006; Wang et al., 1999). By using anon-hydrolyzable ATP analog, a stable Cdc6·ORC com-plex can also be trapped in the absence of DNA (Specket al., 2005). Low-resolution 3D electron microscopyreconstructions of eukaryotic ORC have revealed anelongated, crescent-shaped particle that, in the pres-ence of Cdc6 and ATPcS, transforms into a closed ringwith a large central cavity (Clarey et al., 2006, 2008;Speck et al., 2005; Sun et al., 2012). Structural investiga-tion of D. melanogaster ORC has demonstrated a sub-unit order of Orc1!Orc4!Orc5!Orc3!Orc2 aroundthe ORC ring (Bleichert et al., 2015), with a physical gap

CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY 111

between the terminal Orc subunits that accommodatesCdc6 and is consistent with the direct interactionobserved between Cdc6 and Orc1 (Sun et al., 2012,2013; Wang et al., 1999). The C-terminus of Orc6 hasbeen found to tether this subunit to a conserveddomain insertion within Orc3 (Bleichert et al., 2013,2015); S. cerevisiae Orc6 also binds Orc2 (Chen et al.,2008; Sun et al., 2012), possibly through a region that isspecific to fungal homologs (Bleichert et al., 2013). Theadjoining nature of Orc1 and Orc4 within the ternarycomplex is consistent with the known formation of ajoint ATPase site between the two subunits (Bowerset al., 2004; Chesnokov et al., 2001; Giordano-Coltartet al., 2005; Klemm et al., 1997).

The ORC·Cdc6 assembly represents the functional ini-tiator complex at origins. Interestingly, ORC itself exhib-its differing levels of stability across species, suggestingthat, in certain cases, ORC subcomplexes may besequentially recruited. Indeed, unlike D. melanogasterand S. cerevisiae ORC, which form stable heterohex-amers (Bell & Stillman, 1992; Chesnokov et al., 1999),human and X. laevis ORC appear to have alternativecore subcomplexes. Human Orc1 and Orc6 loosely asso-ciate with an Orc2–5 core (Dhar et al., 2001; Siddiqui &Stillman, 2007; Vashee et al., 2001), whereas in X. laevis,the Orc6 subunit is labile (Gillespie et al., 2001). Theweak interactions of certain Orc subunits likely play an

important role in regulation. For example, vertebrateOrc1 is selectively released from chromatin after initi-ation (Li et al., 2004; Natale et al., 2000; Rowles et al.,1999), which helps to prevent re-initiation and providesa means to alter origin usage in a developmental- ordifferentiation-dependent fashion (Li & DePamphilis,2002). The ability of either Orc1 or Orc6 to dissociatefrom ORC without disrupting the remaining core com-plex is consistent with the terminal position of Orc1within the core ring and the peripheral binding site ofOrc6; one exception to this trend occurs in S. pombe,whereby an ORC pentamer can be purified that lacksOrc4, an internal component of the ORC ring (Kong &DePamphilis, 2001; Moon et al., 1999). Whether S.pombe ORC retains additional stabilizing elements thatcan compensate for the absence of Orc4 in such instan-ces is not known.

A recent atomic-resolution structure of D. mela-nogaster ORC (Bleichert et al., 2015) has helped to clarifyboth our understanding of ORC organization and mech-anistic models for origin engagement and helicase load-ing. ORC adopts a two-tiered, notched ring architecturein which the WH domains of Orc1–5 sit atop a layer ofAAAþ subunits (Figure 3(B)). Interestingly, the arrange-ment of AAAþ and WH domain contacts is domainswapped in ORC, such that the WH domain of Orc1rests on the AAAþ region of Orc4, the WH domain of

Figure 3. ORC architecture. (A) Cdc6/Orc homologs are characterized by three domains, two of which form the AAAþ module(green) and a third that encodes a winged-helix (WH) domain (gray). Bound nucleotide is shown as sticks (PDB ¼1FNN). (B) TheD. melanogaster ORC heterohexamer is a crescent-shaped molecule with the AAAþ (green surface) and WH (gray cartoon)domains forming a domain-swapped arrangement. Orc6 is bound by a domain insertion in the AAAþ domain of Orc3. Althoughthe Orc1/Orc4 active site is required for activity, in the D. melanogaster structure Orc1 is disengaged from Orc4 and positionedabove the plane of the AAAþ ring (PDB ¼4XGC). A color version of this figure is available at www.tandfonline.com/ibmg.

112 M. W. PARKER ET AL.

Orc4 sits on the AAAþ region of Orc5, and so forth. Theopen-ended, pentameric AAAþ core of ORC in principleprovides an opportunity for the formation of fourbipartite AAAþ interfaces. Within the D. melanogasterORC structure, however, only one of the observed sub-unit interfaces evinces a typical AAAþ active-site config-uration (Orc4/Orc5). Two others (Orc5/3 and Orc3/2)approximate the correct protomer alignment but failto reconstitute a canonical active site; in the case ofOrc3/2, the two subunits lack the amino acids necessaryto bind ATP, as predicted. Of the remaining possibleAAAþ interactions, Orc1 was unexpectedly found to berotated more than 90 degrees out-of-plane from Orc4,rendering this catalytic center inoperative (Figure 3(B)).This finding was surprising, as Orc1 alone is capable ofboth binding and hydrolyzing ATP, using an argininefinger donated by Orc4 (Chesnokov et al., 2001; Klemmet al., 1997). Given that the D. melanogaster Orc1 con-formation seen crystallographically is also seen in 3Delectron microscopy reconstructions of ORC (Bleichertet al., 2013, 2015), this observation suggests that meta-zoan ORC can transition between at least two confor-mations, an autoinhibited and active conformation. Inthe future, it will be important to understand how thecell regulates the equilibria of ORC between thesestates, as well as whether this conformational transitionis preserved in other ORC homologs.

A two-state model for ORC origin recognition

Despite the fundamental role of ORC in origin selectionand recognition, the mechanism by which ORC associ-ates with DNA has remained highly enigmatic. The pres-ence of numerous interspecies peculiarities, such asdivergent origin features, species-specific DNA-bindingelements, and the effect of chromatin-bound trans-act-ing factors, have all challenged our understanding ofhow ORC is recruited to and stably binds origins.However, an analysis of ORC behavior across speciessuggests that in all cases, the origin-binding propertiesof ORC can be interpreted within a model containing atleast two states: a transient ORC recruitment event thatis mediated through diverse and sometimes species-specific interactions, and a second, mechanistically con-served step that positions ORC for productive originengagement and that leads to stable Cdc6 association.We will first discuss the conserved mechanism by whichORC stably associates with origins and then detail theinteractions that facilitate ORC recruitment.

Structural studies of origin-bound archaeal Orc haverevealed a coordinated role for both the AAAþ and WHdomains in DNA binding (Dueber et al., 2007; Gaudieret al., 2007). The WH domains show extensive contacts

with origin DNA using a canonical helix-turn-helix (HTH)and b-hairpin wing interface, and a near compete lossof DNA binding by archaeal Orc is observed upon muta-tion or deletion of this region (Dueber et al., 2011;Singleton et al., 2004). Interestingly, the WH-DNA inter-action positions the AAAþ domain in an orientationwhere a characteristic a-helical insertion within the initi-ator/helicase loader subgroup of AAAþ proteins, theinitiator specific motif (ISM), contacts DNA. Given thedifferent subunit compositions and oligomeric statesobserved between archaeal (monomer) and eukaryoticinitiators (heterohexamer) it was initially unclear towhat extent the mechanism of archaeal origin bindingwould be conserved. However, superposing the struc-ture of DNA-bound archaeal Orc onto the D. mela-nogaster ORC crystal structure reveals nearly perfect co-axial positioning of the DNA within the ORC centralchannel (Bleichert et al., 2015). Notably, the eukaryoticISMs and the wings of the WH domains each form acontiguous DNA-binding surface that lines the ORC cen-tral channel, and are thus positioned to engage DNA ina manner similar to that of archaeal Orc (although notidentically, as the HTH motif of ORC’s WH domains areburied between inter-subunit contacts in the complex(Bleichert et al., 2015)). This modeling also accounts foran unresolved region of density in an early 3D electronmicroscopy reconstruction of an ORC·Cdc6·DNA com-plex (Sun et al., 2012). Overall, these data indicate thatarchaeal and eukaryotic ORC engage DNA by a gener-ally conserved mechanism.

The insights gleaned from the available structuralstudies suggest a conserved mechanism for stable ori-gin association by eukaryotic ORC. In this model, duplexDNA is loaded laterally into the ORC central channelthrough the discontinuity between Orc1 and Orc2(Speck et al., 2005). This gap is then blocked off by thesubsequent recruitment of Cdc6, generating a stable,closed-ring conformation that encircles origin DNA(Bleichert et al., 2015). Domain swapping between theWH domain of Orc2 and the AAAþ region of Cdc6 (andbetween the WH domain of Cdc6 and the AAAþ elem-ent of Orc1) are predicted to form, stabilizing the com-plex (Bleichert et al., 2015). The ability of the ORC·Cdc6complex to encircle DNA would be predicted to under-pin ORC’s observed persistence at origins (Duzdevichet al., 2015; Speck et al., 2005) and is consistent with theobservation that both yeast and metazoan Cdc6 canstabilize ORC on DNA (Harvey & Newport, 2003;Houchens et al., 2008).

Interestingly, an additional DNA-binding element hasbeen identified between the N-terminal BAH and AAAþdomain of S. cerevisiae Orc1 that is essential for ARSbinding (Kawakami et al., 2015). While the related

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residues are largely conserved in metazoan Orc1, theyare positioned outside of the ORC central pore and thusit is unclear how this interaction is coordinated withthe encirclement mechanism described here, orwhether it contributes only to the initial recruitmentof ORC to DNA.

Diverse mechanisms for origin recruitment of ORC

Although the encirclement model accounts for stableorigin binding, eukaryotic ORC has been observed tointeract with DNA in a mode that exhibits fast kinetics(Duzdevich et al., 2015; Harvey & Newport, 2003;Houchens et al., 2008; Remus et al., 2004) and, in S.cerevisiae, allows for a one-dimensional linear search forbona fide origin sites (Duzdevich et al., 2015). This initialrecruitment of ORC to DNA likely functions as animportant intermediate on the path towards stableorigin binding. Interestingly, substantial mechanisticplasticity appears to have been introduced to therecruitment step, co-evolving in certain cases with spe-cies-specific features.

In S. pombe ORC, the preference for AT-rich originscorrelates with a unique domain insertion in Orc4 com-prising nine AT-hook motifs (Chuang & Kelly, 1999;Moon et al., 1999), a DNA-binding element that facili-tates interactions with the minor groove of AT-rich DNAsequences (Aravind & Landsman, 1998). Notably, theAT-hook motif is absent in S. cerevisiae and metazoanOrc4. Unlike S. cerevisiae ORC, which utilizes all AAAþdomain-containing Orc subunits for DNA binding (Lee &Bell, 1997), S. pombe Orc4 is necessary and sufficient forinitial origin engagement (Chuang & Kelly, 1999;Gaczynska et al., 2004; Kong & DePamphilis, 2001; Moonet al., 1999), and the AT-hook motifs of Orc4 are add-itionally required for viability (Chuang et al., 2002).Interestingly, S. pombe ORC shows a biphasic mechan-ism of DNA binding, with an initial, salt-sensitive DNAbinding event that precedes the formation of a salt-sta-ble form, a state that in turn can be further stabilizedby the addition of Cdc6 (Houchens et al., 2008). Thesefindings suggest that S. pombe ORC is recruited to chro-mosomes by the Orc4 AT-hook motif, which then leadsto stable DNA association through a mechanism thatlikely involves the encirclement of duplex DNA.

Analogous to S. pombe Orc4, metazoan Orc6 hasbeen shown to have a distinct DNA binding activity.Despite the absence of an ATPase or WH domain, exclu-sion of Orc6 from D. melanogaster ORC results in theloss of ATP-dependent DNA binding, the same effectobserved for Orc1 Walker A (ATP binding) and Walker B(ATP hydrolysis) mutants (Chesnokov et al., 2001).Analysis of the D. melanogaster and human Orc6

N-terminus reveals structural homology with the DNA-binding domain of transcription factor TFIIB (Balasovet al., 2007; Chesnokov et al., 2003), and mutation of theOrc6 TFIIB domain abolishes DNA binding (Liu et al.,2011). Although the Orc6 TFIIB domain was initially con-sidered unique to metazoans, subsequent sequenceanalysis indicates that the domain is conserved in fun-gal Orc6, but lacks specific DNA-binding elements(Bleichert et al., 2013), a finding consistent with S. cerevi-siae Orc6 being dispensable for origin recognition invitro (Chen et al., 2007; Lee & Bell, 1997). Given theavailable data, it seems likely that metazoan ORC uti-lizes Orc6 to loosely tether the complex to DNA in afunctionally analogous manner as S. pombe Orc4, andthat this action aids with the initial ORC recruitmentevent that precedes stable origin association. Thismodel begs the question, however, of how ORC isrecruited to chromosomes in species where Orc6 is alabile subunit (Dhar et al., 2001; Gillespie et al., 2001;Vashee et al., 2001). One possible answer is that chro-matin-bound Orc6 may function as a recruitment plat-form for the core Orc1–5 subunits.

In addition to “hard-wired” DNA binding domains,eukaryotic ORC can also bind a plethora of chromatin-associated factors that provide additional means forrecruiting and regulating ORC’s association with origins(Chakraborty et al., 2011). Early studies recognized thatORC played dual roles in replication and transcriptionalregulation (Bell et al., 1993; Leatherwood & Vas, 2003;Loo et al., 1995; Palacios DeBeer et al., 2003), and anumber of transcriptional regulators have been shownto bind ORC, including HP1, E2F, HMGA1a and Sir1(Bosco et al., 2001; Fox et al., 1997; Gardner et al., 1999;Pak et al., 1997; Prasanth et al., 2004, 2010; Royzmanet al., 1999; Thomae et al., 2008; Triolo & Sternglanz,1996). Although chromatin accessibility and transcrip-tional programs are clearly regulated in an ORC-dependent manner (Bell et al., 1993; Bose et al., 2004;Chesnokov, 2007; Foss et al., 1993; Fox et al., 1995;Huang et al., 1998; Loo et al., 1995; Micklem et al., 1993;Pak et al., 1997; Shor et al., 2009), whether chromatinbound transcriptional regulators also direct ORC originusage is less clear. Certain cases have been investigatedin some detail; for example, ORC can be targeted tochromatin-bound HMGA1a, where it directs assembly ofreplication complexes (Thomae et al., 2008), while gen-ome-wide analysis of metazoan origins reveals ORCassociation with HP1 sites (Cayrou et al., 2011). Thus,transcriptional regulators represent an additional meansof tethering ORC to chromosomes and likely affect ORCfunction in replication.

Eukaryotic ORC is also directly recruited to histonesby the N-terminal Bromo-Adjacent Homology (BAH)

114 M. W. PARKER ET AL.

domain in Orc1 (Noguchi et al., 2006; Muller et al., 2010;Zhang et al., 2002). This element selectively bindsH4K20me2, an interaction that directs replication licens-ing (Kuo et al., 2012). Other histone modifications alsohave been found to direct origin usage (such asH4K20me1, H3K4me2/3, and H4 acetylation), but howthese effect ORC positioning or downstream licensingreactions remains unclear (Aggarwal & Calvi, 2004;Costas et al., 2011; Eaton et al., 2011; Knott et al., 2009;Liu et al., 2012b; Miotto & Struhl, 2010; Rice et al., 2002;Vogelauer et al., 2002; Tardat et al., 2010). A factorknown as ‘ORCA’ (for Origin Recognition ComplexAssociated) additionally has been reported to directlyrecruit human ORC to chromosomes, as well as tobind to other initiation factors that promotereplication licensing and S-phase progression(Shen et al., 2010, 2012). Conversely, the assembly ofthe replication machinery at telomeres has beenreported to be controlled by an interaction betweenORC and the TRF2 subunit of the shelterin complex(Atanasiu et al., 2006; Deng et al., 2009; Higa et al., 2016;Tatsumi et al., 2008).

Collectively, multiple lines of data demonstrate thatlocalizing ORC to DNA prior to productive originengagement (i.e. Cdc6-dependent DNA encircling) is acritical step in replication initiation, and that manydiverse recruitment mechanisms are sufficient todemarcate a replication start site. Consistent with thisidea is the demonstration that a fusion between ORCand the Gal4 DNA binding domain can result in the ini-tiation of DNA replication on a plasmid containing atandem array of Gal4 binding sites (Takeda et al., 2005).In light of the many mechanisms that can facilitate ORCrecruitment, an important future direction will be tounderstand how different recruitment pathwayscooperate or antagonize each other in specifying sitesof replisome assembly.

The minichromosome maintenance (MCM)complex

Following the active designation of an origin byORC, the complex next facilitates initiation at thesesites by loading the heterohexameric MCM2–7 heli-case onto DNA. Studies into MCM2–7 function haverevealed a surprisingly complex enzyme that appearscapable of harnessing ATP to promote both themelting of double-stranded DNA and translocation ofsingle-stranded DNA substrates. Although precisemechanisms have yet to be elaborated, a picture ofhow specific MCM2–7 elements couple ATP turnoverto DNA remodeling and movement is beginning toemerge.

Identification of the MCM2–7 complex

MCM proteins were originally identified in geneticscreens that aimed to uncover factors required for repli-cation and cell cycle progression in yeast (Chen et al.,1992; Hennessy et al., 1990; Maine et al., 1984; Moiret al., 1982; Yan et al., 1991). Homologs were subse-quently identified in D. melanogaster (Treisman et al.,1995), X. laevis (Madine et al., 1995) and in humans(Todorov et al., 1995), and like yeast MCMs, were shownto have an essential role in DNA replication.Investigation into MCM function converged with studiesby Laskey and Blow, who identified a replication licens-ing factor (RLF) that restricted replication to once percell cycle by nuclear exclusion until after nuclear enve-lope breakdown (Blow & Laskey, 1988; Blow, 1993).Indeed, MCM proteins, such as the RLF, showed redistri-bution from the cytosol to nucleus upon completion ofmitosis (Hennessy et al., 1990), and immunodepletion ofMCM proteins inhibited replication (Kubota et al., 1995;Madine et al., 1995). Importantly, a purified RLF fractionwas found to resolve into two separate factors, RLF-Mand RLF-B, with RLF-M containing multiple proteins thatcross-reacted with MCM antibodies (Chong et al., 1995).

Purification of RLF-M, as well as co-immunoprecipita-tion studies, revealed that MCM proteins reside withinlarge multimeric assemblies with other MCM2–7 familymembers (Burkhart et al., 1995; Chong et al., 1995, 1996;Kubota et al., 1995; Kimura et al., 1995; Madine et al.,1995; Musahl et al., 1995; Romanowski et al., 1996a).The predominant assembly in vivo is an MCM2–7 heter-ohexamer, although low levels of subassemblies havebeen reported to exist that may represent intermediates(Adachi et al., 1997; Ishimi, 1997; Lee & Hurwitz, 2000;Prokhorova & Blow, 2000; Su et al., 1996; Thommeset al., 1997). As with Orc1–5, MCM proteins are predi-cated upon a conserved AAAþ ATPase element (Iyeret al., 2004; Koonin, 1993). However, unlike Orc1–5,which show differing levels of active site conservation,the six subunits in the MCM2–7 complex each containthe complete set of catalytic residues expected to benecessary for supporting ATP hydrolysis.

MCM architecture

The basic architecture of an MCM protein is conservedacross archaea and eukaryotes. MCMs contain a centralAAAþ ATPase fold flanked by conserved N- and C-ter-minal domains (termed “NTD” and “CTD”, respectively).Structural analyses of MCMs have revealed three con-served subdomains within the NTD, allowing subdiv-ision of this element into the NTD-A, -B and -C regions(Figure 4(A)) (Bae et al., 2009; Brewster et al., 2008;

CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY 115

Fletcher et al., 2003; Fu et al., 2014; Li et al., 2015; Liuet al., 2008). NTD-C, the most highly conserved NTDsubdomain, possesses an oligonucleotide/oligosacchar-ide-binding (OB) fold. Within an MCM hexamer, interac-tions between adjacent NTD-C elements serve as theprimary interface for subunit assembly, with this inter-action alone sufficient to facilitate hexamerization ofthe NTD (Fletcher et al., 2003; Kasiviswanathan et al.,2004). Although NTD-A is the least conservedN-terminal subdomain, it forms a helical bundle withsimilarity to helix-turn-helix type DNA binding proteins(Costa et al., 2008). The NTD-B element comprises aZn-finger motif that, with the exception of MCM3, isuniversally conserved in MCM2–7 (Li et al., 2015), andthat facilitates higher order organization of single NTDhexamers into a conserved, head-to-head double hex-amer (Figure 4(B)) (Chong et al., 2000; Costa et al., 2014;Evrin et al., 2009; Fletcher et al., 2005; Kelman et al.,1999; Li et al., 2015; Remus et al., 2009; Shechter et al.,2000).

The AAAþ domain is the most highly conservedregion across MCM homologs. The MCM AAAþ fold

contains three unique insertions relative to prototypicalAAAþ proteins, and falls within the pre-sensor II (PSII)insert clade of the ATPase superfamily (Erzberger &Berger, 2006; Iyer et al., 2004). One such insertion is dis-tinctive in that it remodels a portion of the AAAþ cas-sette to orient a catalytic amino acid known as thesensor II motif in trans with the active site of an adja-cent protomer (Bae et al., 2009; Moreau et al., 2007),rather than in cis, which serves as the more usualarrangement in AAAþ oligomers. By comparison, theother two insertions are found in many other AAAþsubgroups and consist of two b–hairpins termed thepre-sensor I (PSI)-insert and the helix 2 (H2)-insert.Within the context of the hexamer, these b-hairpins arepositioned within the central channel and are integralto helicase mechanism (Figure 5). With regard to theMCM catalytic centers, a recent high-resolution cryo-electron microscopy structure of a full-length S. cerevi-siae MCM2–7 double hexamer reveals significant struc-tural variability between the six radially arrangedATPase sites, with a catalytically competent conform-ation but unequal nucleotide occupancy observed for

Figure 4. MCM architecture. (A) MCM homologs are characterized by three domains: NTD, AAAþ, and CTD. The NTD can be sub-divided into NTD-A (a small helical bundle), NTD-B (Zn-finger), and NTD-C (OB fold). The CTD forms a WH domain (for AAAþ andNTD, PDB¼ 3F9V; for WH domain, PDB¼ 2KLQ). (B) Two physiologically relevant MCM oligomers have been observed, a hexamerthat is formed by lateral interactions between the AAAþ and NTD domains of adjacent protomers, and a double hexamer thatis formed by interactions between the NTD-B Zn-finger domains of two MCM2–7 rings. The double hexamer structure from S.cerevisiae Mcm2–7 is shown (Li et al., 2015), with one hexamer faded compared to the other. The inset shows a top-down viewthrough the central cavity and the radial arrangement of eukaryotic MCM2–7 subunits (the double hexamer was built from twocopies of PDB¼ 3JA8 fit to the EM density map EMD-6338 (Li et al., 2015)). A color version of this figure is available at www.tandfonline.com/ibmg.

116 M. W. PARKER ET AL.

four of the sites (Li et al., 2015). This variability is con-sistent with the unequal ATPase activity observed forthe six MCM2–7 active sites (Bochman et al., 2008) andwith the nonequivalent function of these sites acrossthe different stages of helicase recruitment, loading andactivation (Coster et al., 2014; Ilves et al., 2010; Kanget al., 2014).

Insofar as the MCM CTD, solution structures haveshown this region to adopt a canonical winged-helix(WH) domain (Figure 4(A)) (Liu et al., 2012a; Wei et al.,2010; Wiedemann et al., 2015), which based on theweak or absent CTD density in structures of many full-length MCM proteins, appears flexibly tethered to theAAAþ core. Although the position of this domain withrespect to the hexamer was at first unclear, recent struc-tural and biochemical work demonstrates that the WH

domain sits distal to the NTD (Li et al., 2015;Wiedemann et al., 2015; Yuan et al., 2016). The S.solfataricus MCM WH domain can bind single-strandedDNA weakly (Pucci et al., 2007); however, what role thisactivity plays in vivo, and whether the WH domains ofthe eukaryotic MCMs can also bind DNA, are open ques-tions. The majority of data reported thus far suggeststhat, at least in archaea, the MCM WH domain functionsto allosterically modulate the ATP hydrolysis rate of theAAAþ domain (Barry et al., 2007; Jenkinson & Chong,2006; Wiedemann et al., 2015). In addition, this regioncan serve as a protein/protein interaction site, with theWH domains of eukaryotic MCM3 and MCM6 havingbeen shown to engage Cdc6 and Cdt1, respectively(Frigola et al., 2013; Liu et al., 2012a; You & Masai, 2008).Interestingly, recent structural analyses of the activated

Figure 5. Functional elements of MCM helicases. (A) Each MCM monomer contains multiple functional elements, including DNA-binding/sensing motifs, regions that modulate ATPase activity, and loops that communicate between the NTD and AAAþ domain(PDB ¼3JA8, chain 2). (B) In the context of a hexamer, the MCM functional elements (excepting the external b-hairpin) line a cen-tral cavity through which DNA translocates (modeled after PDB ¼3JA8, chains 4, 6, and 7). (C) Symbol key. (D) Detailed functionaldescription for each MCM element known to contribute to activity. A color version of this figure is available at www.tandfonline.com/ibmg.

CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY 117

yeast helicase (the CMG complex) have revealed thatthe Mcm5 and Mcm6WH domains partially occlude theMcm2–7 central channel, thus positioning them to per-haps function in the translocation process (Yuan et al.,2016). Despite this observation, the precise role of theMCM WH domains remains to be determined.

Although the ATPase activity of MCM proteins local-izes exclusively to the core AAAþ domain, hexamer for-mation is facilitated by both the AAAþ and NTDelements in the order MCM5!MCM3!MCM7!MCM4!MCM6!MCM2 (Costa et al., 2011; Crevel et al.,2001; Davey et al., 2003; Schwacha & Bell, 2001 ). A loopwithin each NTD-A OB-fold, termed the allosteric com-munication loop (ACL), contacts the AAAþ domain ofits adjacent protomer, further stabilizing the hexamer(Li et al., 2015; Miller et al., 2014). Collectively, theseinteractions facilitate the formation of a particle consist-ing of two stacked rings (a AAAþ tier and an NTD tier).In single and double MCM2–7 hexamers, the centralchannel is sufficiently large enough to accommodatedouble-stranded DNA (Figure 4(B)) (Costa et al., 2011,2014; Evrin et al., 2009; Li et al., 2015; Lyubimov et al.,2012; Remus et al., 2009); later, in an unknown series ofevents, the helicase undergoes an isomerization reac-tion that appears coupled to the extrusion of one of thetwo strands through a natural discontinuity betweenthe MCM2/5 subunits (Bochman & Schwacha, 2008;Bruck & Kaplan, 2015b; Costa et al., 2011), permittingtranslocation along single-stranded DNA (Fu et al., 2011;Ilves et al., 2010; Ishimi, 1997; Kelman et al., 1999;Shechter et al., 2000).

MCM mechanism in origin melting and processiveunwinding

The MCM2–7 complex matures through a variety ofintermediates before being incorporated into the activehelicase present at replication forks. Within theMCM2–7 lifecycle, two stable complexes are observed:an origin-bound double hexamer (Evrin et al., 2009;Remus et al., 2009) and a replication fork-associated sin-gle hexamer (Duzdevich et al., 2015; Fu et al., 2011).Interestingly, the double hexamer serves not only as aplatform for recruiting helicase-activating factors butalso is likely responsible for the initial origin meltingevent (Bochman & Schwacha, 2015; Li et al., 2015; Sunet al., 2014 ). Thus, the MCM complex must utilize theATP-hydrolysis-driven repositioning of its DNA-bindingelements to carry out two very distinct tasks, originopening as a double hexamer and processive DNAunwinding as a single hexamer. Although the MCM2–7DNA-binding elements are relatively well-defined, it iscurrently unclear whether the two functionalities of the

helicase require an overlapping or mutually exclusiveset of protein interactions with DNA.

The MCM2–7 hexamer has multiple tiers of DNA-binding elements that are positioned to engage DNApassing through the central pore (Figure 5). The NTD-BZn-fingers are situated at one end of the hexamer,forming a skirt of DNA-binding elements that run paral-lel to and extend the central channel. This domain isrequired for double hexamer formation, and mutationor deletion of the archaeal NTD-B reduces DNA bindingand results in loss of helicase activity (Kasiviswanathanet al., 2004; Poplawski et al., 2001; Pucci et al., 2004).The Zn-finger motifs of eukaryotic MCM2–7 also facili-tate double-hexamer formation (Li et al., 2015) and rep-resent a putative DNA-binding region, but the role ofthis domain in helicase activity has not been investi-gated directly. The Zn-finger motif of S. cerevisiae Mcm2and Mcm5 are required for cellular proliferation, sug-gesting an essential and conserved function for thisregion (Dalton & Hopwood, 1997; Yan et al., 1991).

As one moves through the central MCM2–7 pore,from the NTD to the CTD, the next DNA-binding ele-ments encountered are the b–hairpin insertions withinthe OB-fold (NTD-C). The NTD-C b–hairpin constricts thecentral channel (Fletcher et al., 2003; Li et al., 2015) and,in archaea, is electropositive in nature. Mutation of basicresidues within this loop and on the adjacent, pore-lin-ing surface of the OB-fold dramatically reduce bothDNA binding and helicase activity in archaeal MCM(Fletcher et al., 2003, 2008; McGeoch et al., 2005; Pucciet al., 2004). The NTD-C b-hairpin is highly variable inlength and composition within the eukaryotic MCM2–7family, and while a majority are enriched in basic resi-dues, this conservation is not universal. Nevertheless,the NTD-C region appears critical for eukaryotic MCMfunction, as S. cerevisiae strains harboring mutationswithin this b–hairpin of Mcm5 show an increased rateof minichromosome loss (Leon et al., 2008). This effectcan be repressed by the addition of multiple ARSsequences, which suggests that origin recruitment, butnot helicase activity, is defective in this particularmutant.

In addition to the NTD-C b–hairpin, the crystal struc-ture of an archaeal MCM NTD bound to single-strandedDNA has revealed a second DNA-binding region withinthe NTD-C, the MCM-single-stranded-DNA binding motif(MSSB) (Froelich et al., 2014). The MSSB is formed pri-marily by two positively charged residues that extendfrom the OB-fold and, interestingly, bind single-stranded DNA in an orientation perpendicular to thelong axis of the central channel. The MSSB DNA-bindingresidues are conserved among three consecutive subu-nits of the eukaryotic MCM2–7 complex (MCM4, MCM6

118 M. W. PARKER ET AL.

and MCM7), but not the other subunits. It has been sug-gested that the MSSB plays a role in DNA melting, work-ing against a DNA pumping action of the ATPasedomains to induce topological strain that encouragesstrand separation (Froelich et al., 2014). Thus, the MSSBmay have a role in the early steps of initiation, consist-ent with the defects observed for S. cerevisiae MSSBmutants in in vitro loading reactions (Froelich et al.,2014); whether or how this motif contributes to helicasefunction during elongation is unknown.

The AAAþ PSI b–hairpin insertion constitutes themost C-terminal DNA-binding element. This pore-liningloop shows the highest level of conservation of the vari-ous DNA-binding elements discussed thus far, and con-tains an invariant lysine that projects toward the centralpore. The PSI b-hairpin has been compared to the trans-location b–hairpin of SF3 helicases (e.g. the papillomavi-rus E1 protein and the SV40 Large T-antigen), which inthe context of a hexamer forms a vertically aligned,right-handed staircase that tracks the DNA backbonewith a conserved lysine (Enemark & Joshua-Tor, 2006,2008). Consistent with a critical role in helicase function,mutation of the PSI lysine in archaeal MCM weakensDNA binding and ATP hydrolysis, and fully abrogateshelicase activity (McGeoch et al., 2005); alteration of thisresidue within the context of the D. melanogaster CMGsimilarly abolishes helicase function (Petojevic et al.,2015). Interestingly, genetic analysis of the S. cerevisiaePSI b–hairpin lysines has highlighted nonequivalentfunctions for this residue in the six Mcm2–7 homologs,with only the mutation of this residue in Mcm3 abolish-ing an ability to complement deletion strains (Lamet al., 2013; Ramey & Sclafani, 2014).

How are the activities of the MCM2–7 DNA-bindingelements functionally coordinated with ATP turnoverto achieve the different functionalities observed forthe helicase? At least two other loops seem to facili-tate communication between the AAAþ domain andthe DNA-binding elements lining the central channel.One is a motif known as the H2-insert, which sits inthe MCM AAAþ fold and forms an extended loopthat junctions the ATPase and NTD tiers, creating apore-lining feature at the interface (Bae et al., 2009;Brewster et al., 2008; Li et al., 2015). Notably, analysisof the current data suggests that ATP-dependentrepositioning of the H2-insert may switch the helicasebetween an origin melting conformation to one thatfacilitates processive unwinding. Despite a low levelof sequence conservation, the length of the H2-insertis fully conserved in archaea and all MCM2–7 homo-logs and is sufficiently long to directly contact theMSSB. Interestingly, the H2-insert is rich in chargedamino acids and alignment of the six yeast Mcm

proteins reveals two fully conserved acidic residuesthat, in the context of the double hexamer, are posi-tioned to shield the MSSB from binding DNA (Figure5). Consistent with this proposal, the H2-insert dra-matically affects DNA binding in the archaeal MCMcomplex, with removal of the loop enhancing MCMaffinity for both double and single-stranded DNA(Jenkinson & Chong, 2006). The position of the H2-insert is thought to be modulated in an ATP-depend-ent fashion, such that the presence of ATP results ina more buried state for this loop (Jenkinson & Chong,2006). In conclusion, the available data suggest a crit-ical role for the H2-insert in transitioning the helicasebetween different functional states.

In addition to H2-insert-dependent crosstalkbetween MCM subunits, the ACL of NTD-C projectsupward from the OB-fold and toward the AAAþdomain of an adjacent protomer, where it is sand-wiched between the H2-insert and PSI b–hairpin(Barry et al., 2009; Li et al., 2015; Sakakibara et al.,2008). Like the other functional motifs within theNTD, the ACL is absolutely required for helicase activ-ity. However, the ACL does not affect DNA-bindingactivity, but instead modulates the ATPase activity ofthe AAAþ domain; mutation of ACL residues or dele-tion of the region results in markedly reduced ATPaseactivity and inhibition of duplex unwinding (Barryet al., 2009; Sakakibara et al., 2008). Like the H2-insert,the ACL position with respect to the AAAþ domain isaltered under different nucleotide-bound states(Barry et al., 2009).

Overall, the DNA-binding elements within the MCMNTD appear to play a critical function in both originmelting and unwinding. Although functional datapoint to an ATP-controlled connection between thepositional status of specific DNA binding elementswithin the MCM2–7 ring and their ability to grasp orrelease substrate, direct observation of these elementsin either a DNA melting or translocation mode is cur-rently lacking. Recent electron microscopy models ofthe activated eukaryotic MCM2–7 helicase in the con-text of the CMG have revealed that there exists con-formational coupling between the NTD and CTD tier,suggestive of coordinated action between theseregions during processive unwinding (Abid Ali et al.,2016; Yuan et al., 2016). NTD-A also exhibits conform-ational dynamics and has been seen to rotate mark-edly away from the hexamer axis, a movement thatmay be coordinated through direct binding of DNAto the NTD-A (Chen et al., 2005; Costa et al., 2008;Fletcher et al., 2003; Hoang et al., 2007; Liu et al.,2008). As additional substrate-bound structures aresolved for the archaeal and eukaryotic MCM

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homologs, exciting insights into its DNA remodelingand motor mechanisms are certain to emerge.

Loading of the replicative helicase

The defining step in eukaryotic replication licensing isloading of MCM2–7 onto duplex DNA into a stablehead-to-head double hexamer (Figure 1) (Evrin et al.,2009; Gambus et al., 2011; Remus et al., 2009). To attainthis state, single MCM2–7 hexamers are sequentiallyloaded by an interaction with ORC·Cdc6 that is cha-peroned by the Cdc10-dependent transcript 1 (Cdt1)protein. Structural characterization of multiple inter-mediates, together with the ability to reconstitute andstudy loading in vitro, are revealing how the stepwiseand carefully orchestrated exchange of pre-RC factors atorigins underlies this complex reaction (Figure 6).

Mechanism of helicase recruitment to origins

Maturation of the pre-RC occurs in a sequential fashionduring the G1 phase of the cell cycle (Evrin et al., 2009;Gillespie et al., 2001; Romanowski et al., 1996b; Remuset al., 2009; Ticau et al., 2015; Tsakraklides & Bell, 2010;Tsuyama et al., 2005). Demarcation of replication startsites by ORC represents the inaugurating event, and

permits the recruitment of and stable association withCdc6 (Cocker et al., 1996; Coleman et al., 1996; Donovanet al., 1997; Liang et al., 1995; Seki & Diffley, 2000;Tsuyama et al., 2005). In an in vitro setting, Cdc6 regu-lates the fidelity of origin selection by two distinctmechanisms. First, Cdc6 helps to restrict the initiatorfrom acting at illegitimate origins by sequestering thefree initiator (thereby lowering the effective concentra-tion of the initiator to increase origin specificity)(Duzdevich et al., 2015), and by triggering the dissoci-ation of ORC from nonorigin DNA (Mizushima et al.,2000). Second, Cdc6 ATPase activity is enhanced whenbound to non-ARS sequences, which triggers the dis-sociation of Cdc6 from the initiator (Speck et al., 2005;Speck & Stillman, 2007). Ultimately, the secure associ-ation between Cdc6 and ORC at origins primes the initi-ator for helicase recruitment and loading (Donovanet al., 1997; Feng et al., 2000; Perkins & Diffley, 1998;Rowles et al., 1996; Tsuyama et al., 2005); this samecomplex will later facilitate the release of the doublehexamer for subsequent activation at the onset of Sphase (Chang et al., 2015).

The MCM2–7 helicase must first be recruited to ori-gins to initiate the loading reaction. Notably, the heli-case is sensitive to solution conditions and showssubstantial conformational heterogeneity (Bochman &

Figure 6. MCM2–7 complex loading and maturation into the CMG. Two sequential rounds of helicase recruitment and loading atorigins are required for building an MCM2–7 double-hexamer. For both hexamers, DNA is threaded into the central channelthrough a discontinuity between MCM2 and MCM5. The first hexamer is recruited through direct interactions with the initiator(MCM3-Cdc6) and may require Cdt1 for overcoming an MCM6-mediated autoinhibited state of the helicase. After the first hex-amer loads, both Cdc6 and Cdt1 are released. Rebinding of Cdc6 to ORC primes the system for recruiting and loading a secondhexamer in the opposite direction as the first, an event that has been proposed to be controlled by ORC·Cdc6, but templated bythe first MCM2–7 hexamer. Cdt1 and Cdc6 recruitment and ejection are required for both loading events. Phosphorylation of thedouble hexamer and other initiation factors by CDK and DDK facilitate origin melting, GINS and Cdc45 recruitment/assembly, DNAstrand extrusion, and activation of the helicase for DNA unwinding. A color version of this figure is available at www.tandfonline.com/ibmg.

120 M. W. PARKER ET AL.

Schwacha, 2008; Costa et al., 2011; Gomez-Llorenteet al., 2005), a feature that is pertinent for understand-ing the mechanism of association with the origin-boundinitiator. Biochemical and structural analyses of theMCM2–7 complex has revealed that, in the absence ofATP, archaeal, fungal and metazoan complexes canadopt a cracked ring architecture that bears a discon-tinuity, or “gate”, between MCM2 and MCM5 (Bochman& Schwacha, 2008; Boskovic et al., 2016; Costa et al.,2011; Lyubimov et al., 2012; Samson et al., 2016).Nucleotide binding constricts the ring and favors theformation of MCM2/5 interactions across the ring break(Samel et al., 2014), but does not appear to irreversiblyclose the gate in single MCM2–7 hexamers (Costa et al.,2011; Lyubimov et al., 2012). Thus, MCM2–7 may notrequire active ring-opening prior to loading, but insteadmay simply be stably aligned around DNA by origin-bound ORC·Cdc6.

In addition to the ORC·Cdc6 helicase-loader complex,Cdt1 is also required for helicase loading in yeast andmetazoa (Claycomb et al., 2002; Devault et al., 2002;Maiorano et al., 2000; Nishitani et al., 2000; Whittakeret al., 2000). However, the role of this factor has beenenigmatic. Although Cdt1 was initially thought to berequired for helicase recruitment to ORC (Asano et al.,2007; Chen et al., 2007; Chen & Bell, 2011), recent datasuggest that MCM2–7 is recruited by direct interactionswith the initiator (Fernandez-Cid et al., 2013; Frigolaet al., 2013). With the exception of Orc6, all initiationfactors discussed thus far possess a central AAAþATPase unit; Cdt1 represents an additional non-AAAþprotein involved in the process. Cdt1 was first identifiedin S. pombe, and through an ability to induce re-replica-tion in the absence of cell division, was recognized asan essential and potent initiator of replication(Hofmann & Beach, 1994; Nishitani et al., 2000). Cdt1 isuniversally conserved in metazoa but is inconsistentlyfound in other eukaryotic supergroups (Aves et al.,2012; Maiorano et al., 2000; Tada et al., 1999; Whittakeret al., 2000). The weak conservation of Cdt1 acrossspecies aggravated initial attempts to identify an S.cerevisiae homolog, but eventually the TAH11(Topoisomerase-A hypersensitive 11) protein was identi-fied as the budding yeast counterpart (Devault et al.,2002; Tanaka & Diffley, 2002). TAH11 was first isolatedin genetic screens for genes that showed syntheticlethality with a mutant topoisomerase I allele (Fiorani &Bjornsti, 2000), and proteomic studies in human cellshave suggested there exists an interaction betweenCdt1 and both topoisomerase I and topoisomerase IIa(Sugimoto et al., 2008). Archaea also contain a Cdt1-related protein, termed WhiP; however, this factorappears to uniquely function in the Orc-independent

assembly of pre-RCs (Robinson & Bell, 2007; Samsonet al., 2013).

Interestingly, Cdt1 has markedly diverged between S.cerevisiae and metazoa, and functional analyses havesuggested that there exist differing mechanisms forCdt1-dependent action across species. All Cdt1 homo-logs are built from a conserved C-terminal pair of WHdomains (Khayrutdinov et al., 2009; Lee et al., 2004). Bycontrast, the N-terminal region of Cdt1 is not conserved,and while the N-terminal domain of metazoa and S.pombe Cdt1 has a basic pI (human Cdt1 pI¼ 10.5) thatis predicted to be unstructured, the S. cerevisiae N-ter-minus has an acidic pI (pI¼ 5.1) and possesses sufficientsecondary structure to likely constitute an ordereddomain (modeling using Phyre2 (Kelley et al., 2015) sug-gests that this region corresponds to a catalyticallydefunct oxygenase fold). Interestingly, an S. cerevisiaeCdt1 construct lacking the nonconserved N-terminaldomain fails to load the helicase into an activation-com-petent conformation (Fernandez-Cid et al., 2013; Takara& Bell, 2011), whereas the N-terminus of metazoa Cdt1is dispensable for the loading reaction (Ferenbach et al.,2005).

The factors with which Cdt1 stably associates alsoevince species-specific differences. In S. cerevisiae, Cdt1and Mcm2–7 form a tight complex and exhibit inter-dependent nuclear import (Kawasaki et al., 2006; Remuset al., 2009; Takara & Bell, 2011; Tanaka & Diffley, 2002);Cdt1 similarly helps maintain the structural integrity ofthe Mcm2–7 complex (Frigola et al., 2013; Wu et al.,2012). Conversely, Xenopus Cdt1 and MCM2–7 are bio-chemically separable initiation factors, with immunode-pletion of MCM3 from Xenopus egg extracts removingMCM2–7 but having no effect on Cdt1 (Maiorano et al.,2004) (it is worth noting that mouse Cdt1 has beenreported to form a complex with MCM2–7 in vitro (You& Masai, 2008), but the stability of this assembly has notbeen investigated). Multiple explanations could accountfor the differences in behavior between S. cerevisiae andXenopus Cdt1; for example, differences in the relativeaffinities of Cdt1 for the MCM2–7 complex could reflectan alteration to the sequential assembly of the pre-RCfactors in metazoa, such that Cdt1 associates first withORC·Cdc6 rather than with the helicase (Maiorano et al.,2000; Waga & Zembutsu, 2006). Finally, metazoa con-tain an additional Cdt1-binding protein, Geminin, whichnegatively regulates licensing by inhibiting the Cdt1-dependent loading of the helicase into a stable doublehexamer (Edwards et al., 2002; Wohlschlegel et al., 2000;Wu et al., 2014a; Yanagi et al., 2002).

Despite the differences in Cdt1 structure and its pre-ferred binding partners between budding yeast andmetazoans, multiple studies have identified MCM6

CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY 121

as the primary Cdt1 interaction site on the helicasehexamer (Ferenbach et al., 2005; Liu et al., 2012a; Weiet al., 2010; Wu et al., 2012; Yanagi et al., 2002; Zhanget al., 2010) (mouse Cdt1 also interacts with MCM2 (You& Masai, 2008)). Cdt1/MCM6 contacts are facilitated bythe C-terminal WH domain of each protein (Ferenbachet al., 2005; Jee et al., 2010; Khayrutdinov et al., 2009;Takara & Bell, 2011; Teer & Dutta, 2008; Wei et al., 2010),with structural studies of the human proteins revealingthat a helical extension which projects from theCdt1WH region engages the helix-turn-helix motif ofthe MCM6 C-terminus (Liu et al., 2012a; You & Masai,2008). Mutagenesis of residues in the interaction surfaceabolishes helicase loading and DNA replication in bud-ding yeast, suggesting that the MCM6-Cdt1 mechanismof binding is widely conserved (Liu et al., 2012a).Despite this contact, in vitro binding studies have sug-gested that there exist additional undefined sites ofinteraction between Cdt1 and MCM2–7 (Fernandez-Cidet al., 2013; Khayrutdinov et al., 2009).

Saccharomyces cerevisiae Cdt1 has also beenreported to bind Orc6 (Asano et al., 2007; Chen et al.,2007; Chen & Bell, 2011; Semple et al., 2006). This inter-action was initially proposed to facilitate helicaserecruitment by helping to tether MCM2–7 to theORC·Cdc6 complex (it is unclear whether metazoanCdt1 and Orc6 interact (Yanagi et al., 2002)); however,other work has indicated that S. cerevisiae Mcm2–7 con-tains elements that directly engage ORC·Cdc6 and thatare wholly sufficient for recruitment (Fernandez-Cidet al., 2013; Frigola et al., 2013). Cdt1’s participation inmodulating MCM/ORC·Cdc6 interactions is still underdebate, with one study noting that budding yeastCdt1 relieves an Mcm6-dependent autoinhibitorymechanism that prevents Cdt1-independent recruit-ment (i.e. deletion of an Mcm6 autoinhibitory domainresults in helicase recruitment in the absence of Cdt1)(Fernandez-Cid et al., 2013), and another showing thatCdt1 is fully dispensable for helicase recruitment (hereonly an interaction between Cdc6 and the extreme C-terminus of Mcm3 appears necessary for recruitment(Frigola et al., 2013). Consistent with the proposed roleof the eukaryotic MCM3 C-terminus, recent studies inarchaea have found that the MCM C-terminal WHdomain directly binds Orc to facilitate recruitment ofthe helicase to replication origins (Samson et al., 2016).

Together, the available data suggest that Cdt1 maybe dispensable for helicase recruitment, and that thisstep instead predominantly relies on interactionsbetween ORC·Cdc6-bound origins and MCM2–7. Whatthen might be the significance of Cdt1 and its inter-action with Orc6? Beyond a potential ability to relieveMCM6-dependent autoinhibition (Fernandez-Cid et al.,

2013), S. cerevisiae Cdt1 stabilizes Mcm2–7 at origins inan Orc6-dependent manner (Chen et al., 2007), suchthat in the absence of Cdt1, topologically linkedMcm2–7 hexamers fail to load (Chen & Bell, 2011).Notably, an N-terminal deletion construct of buddingyeast Cdt1 permits loading of double hexamers, but ina state that is incapable of being subsequently acti-vated (Takara & Bell, 2011). Thus, Cdt1 is required forevents downstream of recruitment, which appear toinclude the formation of an MCM2–7 double hexamerintermediate that is competent for replication. Overall,the molecular details underlying the role of Cdt1, and inparticular its Orc6 association, are in need of furtherstudy.

Loading the double hexamer

Ultimately, the initiator-dependent recruitment ofMCM2–7 to origins leads to the formation of a stablechromatin-bound MCM2–7 complex that, unlikeORC·Cdc6, is resistant to high-salt extraction (Bowerset al., 2004; Donovan et al., 1997; Edwards et al., 2002).Notably, the MCM2–7 loading reaction has been fullyreconstituted in vitro with purified proteins from S. cere-visiae (Evrin et al., 2009; Remus et al., 2009), which hasin turn allowed the use of single molecule and struc-tural techniques to understand both the nature of thesalt-stable complex and the events leading to its forma-tion. The details of the loading reaction discussed in thefollowing are derived exclusively from the studies on S.cerevisiae initiation factors; given some of the species-specific differences observed to date, it will be import-ant to reconstitute the loading reaction in vitro withother organisms for comparison.

Following the recruitment of Cdt1·Mcm2–7 toORC·Cdc6-bound origins, Mcm2–7 is deposited ontoDNA while both Cdc6 and Cdt1 are ejected from thepre-RC in an ATP hydrolysis-dependent fashion. Cdc6release occurs prior to Cdt1 both for single- and dou-ble-hexamer loading (Evrin et al., 2009; Kang et al.,2014; Remus et al., 2009; Ticau et al., 2015; Tsakraklides& Bell, 2010) (Figure 6). Singly-loaded Mcm2–7 hexam-ers result in the transient formation of a meta-stableORC·Cdc6·Cdt1·Mcm2–7 (OCCM) intermediate, whichcan be stabilized by the presence of a nonhydrolyzableATP analog. This property has been exploited to permitimaging of the complex by 3D cryo-electron microscopy(Sun et al., 2013). Within the OCCM, Cdt1 interfaces withthe N-terminal tier of Mcm2, Mcm6 and Mcm5. TheMcm3 C-terminus resides proximal to Cdc6, consistentwith the observed interaction between these regions invitro and the proposed role of this interaction in heli-case recruitment (Frigola et al., 2013; Sun et al., 2013).

122 M. W. PARKER ET AL.

Cdt1 and Orc6 make no visible contacts in the OCCMstructure; the N-terminal TFIIB domain of Orc6 also doesnot appear ordered. Notably, the Cdt1·Mcm2–7 hep-tamer is oriented such that the MCM C-terminaldomains (the region containing the AAAþ and WHdomains) abut the ORC WH domains (Bleichert et al.,2015), leaving the MCM NTD accessible. Although ana-lysis of the loading reaction in bulk has indicated thatsingly loaded Mcm2–7 hexamers are salt-sensitive (Evrinet al., 2009; Kang et al., 2014; Remus et al., 2009), singlemolecule analysis of the reaction reveals that nearly50% of the singly loaded helicases are salt-stable, sug-gesting that at this stage Mcm2–7 ring closure hasoccurred (Ticau et al., 2015). This discrepancy betweenbulk and single-molecule studies currently lacks explan-ation, although may arise from the relatively high pro-tein concentrations used in the bulk assays and/or bytheir limited kinetic sensitivity.

After the first loading event, Cdc6 re-associates withthe ORC·Mcm2–7 complex, forming a short-livedORC·Cdc6·Mcm2–7 (OCM) intermediate that is compe-tent for recruitment of a second Cdt1·Mcm2–7 hep-tamer (Fernandez-Cid et al., 2013; Ticau et al., 2015).At present, the mechanics by which a second hexameris recruited are unclear. It has been proposed that load-ing of the second hexamer requires equivalent interac-tions as the first, with the Mcm3 C-terminus beingrequired for both loading steps (Frigola et al., 2013).However, if loading the second helicase into a doublehexamer requires an interaction with ORC·Cdc6,this mechanism must involve some fairly complicatedacrobatics, mainly because the first loaded Mcm2–7complex should still be present to block the MCM-bind-ing surface on ORC, and also because the secondMcm2–7 complex must be placed at a location morethan �100Å from ORC’s initial binding site on DNA, inan inverted orientation from the first helicase. This spa-tial problem could be solved if two ORCs were used forthe loading of two helicases with opposing polarity;however, single-molecule analysis has demonstratedthat a single, stably bound ORC is sufficient to load adouble hexamer (Ticau et al., 2015). Because of thesesteric complications, an alternative mechanism hasbeen proposed wherein the first, loaded hexamer tem-plates loading of the second through a distinct mechan-ism that does not require direct ORC·Cdc6 interactions(Ticau et al., 2015). Consistent with this idea, the kineticsof loading the second hexamer are slower than the first(Fernandez-Cid et al., 2013), as is the release of Cdc6and Cdt1 (Ticau et al., 2015), suggesting different reac-tion intermediates are accessed.

While the model described above does not directlyaccount for the demonstrated role of the budding yeast

Mcm3WH domain in the loading of both the first andsecond Mcm2–7 hexamers (Frigola et al., 2013), previouswork has revealed that deletion of the Mcm3 C-ter-minus results in a dramatic loss of Mcm2–7 ATPaseactivity (Sun et al., 2014). Since proper ATPase functionby the Mcm2–7 hexamer is critical at all stages of theloading reaction in S. cerevisiae, this loss of activity inthe Mcm3WH domain mutant could account for theobserved defect in loading the second hexamer.Accordingly, Mcm2–7 complexes with individual activesite mutations also show defective recruitment (particu-larly Mcm2 Walker A and Mcm6 arginine fingermutants), Cdt1 release, and dramatically reduced levelsof salt-stable double hexamers (Coster et al., 2014; Kanget al., 2014). Although a marginal level of double-hex-amer formation can be achieved with particular MCMATPase mutants, these loaded complexes are generallydeficient for downstream activation events and cannotdrive DNA replication in vitro (Kang et al., 2014).Collectively, the present data indicate that ATP bindingand hydrolysis by S. cerevisiae Mcm2–7 is required fromthe early steps of helicase recruitment to the conclud-ing activation events. These results are difficult torationalize with the observation that Xenopus MCM6and MCM7 Walker A mutants are loaded normally(Ying & Gautier, 2005), but this finding may again reflectspecies-specific differences.

Although double-hexamer loading is criticallydependent on the ATPase activity of Mcm2–7, ATPhydrolysis by Cdc6 and ORC perform additional rolesthat are consistent with a requirement for their ATPaseactivity in vivo (Bowers et al., 2004; Schepers & Diffley,2001; Takahashi et al., 2002; Weinreich et al., 1999;Zwerschke et al., 1994). ATP hydrolysis by Orc1 is dis-pensable for the loading reaction (Evrin et al., 2013;Fernandez-Cid et al., 2013), but resets the initiationmachinery to allow reiterative loading of MCM doublehexamers at origins (Bowers et al., 2004). By compari-son, Cdc6 has been reported to function as a qualitycontrol factor, whose ATPase activity ensures thatdefective or incomplete Mcm2–7 complexes arereleased from origins (Coster et al., 2014; Frigola et al.,2013; Kang et al., 2014). Another major function of Cdc6ATPase activity appears to be in the final step of pre-RCformation, where it is responsible for releasing success-fully loaded double hexamers from ORC, an event thatenables the downstream activation of the helicase(Chang et al., 2015). Loss of Cdc6 ATPase activity alsoresults in inefficient Cdt1 release (Randell et al., 2006),although ATP turnover by Mcm2–7 appears primarilyresponsible for Cdt1 ejection (Coster et al., 2014; Kanget al., 2014). Overall, Cdc6 functions as a linchpin factorcritical for multiple events in the loading process.

CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY 123

Helicase activation

Although a double MCM2–7 hexamer is loaded duringpre-RC formation, at replication forks this structure isdissolved to yield two single MCM2–7 hexamers thatmove in opposite directions (Ticau et al., 2015; Yardimciet al., 2010). On their own, single MCM2–7 hexamerspossess little or no helicase activity (Ishimi, 1997; Ilveset al., 2010; Kaplan et al., 2003; Lee et al., 2000; Youet al., 2003). Initially, this observation was difficult torationalize with the strong evidence that pointedtoward a role for the complex as the replicative heli-case, but it is now appreciated that MCM2–7 is incorpo-rated into a larger Cdc45·MCM2–7·GINS (CMG) complexthat possesses robust helicase activity. Like pre-RCassembly, double hexamer dissolution and CMG forma-tion occur through a highly orchestrated and intercon-nected series of events that provide yet another meansfor regulating the DNA replication initiation reaction.

Identification of the Cdc45·MCM2–7·GINS (CMG)complex

The MCM2–7 genes are essential proteins in DNA repli-cation (Dalton & Whitbread, 1995; Gibson et al., 1990;Moir et al., 1982; Maine et al., 1984) and their phylogen-etic lineage to helicases led to early suggestions thatthese factors might function as nucleic acid motor pro-teins (Koonin, 1993). Together with work on archaealMCMs, which have been found to possess robust heli-case activity (Chong et al., 2000; Kelman et al., 1999;Shechter et al., 2000), these data indicated that theMCM2–7 complex likely served as the replicative heli-case in eukaryotes. Interestingly, initial in vitro studieswere unable to detect helicase activity from the eukary-otic MCM2–7 complex, although an MCM subcomplexcomposed of subunits (4,6,7)2 showed weak helicaseactivity (Ishimi, 1997; Kaplan et al., 2003; Lee et al., 2000;You et al., 2003). These data suggested that theMCM2–7 assembly might play a more limited role inreplication, such as origin melting. However, the persist-ence of MCM2–7 at replication forks (Aparicio et al.,1997), coupled with the observed S-phase arrest afterMCM2–7 inactivation (Labib et al., 2000; Pacek & Walter,2004; Shechter et al., 2004), provided irrefutable evi-dence for the role of the complex in elongation (Krudeet al., 1996; Todorov et al., 1995). Nevertheless, the dem-onstration of helicase activation remained elusive.

Eventually, conditions were identified that recoveredin vitro helicase activity for budding yeast Mcm2–7(Bochman & Schwacha, 2008). Roughly contemporan-eously, several lines of analysis began to reveal that thehelicase also existed within a larger macromolecular

assembly (Calzada et al., 2005; Gambus et al., 2006;Masuda et al., 2003; Moyer et al., 2006; Pacek et al.,2006). Three parallel studies – two using an unbiasedassessment of Cdc45 and GINS interacting factors and athird using a candidate-based approach to determinethe composition of the helicase stalled at replicationforks – have now demonstrated that MCM2–7 forms astable complex with Cdc45 and GINS (Gambus et al.,2006; Moyer et al., 2006; Pacek et al., 2006). Cdc45 andGINS are essential replication fork components (Dalton& Hopwood, 1997; Hopwood & Dalton, 1996; Kamimuraet al., 1998; Kanemaki et al., 2003; Masuda et al., 2003;Moir et al., 1982; Zou et al., 1997), which mimic thechromatin association pattern of MCM subunits (Kubotaet al., 2003; Takayama et al., 2003). Each MCM2–7 bindsa single copy of Cdc45 and one GINS tetramer to form astable, 11-protein complex termed the CMG (forCdc45·MCM2–7·GINS). The CMG has been shown to pos-sess robust helicase activity (Ilves et al., 2010; Moyeret al., 2006), and bioinformatics analyses have indicatedthat a full set of CMG factors are likely conserved fromarchaea to eukaryotes (Makarova et al., 2012), withrecent work demonstrating that, as observed for theeukaryotic accessory factors, archaeal Cdc45 and GINShomologs activate MCM in vitro and in vivo (Xu et al.,2016).

Regulating helicase activation

Appropriate recruitment of the factors necessary forCMG formation constitutes a highly regulated steptoward replisome formation. Consistent with the Cdc6-dependent release of the MCM2–7 double hexamerfrom ORC (Chang et al., 2015), helicase activation nolonger requires the function of either ORC or Cdc6 (Hua& Newport, 1998; Jares & Blow, 2000; Rowles et al.,1999; Walter, 2000; Yeeles et al., 2015). The newlyformed MCM2–7 double hexamer serves itself as a bind-ing platform and target for the Dbf4-dependent kinase(DDK) that, in collaboration with S-phase cyclin-depend-ent kinase (S-CDK), constitute the minimal set of kinasesneeded to regulate the recruitment of Cdc45 and GINS.Together, the activity of these two kinases coordinatethe assembly of a pre-IC that is poised for the activationand formation of a bidirectional replication fork(Figure 6) (Aladjem, 2007).

DDK is formed by the direct association of the Cdc7kinase with Dbf4, a protein co-factor that relieves Cdc7autoinhibition (Dowell et al., 1994; Jackson et al., 1993;Kitada et al., 1992) and that is regulated in a cell-cycle-dependent manner (Cheng et al., 1999; Oshiro et al.,1999; Nougarede et al., 2000). DDK phosphorylates mul-tiple MCM subunits (Cho et al., 2006; Lei et al., 1997;

124 M. W. PARKER ET AL.

Masai et al., 2006; Montagnoli et al., 2006; Sheu &Stillman, 2006; Weinreich & Stillman, 1999; Tsuji et al.,2006) and is required for initiating DNA replication(Chapman & Johnston, 1989; Hollingsworth & Sclafani,1990; Jiang et al., 1999a). Like DDK, S-CDK is requiredfor replication (Broek et al., 1991; Hayles et al., 1994)and exists in an inactive conformation by engagingSic1, a repressive protein that restricts S-CDK activityinto late G1/S phase (Donovan et al., 1994; Knapp et al.,1996; Mendenhall, 1993; Schneider et al., 1996; Schwobet al., 1994). Redundant regulation of S-CDK occurs bythe cell-cycle-dependent expression of activating cyclins(Abreu et al., 2013). It is currently unclear whether thereis a conserved sequential order for DDK and CDK activ-ity. In a X. laevis egg extract system, DDK activity is inde-pendent of CDK, and exposing the pre-RC first to CDKfully ablates DNA replication initiation (Jares & Blow,2000; Walter, 2000), suggesting that DDK acts first. Sucha consensus has not been found with S. cerevisiae pro-teins, where the sequential action of DDK and CDKdepends on the experimental setup (Heller et al., 2011;Nougarede et al., 2000) and has been found to beinconsequential in a fully in vitro reconstituted DNA rep-lication system (Yeeles et al., 2015).

DDK is recruited to Mcm2–7 through interactionswith Mcm4 and Mcm2 (Ramer et al., 2013; Sheu &Stillman, 2010) and targets specific MCM subunitswithin the context of a double, but not a single,MCM2–7 hexamer (Costa et al., 2014; Evrin et al., 2014;Francis et al., 2009; Kang et al., 2014; Sun et al., 2014).Notably, the MCM subunits responsible for recruitingDDK are nonadjacent within the context of a single hex-amer; however, the rotational offset within the doublehexamer results in the adjacent placement of MCM2and MCM4 from separate hexamers (Costa et al., 2014;Li et al., 2015; Sun et al., 2014), thus forming a compos-ite DDK-interaction interface. In addition, MCM2, MCM4and MCM6, which are all phosphorylated by DDK (Bruck& Kaplan, 2009; Cho et al., 2006; Lei et al., 1997; Masaiet al., 2006; Randell et al., 2010; Sheu & Stillman, 2006;Tsuji et al., 2006), reside in close proximity within thedouble hexamer and the interface between the twohexamers is somewhat splayed apart at this position,creating a gap that may provide DDK access to the N-terminal serine/threonine-rich domains (NSD) of itsMCM targets (Li et al., 2015; Sheu & Stillman, 2010; Sunet al., 2014). Collectively, the available data suggest thatthe DDK-dependent activation of MCM2–7 is temporallyregulated by selective recruitment of the kinase to andaction upon a double hexameric MCM2–7, a structurethat uniquely positions the relevant MCM subunits inclose proximity (Sun et al., 2014). In addition to DDK,budding yeast Mcm2–7 has been shown to be

phosphorylated by CDK and Mec1 kinases, which sensi-tize Mcm2–7 to DDK activity (Devault et al., 2008;Randell et al., 2010); however, DDK alone is required forthe activation of S. cerevisiae Mcm2–7 in vitro (Yeeleset al., 2015).

In S. cerevisiae, the DDK-dependent phosphorylationof Mcm2–7 is important for facilitating the Sld3 andSld7-chaperoned recruitment of Cdc45 onto the doublehexamer (Masai et al., 2006; Sheu & Stillman, 2006;Tanaka et al., 2011a; Yabuuchi et al., 2006). In humans,two proteins known as Treslin and MTBP (for MDM2binding protein) have been proposed to represent func-tional homologs of budding yeast Sld3 and Sld7,respectively (Boos et al., 2013; Bruck & Kaplan, 2015c;Kumagai et al., 2010; Matsuno et al., 2006; Sanchez-Pulido et al., 2010; Sansam et al., 2010). In vitro, Cdc45recruitment depends on the prior association of Sld3and Sld7 with the DDK-phosphorylated N-termini ofMcm2, Mcm4 and Mcm6 (Deegan et al., 2016; Fanget al., 2016); however, in vivo Sld3/7 and Cdc45 form astable complex and may be co-recruited to double hex-amers (Kamimura et al., 2001; Tanaka et al., 2011a,).Notably, in the absence of DDK, no additional initiationfactors are recruited to Mcm2–7 double hexamers.Conversely, in the absence of CDK activity, Cdc45 aswell as Sld3 and Sld7 are still efficiently recruited (Helleret al., 2011; Yeeles et al., 2015). Thus, recruitment ofCdc45 appears to rely exclusively on DDK.

How does DDK-dependent MCM2–7 phosphorylationfacilitate Cdc45 recruitment? Although we lack a com-plete mechanistic picture for DDK-dependent activation,three distinct MCM modifications have been identifiedthat result in DDK bypass, suggesting there exists acomplex role for DDK that likely involves a concertedchange within multiple MCM subunits. First, hyperphos-phorylation of the S. cerevisiae Mcm4 N-terminusrelieves an autoinhibitory function that this regionimposes on the helicase (indeed, deletion of the Mcm4N-terminus bypasses the DDK requirement) (Sheu &Stillman, 2006, 2010). Although a mechanism entailing asteric block to the Cdc45-binding site or the adoptionof a different nonpermissive orientation would satisfythis observation, within the atomic structure of the S.cerevisiae Mcm2–7 double hexamer the Mcm4 N-ter-minus is disordered, suggesting that autoinhibition isrelieved by an alternative mechanism (Li et al., 2015).Second, phosphorylation of Mcm4 and Mcm6 generatesa phosphopeptide-binding site for recruiting Sld3(phosphomimetic mutants of these MCM subunits canbypass the requirement for DDK) (Deegan et al., 2016).Finally, a mutation within budding yeast Mcm5, knownas the bob1 allele, has been found to bypass DDK-activated initiation (Hardy et al., 1997).

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In addition to DDK-dependent recruitment of Cdc45,Mcm2–7 activation requires an association with GINSand, for origin firing, with Replication Protein A (RPA),MCM10, Pola, Pole and CTF4 (van Deursen et al., 2012,Yeeles et al., 2015). Notably, CDK’s action on the grow-ing, chromatin-associated replication assembly, as wellas on replicative proteins yet to be integrated into thereplisome, results in the recruitment of the remaininginitiation factors that are needed for origin firing. In thisswarm of activity, CDK targets Cdc45, Sld2, Sld3 andSld7 (Heller et al., 2011; Masumoto et al., 2002;Muramatsu et al., 2010; Tanaka et al., 2007; Yeeles et al.,2015; Zegerman & Diffley, 2007), and, together with aprotein known as Dpb11, facilitates recruitment of amultiprotein assembly composed of GINS, Sld2 and Poleto the growing complex (Araki et al., 1995; Kamimuraet al., 1998, 2001; Muramatsu et al., 2010; Takayamaet al., 2003; Tanaka et al., 2013). Thus, DDK and CDKdrive formation of an active helicase by phosphorylat-ing key assembly factors that allow for the sequentialrecruitment of Cdc45 and GINS. For its part, MCM10 isrecruited by interactions with the CMG (Douglas &Diffley, 2016; Homesley et al., 2000; Wohlschlegel et al.,2002) and further appears to help stabilize the repli-some once formed (Gregan et al., 2003; Ricke &Bielinsky, 2004). Pole is likewise recruited directly to theCMG by an interaction with GINS (Araki et al., 1995;Muramatsu et al., 2010), whereas Pola is physicallycoupled to the helicase by CTF4 (Simon et al., 2014;Villa et al., 2016).

Many questions remain for the events that transitionthe growing MCM2–7 complex into a competent bi-dir-ectional replication fork. For example, it is currentlyunclear at what point and how origin melting occurs, orhow an MCM2–7 double hexamer, which encirclesduplex DNA, transitions into two uncoupled CMG par-ticles that encircle single-stranded DNA (Ticau et al.,2015; Yardimci et al., 2010). Interestingly, recent datasuggests that the process of origin melting and strandextrusion may be interdependent and temporally inter-twined with CMG assembly. Indeed, S. cerevisiae Sld2,Sld3 and Dpb11, which chaperone GINS into a complexwith the helicase, compete with GINS for Mcm2–7 bind-ing, a function that is relieved in the presence of single-stranded DNA (Bruck et al., 2011; Bruck & Kaplan, 2011,2014, 2015c; Dhingra et al., 2015) (the metazoan Sld2and Sld3 equivalents also bind single-stranded DNA(Bruck & Kaplan, 2015c; Ohlenschlager et al., 2012;Sangrithi et al., 2005)). Thus, GINS binding may not onlybe anticipated by origin melting and the generation ofsingle-stranded DNA, but by strand extrusion as well, asthe separated origin strands may be contained withinthe Mcm2–7 double hexamer for a period of time

(Li et al., 2015; Sun et al., 2014). The topological transfor-mations to DNA in the context of the maturing CMGcomplex are likely aided in by the phosphorylation ofMcm2 by DDK (Bruck & Kaplan, 2009; Lei et al., 1997),an event that weakens the Mcm2/5 gate and promotesring opening (Bruck & Kaplan, 2015a,b). Consistent withthis idea, origin melting and strand extrusion have beenreported to occur concomitantly with DDK-dependentactivation but prior to complete CMG assembly (Bruck &Kaplan, 2015a). Interestingly, recent data suggests thatthe single-stranded DNA-binding activity of buddingyeast Mcm10 is required for initiation and that it mayhelp stabilize the origin melting reaction (Perez-Arnaiz& Kaplan, 2016).

Despite the available insights, the mechanisms thattransition a loaded double hexamer into an active, bi-directional replication fork remain shrouded in mystery.An important future direction will be to understand thetiming of different events in the lifecycle of the helicase,such as when double-hexamer dissociation and repli-some factor recruitment occurs (such as MCM10, Polaand RPA), as compared to origin melting and strandextrusion. The physical mechanisms that coordinateCMG structural changes with origin remodeling likewiseremain ill-defined, and require structural studies of dif-ferent intermediates that, at present, have not been sta-bly isolated.

Mechanism of CMG function

The cracked ring architecture of the MCM2–7 complexis an asset when it comes to loading the helicasearound double-stranded DNA, but would seem to be apotential liability in terms of processive DNA unwindingfunctions (Bochman & Schwacha, 2008; Ilves et al., 2010;Petojevic et al., 2015). Although nucleotide binding byMCM2–7 helps to overcome a natural tendency of thehelicase to splay open (Samel et al., 2014), cracked-ringconformations still readily form (Costa et al., 2011;Lyubimov et al., 2012), providing a physical basis for theweak in vitro helicase activity of MCM2–7 (Bochman &Schwacha, 2008; Ilves et al., 2010). Incorporation ofMCM2–7 into the CMG counteracts the physical defi-ciencies of the MCM2–7 complex for DNA unwinding.3D electron microscopic reconstructions of both theDrosophila and budding yeast CMG have revealed thatGINS and Cdc45 seal off the MCM2/MCM5 discontinuityand reduce the conformational dynamics of the MCM2/5 gate, likely as a means to favor productive ATPaseinterface contacts (Figure 7(A,B)) (Abid Ali et al., 2016;Costa et al., 2011, 2014; Yuan et al., 2016). Thus, insteadof being directly involved in translocation per se, itwould seem that GINS and Cdc45 help shift the

126 M. W. PARKER ET AL.

structural disposition of the MCM2–7 AAAþ domainsinto productive conformations. Consistent with thisnotion, incorporation of Drosophila MCM2–7 into theCMG increases the vmax of ATP hydrolysis by over 300-fold and results in a 10-fold higher affinity for DNA,resulting in a dramatic increase in helicase activity (Ilveset al., 2010). Interestingly, in addition to participating inthe control of MCM2–7 ring status, both GINS (Boskovicet al., 2007; Ilves et al., 2010) and Cdc45 (Bruck &Kaplan, 2013; Krastanova et al., 2012; Szambowska et al.,2014) have been shown to bind DNA. Cdc45, which is acatalytically defunct homolog of the RecJ exonuclease(Sanchez-Pulido & Ponting, 2011; Simon et al., 2016),also appears to assist with capture of the leadingstrand during transient CMG gate opening

(Petojevic et al., 2015), thereby possibly playing a pro-tective role during replication fork stalling (Bruck &Kaplan, 2013).

Structural analysis of the CMG reveals the presenceof two distinct conformers that are, as observed for theMCM2–7 helicase core, differentiated by the relativepositioning of the gating subunits (Abid Ali et al., 2016;Yuan et al., 2016). As the MCM NTD is stabilized byextensive contacts with GINS and Cdc45, changeswithin the AAAþ ATPase ring appear primarily respon-sible for the alterations observed between the two con-formational states (Abid Ali et al., 2016; Costa et al.,2011, 2014; Yuan et al., 2016). Altering the position ofthe MCM2/5 gate appears to propagate structuralchanges around the MCM2–7 ring that result in

Figure 7. CMG helicase organization and dynamics. (A) MCM2–7 adopts a spiral, cracked-ring architecture with a discontinuitybetween MCM2 and MCM5. Upon incorporation into the CMG, Cdc45 and GINS convert the helicase into a planar form and sealoff the MCM2/5 gate (EM density maps¼ EMD-1835 and EMD-1833 for MCM2–7 and CMG, respectively). (B) A view of the CMGfrom the AAAþ face illustrating the overall architecture of the complex (PDB¼ 3JC5 (Yuan et al., 2016)). (C) At least two conform-ational states exist for the CMG, a constricted state in which the AAAþ and NTD rings are coplanar (bottom panel), and a relaxedstate where one end of the AAAþ tier lifts up from NTD tier (top panel). These conformations appear coupled to alterations inthe relative disposition of the gating subunits, MCM2 and MCM5 (constricted and relaxed conformer PDB codes¼ 3JC5 and 3JC7,respectively). (D) The activated CMG helicase is thought to translocate along single-stranded DNA, unwinding downstreamtemplate through a combined steric exclusion and DNA wrapping mechanism. A color version of this figure is available at www.tandfonline.com/ibmg.

CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY 127

significant translational and rotational rearrangement ofMCM6 and, to a lesser extent, MCM4. These changes inturn are coupled to a transition from a constricted con-former state, where the MCM2–7 NTD and CTD tiers areco-planar, to a relaxed, tilted form, which generates anasymmetric expansion of the gap between the NTD andCTD tiers on one side of the ring (Figure 7(C)). It hasbeen suggested that switching between these confor-mations could elicit a pumpjack motion, with the CTDtier rocking with respect to a stable NTD tier duringDNA translocation (Yuan et al., 2016), and that thesedynamics could be used to drive a linear, as opposed torotational, mechanism of DNA translocation (Abid Aliet al., 2016; Yuan et al., 2016). Higher-resolution struc-tures of distinct translocation intermediates, coupledwith single-molecule measurements of ring dynamicsand displacement, will be needed to test these ideas.

An interesting point of discussion in the field hasbeen whether the ATPase regions of MCMs serve as sin-gle or double-stranded DNA translocases. Based on adistant phylogenetic kinship to RuvB, a known double-strand translocase, MCM2–7 was initially proposed tomove along duplex DNA, with the NTD-associatedGINS/Cdc45 accessory subunits serving as a plowshareto separate the two strands (Takahashi et al., 2005).However, subsequent biochemical studies of theDrosophila CMG have shown that the complex requiresa fork for unwinding (Ilves et al., 2010), consistent withboth archaeal MCM and metazoan MCM4,6,7 function-ing as single-stranded DNA translocases on model sub-strates (Ishimi, 1997; Kang et al., 2012; Kelman et al.,1999; Moyer et al., 2006; Shechter et al., 2000). Alongthese lines, studies in archaea have reported that anMCM construct lacking the NTD tier (and that has noassociated GINS/Cdc45) still possesses helicase activity(Barry et al., 2007). Moreover, replisomes have beenchallenged with leading and lagging strand roadblocksin an X. laevis extract system, with only leading strandroadblocks proving capable of stalling fork progression(MCM2–7 is a 30!50 helicase that tracks on the leadingstrand) (Fu et al., 2011). Consistent with this observa-tion, recent analysis of the pattern of forked DNA cross-linking to the CMG reveals that the leading strandshows robust crosslinking to all MCM2–7 subunits, aswell as Cdc45, while lagging strand crosslinks are mark-edly fewer and those that occur are all on the exteriorsurface of MCM2–7 (Petojevic et al., 2015). Collectively,these data strongly indicate that the CMG functions asa single-stranded DNA translocase.

Structural analysis of the apo and substrate-boundCMG states also support a single-strand trackingmodel. In particular, upon binding of the CMG to aforked duplex substrate, nucleic acid density can be

visualized within the central channel whose dimen-sions appear sufficient to accommodate single butnot double-stranded DNA (Abid Ali et al., 2016). Thisfinding is consistent with a previous structural ana-lysis of the CMG bound to duplex DNA containing asingle 30-tail, which showed that the duplex region ofa DNA fork is positioned outside the central channelin an orientation consistent with the known 30!50

polarity of DNA engagement (Costa et al., 2014).Interestingly, the high-resolution structure of the S.cerevisiae CMG reveals that the WH domains of Mcm5and Mcm6, which are disordered within the contextof the double hexamer (Li et al., 2015), becomeordered and take up a position within the centralpore of the CMG that constricts the AAAþ pore to apoint where it is too small to accommodate double-stranded DNA (Yuan et al., 2016). Overall, these linesof evidence largely corroborate the view that theCMG tracks along single-stranded templates, albeitwith a potential capacity to switch to a double-stranded DNA mode of translocation, as might occurduring replication termination, when forks converge(Dewar et al., 2015) (parenthetically, there is evidencethat the E. coli DnaB helicase also can undergo atransition between single and double-stranded trans-location modes (Kaplan, 2000; Kaplan & O'Donnell,2002)). Understanding the role of the WH domainsawaits determination of substrate-bound CMG struc-tures in which the disposition of the nucleic acid sub-strate and the WH domains are fully defined.

Overall, while a variety of mechanistic models existto account for hexameric helicase activity, the so-called steric exclusion model appears most compat-ible with the observed data to date for the CMG. Thesteric exclusion model posits that translocation occursalong single-stranded DNA with the nontemplatestrand being excluded from the central channel. Oneassumption of this framework has been that the lag-ging strand is functionally passive, implying that oncedisplaced, it is inconsequential to helicase function.However, emerging lines of evidence are starting tosuggest that the lagging strand may wrap around theexternal surface of the CMG and support helicaseactivity (Figure 7(D)). Consistent with this notion, DNAinteractions with the external surface of archaealMCM is integral to helicase function (Costa et al.,2008; Graham et al., 2011, 2016; Rothenberg et al.,2007). DNA binding to the MCM external surfaceappears at least partially facilitated by the NTD-Aregion, which can undergo large structural rearrange-ments to expose a DNA-binding site (Chen et al.,2005; Costa et al., 2008; Fletcher et al., 2003; Milleret al., 2014). Whether the eukaryotic MCM NTD-A also

128 M. W. PARKER ET AL.

facilitates DNA binding is unclear; however, the MCMAAAþ domain contains a conserved, surface-exposedb–hairpin that has been reported to directly engageDNA (Brewster et al., 2010; Graham et al., 2011) and,in the context of the eukaryotic CMG, is integral tohelicase function (Petojevic et al., 2015). Thus, theCMG may function by a mechanism of single-strandedDNA translocation that results in both steric exclusionand the wrapping of lagging strand DNA (Grahamet al., 2011).

Regulatory mechanisms for preventingre-initiation

Origin licensing and firing are the most tightly regu-lated events in the process of DNA replication initi-ation. Multiple interdependent and redundantmechanisms ensure that origins can be marked andutilized only within a certain window of the cell cycle.While the previous discussion of helicase activationhas demonstrated the role of CDK and DDK in posi-tively regulating initiation, a plethora of mechanismsexist to negatively regulate replication licensing andprevent re-replication as well. This section will discusssome general principles for preventing re-replication;although regulatory approaches show substantialevolutionary diversification and expansion, certaincommon themes do exist. For a more thorough dis-cussion of the regulation of DNA replication initiationin eukaryotes, we point the reader to a number ofdedicated reviews on the topic (see (Arias & Walter,2007; Araki, 2010; Hook et al., 2007; Masai et al.,2010; Siddiqui et al., 2013)).

Initiator and helicase regulation

By temporally separating helicase loading from activa-tion, S-CDK can simultaneously activate the helicasewhile also inhibiting helicase-loading factors to preventorigin re-firing. A major means by which CDK protectsagainst re-licensing in S-phase is to regulate initiatorinteractions and their association with chromatin.Although budding yeast ORC remains stably boundthroughout the cell cycle (Diffley et al., 1994; Fujitaet al., 1998; Liang & Stillman, 1997), vertebrate Orc1 isthe target of CDK-dependent and CDK-independentmechanisms that remove it from chromatin (Figure 8)(Findeisen et al., 1999; Kreitz et al., 2001; Li &DePamphilis, 2002; Li et al., 2004; Natale et al., 2000;Rowles et al., 1999; Sun et al., 2002). In both yeastand metazoans, initiation factors are targeted for prote-olysis in a CDK-dependent manner, with buddingyeast exclusively targeting Cdc6 (Drury et al., 1997;

Mimura et al., 2004; Perkins et al., 2001; Piatti et al.,1995) and metazoans targeting both Orc1 and Cdc6(Kalfalah et al., 2015; Lidonnici et al., 2004; Mendezet al., 2002; Ohta et al., 2003; Tatsumi et al., 2003). In S.pombe, Orc2 is also targeted by CDK to prevent re-licensing, but the mechanism utilized in this instance isunclear (Vas et al., 2001; Wuarin et al., 2002). In S. cerevi-siae, CDK-dependent control is further extended toregulate Cdc6 transcription and nuclear localization(Honey & Futcher, 2007; Moll et al., 1991); metazoanCdc6 likewise undergoes CDK-dependent nuclearexport to limit replication (Jiang et al., 1999b; Petersenet al., 1999; Saha et al., 1998). Interestingly, buddingyeast Orc6 and Cdc6 stably associate with CDK duringS-phase and this interaction sterically inhibits interac-tions necessary for pre-RC assembly, such as with Cdt1(Chen & Bell, 2011; Mimura et al., 2004; Wilmes et al.,2004). In metazoans, Cdt1 has not yet been reported tointeract with Orc6.

In addition to regulating ORC and Cdc6, in S.cerevisiae CDK targets Mcm2–7 to prevent new, pro-ductive interactions with chromatin-bound ORC·Cdc6complexes. This action occurs through the CDK-dependent nuclear exclusion and export of Mcm2–7(Labib et al., 1999; Liku et al., 2005; Nguyen et al.,2001; Tanaka & Diffley, 2002). There is currently noevidence that CDK targets metazoan MCM2–7 directly,although phosphorylation of the initiator reduces

Figure 8. Mechanisms to prevent re-replication. Multiple,redundant mechanisms are utilized to prevent re-licensing oforigins after S-phase has initiated. Whereas yeast seems toexclusively utilize CDK-dependent mechanisms to preventre-licensing, metazoans also employ CDK-independent path-ways for negatively regulating Cdt1 activity. A color version ofthis figure is available at www.tandfonline.com/ibmg.

CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY 129

helicase association with origins (Findeisen et al.,1999). Interestingly, a new, CDK-independent MCMregulatory mechanism has recently been identified inbudding yeast that involves the SUMOylation of allmembers of the Mcm2–7 hexamer in G1. The SUMOmodification in turn, through phosphatase recruit-ment, prevents phosphorylation-dependent helicaseactivation and thus negatively regulates initiation(Wei & Zhao, 2016). How the SUMO pathway integra-tes with the other mechanisms that regulate initiationis currently unclear.

Cdt1: a master regulatory nexus

Cdt1 plays an essential part in the loading of MCM2–7onto DNA as a stable double hexamer. As such, Cdt1turns out to be a common regulatory point.Saccharomyces cerevisiae Cdt1 activity is restricted bythe CDK-dependent inhibition of its interaction withOrc6 (Chen & Bell, 2011) and by nuclear export of Cdt1in G1 (Tanaka & Diffley, 2002). Conversely, metazoanand S. pombe Cdt1 protein levels are regulated in acell-cycle-dependent manner such that Cdt1 isactively degraded upon S-phase entry (Figure 8)(Gopalakrishnan et al., 2001; Nishitani et al., 2000, 2001,2004; Wohlschlegel et al., 2000; Zhong et al., 2003).Degradation of Cdt1 is restricted to S-phase by an inter-action with chromatin-bound PCNA, which serves as aplatform for Cdt1 recognition and ubiquitination by theCullin-RING ligase 4 (CRL4) ubiquitin ligase (Arias &Walter, 2005, 2006; Guarino et al., 2011; Hu & Xiong,2006; Senga et al., 2006). A conserved PCNA-interactingprotein (PIP) degron within the Cdt1 N-terminus facili-tates the interaction with chromatin-bound PCNA and isrequired for degradation (Havens & Walter, 2009, 2011;Senga et al., 2006). Similarly, the CDK-dependent phos-phorylation of Cdt1 leads to its recognition and ubiqui-tination by the SCFSkp2 E3 ubiquitin ligase, providing anadditional mechanism to limit Cdt1 protein levels andprevent re-replication (Kondo et al., 2004; Li et al., 2003;Liu et al., 2004; Nishitani et al., 2006; Sugimoto et al.,2004; Thomer et al., 2004 ). Notably, Cdt1 ubiquitinationcan be reversed through the function of the ubiquitinhydrolase USP37, whose activity stabilizes Cdt1 and pro-motes helicase loading (Hernandez-Perez et al., 2016).

Metazoan Cdt1 is also uniquely regulated by bindingto a partner protein, Geminin. Geminin is a coiled-coilprotein (Lee et al., 2004; Saxena et al., 2004) that wasinitially identified in Xenopus egg extract screens forproteins destabilized in mitosis (McGarry & Kirschner,1998). This study, along with many others, revealed thatGeminin, a nuclear protein whose levels become ele-vated during S-phase, targets and restricts Cdt1 activity

to G1, thus preventing Cdt1-induced re-replication(Cook et al., 2004; Kerns et al., 2007; Lutzmann et al.,2006; Mihaylov et al., 2002; Quinn et al., 2001; Tadaet al., 2001; Yoshida et al., 2005). Geminin binds directlyto Cdt1 (De Marco et al., 2009; Lutzmann et al., 2006;Wohlschlegel et al., 2000), and this interaction has beenshown to inhibit Cdt1 binding to mouse MCM6 (Yanagiet al., 2002). Recent work demonstrates that Gemininbinding to Cdt1 inhibits ORC-dependent helicaseloading but not recruitment (Wu et al., 2014a), afinding consistent with studies in budding yeast show-ing Cdt1-independent recruitment of the helicase tothe origin-bound initiator (Fernandez-Cid et al., 2013;Frigola et al., 2013).

Interestingly, Geminin serves two seemingly dichot-omous functions. Although clearly an inhibitor of licens-ing, Geminin is also required for replication initiationthrough its ability to stabilize Cdt1 levels by protectingthe protein from degradation (Ballabeni et al., 2004;Narasimhachar & Coue, 2009). Thus, metazoans regulateGeminin protein levels and cellular localization to liber-ate Cdt1 from Geminin and promote pre-RC assemblyin G1 (Dimaki et al., 2013; McGarry & Kirschner, 1998;Tsunematsu et al., 2013). In addition, the stoichiometryof the Cdt1·Geminin complex has been suggested toregulate Geminin’s activity, such that a lower orderCdt1:Geminin complex (1:2) is permissive to pre-RCassembly whereas a higher order (2:4) complex is not(De Marco et al., 2009). How this stoichiometry is regu-lated in the context of the other regulatory mechanismsgoverning Geminin and Cdt1 protein levels and cellularlocalization is not known.

Concluding remarks

At this point, many of the major events and inter-mediates that facilitate pre-RC and pre-IC assemblyhave been defined. However, between these stableintermediates there exist multiple, undefined dynamicand transient protein interactions, modifications andexchanges that represent critical steps toward build-ing a replisome. The questions that remain are bothbroad and specific: how does ORC balance sequencepreference and trans-acting chromatin contextualcues when selecting origins? What is the role of Cdt1in MCM2–7 loading and, given the minimal conserva-tion of this protein, is Cdt1 mechanism conservedacross eukarya? Does MCM2–7 melt origins andunwind duplex DNA (as part of the CMG) using anoverlapping or mutually exclusive set of protein-nucleic acid interactions, and how is this functionalswitch regulated? What type of molecular dancemust take place to transition the pre-RC product, the

130 M. W. PARKER ET AL.

double-stranded DNA-bound MCM2–7double hexamer, into the CMG, and then again intoa bi-directional replication fork defined by two singlehelicases bound to single-stranded DNA? It will becritical to address these and other questions in mul-tiple model eukaryotic organisms to begin to under-stand what level of conservation can be expected forsuch a complex process. Future work that aims toresolve these and other questions will undoubtedlyreveal exciting new mechanisms that underlie thehighly orchestrated and regulated process of replica-tion initiation in eukaryotes.

Acknowledgements

The authors acknowledge Berger and Botchan lab memberspast and present for valuable discussion and helpful advicein preparing this review.

Disclosure statement

The authors report no conflicts of interest. The authors aloneare responsible for the content and writing of this article.

Funding

This work was supported by an NIH NRSA postdoctoral fel-lowship (F32GM116393, to MWP) and by the NCI(R01CA030490, to JMB and MRB).

ORCID

James M. Berger http://orcid.org/0000-0003-0666-1240

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