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Origins of DNA replication

Affiliation Quantitative Biology, Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland

* E-mail: [email protected]

• Babatunde Ekundayo, 
• Franziska Bleichert

Published: September 12, 2019

• https://doi.org/10.1371/journal.pgen.1008320
• Reader Comments

19 Dec 2019: The PLOS Genetics Staff (2019) Correction: Origins of DNA replication. PLOS Genetics 15(12): e1008556. https://doi.org/10.1371/journal.pgen.1008556 View correction

In all kingdoms of life, DNA is used to encode hereditary information. Propagation of the genetic material between generations requires timely and accurate duplication of DNA by semiconservative replication prior to cell division to ensure each daughter cell receives the full complement of chromosomes . DNA synthesis of daughter strands starts at discrete sites, termed replication origins, and proceeds in a bidirectional manner until all genomic DNA is replicated. Despite the fundamental nature of these events, organisms have evolved surprisingly divergent strategies that control replication onset. Here, we discuss commonalities and differences in replication origin organization and recognition in the three domains of life.

Citation: Ekundayo B, Bleichert F (2019) Origins of DNA replication. PLoS Genet 15(9): e1008320. https://doi.org/10.1371/journal.pgen.1008320

Copyright: © 2019 Ekundayo, Bleichert. This is an open access article distributed under the terms of the Creative Commons Attribution License , which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: This work was supported by the Novartis Research Foundation and the European Research Council under the European Union’s Horizon 2020 research and innovation program (ERC-STG-757909). The funders had no role in the preparation of the article.

Competing interests: The authors have declared that no competing interests exist.

Wikipedia Version : https://en.wikipedia.org/wiki/origins_of_dna_replication .

Introduction

In the second half of the 19th century, Gregor Mendel's pioneering work on the inheritance of traits in pea plants suggested that specific “factors” (today established as genes) are responsible for transferring organismal traits between generations [ 1 ]. Although proteins were initially assumed to serve as the hereditary material, Avery, MacLeod and McCarty established a century later DNA, which had been discovered by Friedrich Miescher , as the carrier of genetic information [ 2 ]. These findings paved the way for research uncovering the chemical nature of DNA and the rules for encoding genetic information, and ultimately led to the proposal of the double-helical structure of DNA by Watson and Crick [ 3 ]. This three-dimensional model of DNA illuminated potential mechanisms by which the genetic information could be copied in a semiconservative manner prior to cell division, a hypothesis that was later experimentally supported by Meselson and Stahl using isotope incorporation to distinguish parental from newly synthesized DNA [ 4 ][ 5 ]. The subsequent isolation of DNA polymerases, the enzymes that catalyze the synthesis of new DNA strands, by Kornberg and colleagues pioneered the identification of many different components of the biological DNA replication machinery, first in the bacterial model organism E . coli , but later also in eukaryotic life forms [ 6 ].

A key prerequisite for DNA replication is that it must occur with extremely high fidelity and efficiency exactly once per cell cycle to prevent the accumulation of genetic alterations with potentially deleterious consequences for cell survival and organismal viability [ 7 ]. Incomplete, erroneous, or untimely DNA replication events can give rise to mutations, chromosomal polyploidy or aneuploidy , and gene copy number variations, each of which in turn can lead to diseases, including cancer [ 8 ][ 9 ]. To ensure complete and accurate duplication of the entire genome and the correct flow of genetic information to progeny cells, all DNA replication events are not only tightly regulated with cell cycle cues but are also coordinated with other cellular events such as transcription and DNA repair [ 10 ][ 11 ][ 12 ].

DNA replication is divided into different stages ( Fig 1 ). During initiation, the replication machineries–termed replisomes–are assembled on DNA in a bidirectional fashion. These assembly loci constitute the start sites of DNA replication or replication origins. In the elongation phase, replisomes travel in opposite directions with the replication forks, unwinding the DNA helix and synthesizing complementary daughter DNA strands using both parental strands as templates. Once replication is complete, specific termination events lead to the disassembly of replisomes. As long as the entire genome is duplicated before cell division, one might assume that the location of replication start sites does not matter; yet, it has been shown that many organisms use preferred genomic regions as origins [ 13 ][ 14 ]. The necessity to regulate origin location likely arises from the need to coordinate DNA replication with other processes that act on the shared chromatin template to avoid DNA strand breaks and DNA damage [ 8 ][ 12 ][ 15 ][ 16 ][ 17 ][ 18 ][ 19 ].

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Models for bacterial ( A ) and eukaryotic ( B ) DNA replication initiation. A ) Circular bacterial chromosomes contain a cis -acting element, the replicator, that is located at or near replication origins. i ) The replicator recruits initiator proteins in a DNA sequence-specific manner, which results in melting of the DNA helix and loading of the replicative helicase onto each of the single DNA strands ( ii ). iii ) Assembled replisomes bidirectionally replicate DNA to yield two copies of the bacterial chromosome. B ) Linear eukaryotic chromosomes contain many replication origins. Initiator binding ( i ) facilitates replicative helicase loading ( ii ) onto duplex DNA to license origins. iii ) A subset of loaded helicases is activated for replisome assembly. Replication proceeds bidirectionally from origins and terminates when replication forks from adjacent active origins meet ( iv ).

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The replicon model

More than five decades ago, Jacob , Brenner , and Cuzin proposed the replicon hypothesis to explain the regulation of chromosomal DNA synthesis in E . coli [ 20 ]. The model postulates that a diffusible, trans -acting factor, a so-called initiator, interacts with a cis -acting DNA element, the replicator, to promote replication onset at a nearby origin ( Fig 1A , i ). Once bound to replicators, initiators (often with the help of co-loader proteins) deposit replicative helicases onto DNA, which subsequently drive the recruitment of additional replisome components and the assembly of the entire replication machinery ( Fig 1A , ii ). The replicator thereby specifies the location of replication initiation events, and the chromosome region that is replicated from a single origin or initiation event is defined as the replicon.

A fundamental feature of the replicon hypothesis is that it relies on positive regulation to control DNA replication onset, which can explain many experimental observations in bacterial and phage systems [ 20 ]. For example, it accounts for the failure of extrachromosomal DNAs without origins to replicate when introduced into host cells. It further rationalizes plasmid incompatibilities in E . coli , where certain plasmids destabilize each other’s inheritance due to competition for the same molecular initiation machinery [ 21 ]. By contrast, a model of negative regulation (analogous to the replicon-operator model for transcription) fails to explain the above findings [ 20 ]. Nonetheless, research subsequent to Jacob’s, Brenner’s and Cuzin’s proposal of the replicon model has discovered many additional layers of replication control in bacteria and eukaryotes that comprise both positive and negative regulatory elements, highlighting both the complexity and the importance of restricting DNA replication temporally and spatially [ 22 ][ 23 ][ 24 ].

The concept of the replicator as a genetic entity has proven very useful in the quest to identify replicator DNA sequences and initiator proteins in prokaryotes , and to some extent also in eukaryotes , although the organization and complexity of replicators differ considerably between the domains of life (for reviews, see [ 25 ][ 26 ]). While bacterial genomes typically contain a single replicator that is specified by consensus DNA sequence elements and that controls replication of the entire chromosome ( Fig 1A ), most eukaryotic replicators–with the exception of budding yeast–are not defined at the level of DNA sequence; instead, they appear to be specified combinatorially by local DNA structural and chromatin cues [ 27 ][ 28 ][ 29 ][ 30 ] [ 31 ][ 32 ][ 33 ][ 34 ][ 35 ][ 36 ]. Eukaryotic chromosomes are also much larger than their bacterial counterparts, raising the need for initiating DNA synthesis from many origins simultaneously to ensure timely replication of the entire genome ( Fig 1B ). Additionally, many more replicative helicases are loaded than activated to initiate replication in a given cell cycle ( Fig 1B ). The context-driven definition of replicators and selection of origins suggests a relaxed replicon model in eukaryotic systems that allows for flexibility in the DNA replication program [ 25 ]. Although replicators and origins can be spaced physically apart on chromosomes, they often co-localize or are located in close proximity; for simplicity, we will thus refer to both elements as ‘origins’ throughout this review. Taken together, the discovery and isolation of origin sequences in various organisms represents a significant milestone towards gaining mechanistic understanding of replication initiation. In addition, these accomplishments had profound biotechnological implications for the development of shuttle vectors that can be propagated in bacterial, yeast, and mammalian cells [ 37 ][ 38 ][ 39 ].

Bacterial replication origins

Most bacterial chromosomes are circular and contain a single origin of chromosomal replication ( oriC ). Bacterial oriC regions are surprisingly diverse in size (ranging from 250 bp to 2 kbp), sequence, and organization [ 41 ][ 42 ]; nonetheless, their ability to drive replication onset typically depends on sequence-specific readout of consensus DNA elements by the bacterial initiator, a protein called DnaA [ 43 ][ 44 ][ 45 ][ 46 ]. Origins in bacteria are either continuous or bipartite and contain three functional elements that control origin activity: conserved DNA repeats that are specifically recognized by DnaA (called DnaA-boxes), an AT-rich DNA unwinding element (DUE), and binding sites for proteins that help regulate replication initiation (for reviews, see [ 13 ][ 47 ][ 48 ]; Fig 2A ). Interactions of DnaA both with the double-stranded (ds) DnaA-box regions and with single-stranded (ss) DNA in the DUE are important for origin activation and are mediated by different domains in the initiator protein: a helix-turn-helix (HTH) DNA binding element and an ATPase associated with various cellular activities ( AAA+ ) domain, respectively ( Fig 2B ) [ 49 ][ 50 ][ 51 ][ 52 ][ 53 ][ 54 ][ 55 ][ 56 ]. While the sequence, number, and arrangement of origin-associated DnaA-boxes vary throughout the bacterial kingdom, their specific positioning and spacing in a given species are critical for oriC function and for productive initiation complex formation [ 41 ][ 42 ][ 57 ][ 58 ][ 59 ][ 60 ][ 61 ].

A ) Schematic of the architecture of E . coli origin oriC , Thermotoga maritima oriC , and the bipartite origin in Helicobacter pylori . The DUE is flanked on one side by several high- and weak-affinity DnaA-boxes as indicated for E . coli oriC . B ) Domain organization of the E . coli initiator DnaA. The magenta circle indicates the single-strand DNA binding site. C ) Models for origin recognition and melting by DnaA. In the two-state model (left panel), the DnaA protomers transition from a dsDNA binding mode (mediated by the HTH-domains recognizing DnaA-boxes) to an ssDNA binding mode (mediated by the AAA+ domains). In the loop-back model, the DNA is sharply bent backwards onto the DnaA filament (facilitated by the regulatory protein IHF [ 40 ]) so that a single protomer binds both duplex and single-stranded regions. In either instance, the DnaA filament melts the DNA duplex and stabilizes the initiation bubble prior to loading of the replicative helicase (DnaB in E . coli ). HTH–helix-turn-helix domain, DUE–DNA unwinding element, IHF–integration host factor.

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Among bacteria, E . coli is a particularly powerful model system to study the organization, recognition, and activation mechanism of replication origins. E . coli oriC comprises an approximately 260 bp region containing four types of initiator binding elements that differ in their affinities for DnaA and their dependencies on the co-factor ATP ( Fig 2A ). DnaA-boxes R1, R2, and R4 constitute high-affinity sites that are bound by the HTH domain of DnaA irrespective of the nucleotide-binding state of the initiator [ 43 ][ 62 ][ 63 ][ 64 ][ 65 ][ 66 ]. By contrast, the I, τ, and C-sites, which are interspersed between the R-sites, are low-affinity DnaA-boxes and associate preferentially with ATP-bound DnaA, although ADP-DnaA can substitute for ATP-DnaA under certain conditions [ 67 ][ 68 ][ 69 ][ 60 ]. Binding of the HTH domains to the high- and low-affinity DnaA recognition elements promotes ATP-dependent higher-order oligomerization of DnaA’s AAA+ modules into a right-handed filament that wraps duplex DNA around its outer surface, thereby generating superhelical torsion that facilitates melting of the adjacent AT-rich DUE ( Fig 2C ) [ 49 ][ 70 ][ 71 ][ 72 ]. DNA strand separation is additionally aided by direct interactions of DnaA’s AAA+ ATPase domain with triplet repeats, so-called DnaA-trios, in the proximal DUE region [ 73 ]. The engagement of single-stranded trinucleotide segments by the initiator filament stretches DNA and stabilizes the initiation bubble by preventing reannealing [ 53 ]. The DnaA-trio origin element is conserved in many bacterial species, indicating it is a key element for origin function [ 73 ]. After melting, the DUE provides an entry site for the E . coli replicative helicase DnaB, which is deposited onto each of the single DNA strands by its loader protein DnaC.

Although the different DNA binding activities of DnaA have been extensively studied biochemically and various apo , ssDNA-, or dsDNA-bound structures have been determined [ 52 ][ 53 ][ 54 ][ 71 ], the exact architecture of the higher-order DnaA- oriC initiation assembly remains unclear. Two models have been proposed to explain the organization of essential origin elements and DnaA-mediated oriC melting. The two-state model assumes a continuous DnaA filament that switches from a dsDNA binding mode (the organizing complex) to an ssDNA binding mode in the DUE (the melting complex) ( Fig 2C , left panel ) [ 71 ][ 74 ]. By contrast, in the loop-back model, the DNA is sharply bent in oriC and folds back onto the initiator filament so that DnaA protomers simultaneously engage double- and single-stranded DNA regions ( Fig 2C , right panel ) [ 75 ]. Elucidating how exactly oriC DNA is organized by DnaA remains thus an important task for future studies. Insights into initiation complex architecture will help explain not only how origin DNA is melted, but also how a replicative helicase is loaded directionally onto each of the exposed single DNA strands in the unwound DUE, and how these events are aided by interactions of the helicase with the initiator and specific loader proteins.

Archaeal replication origins

Archaeal replication origins share some but not all of the organizational features of bacterial oriC . Unlike bacteria, archaea often initiate replication from multiple origins per chromosome (one to four have been reported) [ 76 ][ 77 ][ 78 ][ 79 ][ 80 ][ 81 ][ 82 ] [ 83 ][ 42 ]; yet, archaeal origins also bear specialized sequence regions that control origin function (for recent reviews, see [ 84 ][ 85 ][ 86 ]). These elements include both DNA sequence-specific origin recognition boxes (ORBs or miniORBs) and an AT-rich DUE that is flanked by one or several ORB regions [ 82 ][ 87 ]. ORB elements display a considerable degree of diversity in terms of their number, arrangement, and sequence, both among different archaeal species and among different origins within in a single species [ 77 ][ 82 ][ 88 ]. An additional degree of complexity is introduced by the initiator, Orc1/Cdc6 in archaea, which binds to ORB regions. Archaeal genomes typically encode multiple paralogs of Orc1/Cdc6 that vary substantially in their affinities for distinct ORB elements and that differentially contribute to origin activities [ 82 ][ 89 ][ 90 ][ 91 ]. In Sulfolobus solfataricus , for example, three chromosomal origins have been mapped (oriC1, oriC2, and oriC3; Fig 3A ), and biochemical studies have revealed complex binding patterns of initiators at these sites ( Fig 3B ) [ 82 ][ 83 ][ 92 ][ 93 ]. The cognate initiator for oriC1 is Orc1-1, which associates with several ORBs at this origin [ 82 ][ 90 ]. OriC2 and oriC3 are bound by both Orc1-1 and Orc1-3 [ 82 ][ 90 ][ 93 ]. Conversely, a third paralog, Orc1-2, footprints at all three origins but has been postulated to negatively regulate replication initiation [ 82 ][ 93 ]. Additionally, the WhiP protein, an initiator unrelated to Orc1/Cdc6, has been shown to bind all origins as well and to drive origin activity of oriC3 in the closely related Sulfolobus islandicus [ 90 ][ 92 ]. Because archaeal origins often contain several adjacent ORB elements, multiple Orc1/Cdc6 paralogs can be simultaneously recruited to an origin and oligomerize in some instances [ 91 ][ 94 ]; however, in contrast to bacterial DnaA, formation of a higher-order initiator assembly does not appear to be a general prerequisite for origin function in the archaeal domain.

A ) The circular chromosome of Sulfolobus solfataricus contains three different origins. B ) Arrangement of initiator binding sites at two S . solfataricus origins, oriC1 and oriC2. Orc1-1 association with ORB elements is shown for oriC1. Recognition elements for additional Orc1/Cdc6 paralogs are also indicated, while WhiP binding sites have been omitted. C ) Domain architecture of archaeal Orc1/Cdc6 paralogs. The orientation of ORB elements at origins leads to directional binding of Orc1/Cdc6 and MCM loading in between opposing ORBs (in B ). (m)ORB–(mini-)origin recognition box, DUE–DNA unwinding element, WH–winged-helix domain.

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Structural studies have provided insights into how archaeal Orc1/Cdc6 recognizes ORB elements and remodels origin DNA [ 94 ][ 95 ]. Orc1/Cdc6 paralogs are two-domain proteins and are composed of a AAA+ ATPase module fused to a C-terminal winged-helix fold ( Fig 3C ) [ 96 ][ 97 ][ 98 ]. DNA-complexed structures of Orc1/Cdc6 revealed that ORBs are bound by an Orc1/Cdc6 monomer despite the presence of inverted repeat sequences within ORB elements [ 94 ][ 95 ]. Both the ATPase and winged-helix regions interact with the DNA duplex but contact the palindromic ORB repeat sequence asymmetrically, which orients Orc1/Cdc6 in a specific direction on the repeat [ 94 ][ 95 ]. Interestingly, the DUE-flanking ORB or miniORB elements often have opposite polarities [ 77 ][ 82 ][ 91 ][ 99 ][ 100 ], which predicts that the AAA+ lid subdomains and the winged-helix domains of Orc1/Cdc6 are positioned on either side of the DUE in a manner where they face each other ( Fig 3B , bottom panel ) [ 94 ][ 95 ]. Since both regions of Orc1/Cdc6 associate with the minichromosome maintenance (MCM) replicative helicase [ 101 ][ 102 ], this specific arrangement of ORB elements and Orc1/Cdc6 is likely important for loading two MCM complexes symmetrically onto the DUE ( Fig 3B ) [ 82 ]. Surprisingly, while the ORB DNA sequence determines the directionality of Orc1/Cdc6 binding, the initiator makes relatively few sequence-specific contacts with DNA [ 94 ][ 95 ]. However, Orc1/Cdc6 underwinds and bends DNA, suggesting that it relies on a mix of both DNA sequence and context-dependent DNA structural features to recognize origins [ 94 ][ 95 ][ 103 ]. Notably, base pairing is maintained in the distorted DNA duplex upon Orc1/Cdc6 binding in the crystal structures [ 94 ][ 95 ], whereas biochemical studies have yielded contradictory findings as to whether archaeal initiators can melt DNA similarly to bacterial DnaA [ 90 ][ 91 ][ 104 ]. Although the evolutionary kinship of archaeal and eukaryotic initiators and replicative helicases indicates that archaeal MCM is likely loaded onto duplex DNA (see next section), the temporal order of origin melting and helicase loading, as well as the mechanism for origin DNA melting, in archaeal systems remains therefore to be clearly established. Likewise, how exactly the MCM helicase is loaded onto DNA needs to be addressed in future studies.

Eukaryotic replication origins

Origin organization, specification, and activation in eukaryotes are more complex than in bacterial or archaeal kingdoms and significantly deviate from the paradigm established for prokaryotic replication initiation. The large genome sizes of eukaryotic cells, which range from 12 Mbp in S . cerevisiae to 3 Gbp in humans, necessitates that DNA replication starts at several hundred (in budding yeast) to tens of thousands (in humans) origins to complete DNA replication of all chromosomes during each cell cycle (for recent reviews, see [ 32 ][ 23 ]). With the exception of S . cerevisiae and related Saccharomycotina species, eukaryotic origins do not contain consensus DNA sequence elements but their location is influenced by contextual cues such as local DNA topology, DNA structural features, and chromatin environment [ 25 ][ 31 ][ 33 ]. Nonetheless, eukaryotic origin function still relies on a conserved initiator protein complex to load replicative helicases onto DNA during the late M and G1 phases of the cell cycle, a step known as origin licensing ( Fig 1B ). [ 107 ] In contrast to their bacterial counterparts, replicative helicases in eukaryotes are loaded onto origin duplex DNA in an inactive, double-hexameric form and only a subset of them (10–20% in mammalian cells) is activated during any given S phase, events that are referred to as origin firing ( Fig 1B ) [ 108 ][ 109 ][ 110 ]. The location of active eukaryotic origins is therefore determined on at least two different levels, origin licensing to mark all potential origins, and origin firing to select a subset that permits assembly of the replication machinery and initiation of DNA synthesis. The extra licensed origins serve as backup and are activated only upon slowing or stalling of nearby replication forks, ensuring that DNA replication can be completed when cells encounter replication stress [ 111 ][ 112 ]. Together, the excess of licensed origins and the tight cell cycle control of origin licensing and firing embody two important strategies to prevent under- and overreplication and to maintain the integrity of eukaryotic genomes.

Early studies in S . cerevisiae indicated that replication origins in eukaryotes might be recognized in a DNA-sequence-specific manner analogously to those in prokaryotes. In budding yeast, the search for genetic replicators lead to the identification of autonomously replicating sequences (ARS) that support efficient DNA replication initiation of extrachromosomal DNA [ 113 ][ 114 ][ 115 ]. These ARS regions are approximately 100–200 bp long and exhibit a multipartite organization, containing A, B1, B2, and sometimes B3 elements that together are essential for origin function ( Fig 4 ) [ 116 ][ 117 ]. The A element encompasses the conserved 11 bp ARS consensus sequence (ACS) [ 118 ][ 119 ], which, in conjunction with the B1 element, constitutes the primary binding site for the heterohexameric origin recognition complex (ORC), the eukaryotic replication initiator [ 120 ][ 121 ][ 122 ][ 123 ]. Within ORC, five subunits are predicated on conserved AAA+ ATPase and winged-helix folds and co-assemble into a pentameric ring that encircles DNA ( Fig 4 ) [ 123 ][ 124 ][ 125 ]. In budding yeast ORC, DNA binding elements in the ATPase and winged-helix domains, as well as adjacent basic patch regions in some of the ORC subunits, are positioned in the central pore of the ORC ring such that they aid the DNA-sequence-specific recognition of the ACS in an ATP-dependent manner [ 123 ][ 126 ]. By contrast, the roles of the B2 and B3 elements are less clear. The B2 region is similar to the ACS in sequence and has been suggested to function as a second ORC binding site under certain conditions, or as a binding site for the replicative helicase core [ 127 ][ 128 ][ 129 ][ 130 ][ 131 ]. Conversely, the B3 element recruits the transcription factor Abf1, albeit B3 is not found at all budding yeast origins and Abf1 binding does not appear to be strictly essential for origin function [ 116 ][ 132 ][ 133 ].

Specific DNA elements and epigenetic features involved in ORC recruitment and origin function are summarized for S . cerevisiae , S . pombe , and metazoan origins. A schematic of the ORC architecture is also shown, highlighting the arrangement of the AAA+ and winged-helix domains into a pentameric ring that encircles origin DNA. Ancillary domains of several ORC subunits involved in targeting ORC to chromosomes are included. Other regions in ORC subunits may also be involved in initiator recruitment, either by directly or indirectly associating with partner proteins. A few examples are listed. Note that the BAH domain in S . cerevisiae Orc1 binds nucleosomes [ 105 ] but does not recognize H4K20me2 [ 106 ]. BAH–bromo-adjacent homology domain, WH–winged-helix domain, TFIIB–transcription factor II B-like domain in Orc6, G4 –G quadruplex, OGRE–origin G-rich repeated element.

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Origin recognition in eukaryotes other than S . cerevisiae or its close relatives does not conform to the sequence-specific readout of conserved origin DNA elements. Pursuits to isolate specific chromosomal replicator sequences more generally in eukaryotic species, either genetically or by genome-wide mapping of initiator binding or replication start sites, have failed to identify clear consensus sequences at origins [ 134 ][ 135 ][ 136 ][ 137 ][ 138 ][ 139 ][ 140 ][ 141 ][ 142 ][ 143 ][ 144 ][ 145 ]. Thus, sequence-specific DNA-initiator interactions in budding yeast signify a specialized mode for origin recognition in this system rather than an archetypal mode for origin specification across the eukaryotic domain. Nonetheless, DNA replication does initiate at discrete sites that are not randomly distributed across eukaryotic genomes, arguing that alternative means determine the chromosomal location of origins in these systems. These mechanisms involve a complex interplay between DNA accessibility, nucleotide sequence skew (both AT-richness and CpG islands have been linked to origins), nucleosome positioning, epigenetic features, DNA topology and certain DNA structural features (e.g., G4 motifs), as well as regulatory proteins and transcriptional interference [ 13 ][ 14 ][ 30 ][ 31 ][ 33 ][ 146 ][ 147 ][ 139 ][ 148 ]. Importantly, origin properties vary not only between different origins in an organism and among species, but some can also change during development and cell differentiation. The chorion locus in Drosophila follicle cells constitutes a well-established example for spatial and developmental control of initiation events. This region undergoes DNA-replication-dependent gene amplification at a defined stage during oogenesis and relies on the timely and specific activation of chorion origins, which in turn is regulated by origin-specific cis-elements and several protein factors, including the Myb complex, E2F1, and E2F2 [ 149 ][ 150 ][ 151 ][ 152 ][ 153 ]. This combinatorial specification and multifactorial regulation of metazoan origins has complicated the identification of unifying features that determine the location of replication start sites across eukaryotes more generally.

To facilitate replication initiation, ORC assemblies from various species have evolved specialized auxiliary domains that are thought to aid initiator targeting to chromosomal origins or chromosomes in general ( Fig 4 ). For example, the Orc4 subunit in S . pombe ORC contains several AT-hooks that preferentially bind AT-rich DNA [ 154 ], while in metazoan ORC the TFIIB-like domain of Orc6 is thought to perform a similar function [ 155 ]. Metazoan Orc1 proteins also harbor a bromo-adjacent homology (BAH) domain that interacts with H4K20me2-nucleosomes [ 106 ]. Particularly in mammalian cells, H4K20 methylation has been reported to be required for efficient replication initiation, and the Orc1-BAH domain facilitates ORC association with chromosomes and Epstein-Barr virus origin-dependent replication [ 156 ][ 157 ][ 158 ][ 159 ][ 160 ]. Therefore, it is intriguing to speculate that both observations are mechanistically linked at least in a subset of metazoa, but this possibility needs to be further explored in future studies. In addition to the recognition of certain DNA or epigenetic features, ORC also associates directly or indirectly with several partner proteins that could aid initiator recruitment, including LRWD1, PHIP (or DCAF14), HMGA1a, among others ( Fig 4 ) [ 29 ][ 161 ][ 162 ][ 163 ][ 164 ][ 165 ][ 166 ][ 167 ]. Interestingly, Drosophila ORC, like its budding yeast counterpart, bends DNA and negative supercoiling has been reported to enhance DNA binding of this complex, suggesting that DNA topology and malleability might influence the location of ORC binding sites across metazoan genomes [ 27 ][ 123 ][ 168 ][ 169 ][ 170 ]. A molecular understanding for how ORC’s DNA binding regions might support the readout of structural properties of the DNA duplex in metazoans rather than of specific DNA sequences as in S . cerevisiae awaits high-resolution structural information of DNA-bound metazoan initiator assemblies. Likewise, how different epigenetic factors contribute to initiator recruitment in metazoan systems is poorly defined and is an important question that needs to be addressed in more detail.

Once recruited to origins, ORC and its co-factors Cdc6 and Cdt1 drive the deposition of the minichromosome maintenance 2–7 (Mcm2-7) complex onto DNA (for reviews see [ 107 ][ 171 ]). Like the archaeal replicative helicase core, Mcm2-7 is loaded as a head-to-head double hexamer onto DNA to license origins ( Fig 1B ) [ 108 ][ 109 ][ 110 ]. In S-phase, Dbf4-dependent kinase (DDK) and cyclin-dependent kinase (CDK) phosphorylate several Mcm2-7 subunits and additional initiation factors to promote the recruitment of the helicase co-activators Cdc45 and GINS, DNA melting, and ultimately bidirectional replisome assembly at a subset of the licensed origins ( Fig 1B ) [ 172 ][ 24 ]. In both yeast and metazoans, origins are free or depleted of nucleosomes, a property that is crucial for Mcm2-7 loading, indicating that chromatin state at origins regulates not only initiator recruitment but also helicase loading [ 140 ][ 173 ][ 174 ][ 175 ][ 176 ][ 177 ]. A permissive chromatin environment is further important for origin activation and has been implicated in regulating both origin efficiency and the timing of origin firing. Euchromatic origins typically contain active chromatin marks, replicate early, and are more efficient than late-replicating, heterochromatic origins, which conversely are characterized by repressive marks [ 23 ][ 175 ][ 178 ]. Not surprisingly, several chromatin remodelers and chromatin-modifying enzymes have been found to associate with origins and certain initiation factors [ 179 ][ 180 ], but how their activities impact different replication initiation events remains largely obscure. Remarkably, cis-acting “early replication control elements” (ERCEs) have recently also been identified to help regulate replication timing and to influence 3D genome architecture in mammalian cells [ 181 ]. Understanding the molecular and biochemical mechanisms that orchestrate this complex interplay between 3D genome organization, local and higher-order chromatin structure, and replication initiation is an exciting topic for further studies.

Why have metazoan replication origins diverged from the DNA sequence-specific recognition paradigm that determines replication start sites in prokaryotes and budding yeast? Observations that metazoan origins often co-localize with promoter regions in Drosophila and mammalian cells and that replication-transcription conflicts due to collisions of the underlying molecular machineries can lead to DNA damage suggest that proper coordination of transcription and replication is important for maintaining genome stability [ 135 ][ 137 ][ 139 ][ 142 ][ 182 ][ 16 ][ 17 ][ 19 ]. Recent findings also point to a more direct role of transcription in influencing the location of origins, either by inhibiting Mcm2-7 loading or by repositioning of loaded Mcm2-7 on chromosomes [ 183 ][ 148 ]. Sequence-independent (but not necessarily random) initiator binding to DNA additionally allows for flexibility in specifying helicase loading sites and, together with transcriptional interference and the variability in activation efficiencies of licensed origins, likely determines origin location and contributes to the co-regulation of DNA replication and transcriptional programs during development and cell fate transitions. Computational modeling of initiation events in S . pombe , as well as the identification of cell-type specific and developmentally-regulated origins in metazoans, are in agreement with this notion [ 136 ][ 144 ][ 184 ][ 185 ][ 186 ][ 187 ][ 188 ][ 148 ]. However, a large degree of flexibility in origin choice also exists among different cells within a single population [ 139 ][ 145 ][ 185 ], and the molecular mechanisms that lead to the heterogeneity in origin usage remain ill-defined. Mapping origins in single cells in metazoan systems and correlating these initiation events with single-cell gene expression and chromatin status will be important to elucidate whether origin choice is purely stochastic or controlled in a defined manner.

Concluding remarks

Although DNA replication is essential for genetic inheritance, defined, site-specific replication origins are technically not a requirement for genome duplication as long as all chromosomes are copied in their entirety to maintain gene copy numbers. Certain bacteriophages and viruses, for example, can initiate DNA replication by homologous recombination independent of dedicated origins [ 189 ]. Likewise, the archaeon Haloferax volcanii uses recombination-dependent initiation to duplicate its genome when its endogenous origins are deleted [ 78 ]. Similar non-canonical initiation events through break-induced or transcription-initiated replication have been reported in E . coli and S . cerevisiae [ 190 ][ 191 ][ 192 ][ 193 ][ 194 ]. Nonetheless, despite the ability of cells to sustain viability under these exceptional circumstances, origin-dependent initiation is a common strategy universally adopted across different domains of life. The controlled assembly of the replication machinery at origins likely confers long-term advantage to cells by allowing tight cell cycle regulation and by maintaining a specific replication dynamics. The divergent origin specification modes between prokaryotes and budding yeast on the one hand and metazoans on the other hand appear to reflect distinct needs to coordinate the spatiotemporal replication program with gene expression and cell differentiation programs to ensure not only genetic but also epigenetic inheritance and to preserve cell identity. Deciphering the underlying molecular mechanisms that modulate origin location, usage, and timing to define the replication program in metazoan systems represents an important major challenge in the field and will be essential to understand how dysregulation of these events are linked to human diseases. In addition, detailed studies of replication initiation have focused on a limited number of model systems. The extensively studied fungi and metazoa are both members of the opisthokont supergroup and exemplify only a small fraction of the evolutionary landscape in the eukaryotic domain [ 195 ]. Comparably few efforts have been directed at other eukaryotic model systems, such as kinetoplastids or tetrahymena [ 196 ][ 197 ][ 198 ][ 199 ][ 200 ] [ 201 ][ 202 ]. Surprisingly, these studies have revealed interesting differences both in origin properties and in initiator composition compared to yeast and metazoans. Further exploration of replication initiation mechanisms across different branches of the eukaryotic domain will likely yield unexpected insight into the diversity and evolution of this fundamental biological process.

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Microbe Notes

Prokaryotic DNA Replication- Enzymes, Steps and Significance

• DNA replication is the process by which an organism duplicates its DNA into another copy that is passed on to daughter cells.
• Replication occurs before a cell divides to ensure that both cells receive an exact copy of the parent’s genetic material. 
• DNA replication uses a semi-conservative method that results in a double-stranded DNA with one parental strand and a new daughter strand.
• Prokaryotic DNA replication is often studied in the model organism  coli , but all other prokaryotes show many similarities.

Table of Contents

Interesting Science Videos

Features of Prokaryotic DNA Replication

• Replication is bi-directional and originates at a single origin of replication (OriC).
• Takes place in the cell cytoplasm.
• Synthesis occurs only in the 5′to 3′direction.
• Individual strands of DNA are manufactured in different directions, producing a leading and a lagging strand.
• Lagging strands are created by the production of small DNA fragments called Okazaki fragments that are eventually joined together.

Enzymes of DNA Replication

• Helicases: Unwind the DNA helix at the start of replication.
• SSB proteins: Bind to the single strands of unwound DNA to prevent reformation of the DNA helix during replication.
• Primase: Synthesizes the RNA primer needed for the initiation of DNA chain synthesis.
• DNA Polymerase III (DNAP III): Elongates DNA strand by adding deoxyribonucleotides to the 3′end of the chain. Synthesis can only occur in the 5′to 3′direction because of DNAP III.
• DNA Polymerase I (DNAP I): Replaces RNA primer with the appropriate deoxynucleotides.
• DNA topoisomerase I: Relaxes the DNA helix during replication through creation of a nick in one of the DNA strands.
• DNA topoisomerase II: Relieves the strain on the DNA helix during replication by forming supercoils in the helix through the creation of nicks in both strands of DNA.
• DNA ligase: Forms a 3′-5′phosphodiester bond between adjacent fragments of DNA.

Steps of DNA Replication

• DNA replication begins at a specific spot on the DNA molecule called the origin of replication.
• At the origin, enzymes unwind the double helix making its components accessible for replication. 
• The helix is unwound by helicase to form a pair of replication forks.
• The unwound helix is stabilized by SSB proteins and DNA topoisomerases.
• Primase forms RNA primers (10 bases), which serve to initiate synthesis of both the leading and lagging strand.
• The leading strand is synthesized continuously in the 5′to 3′ direction by DNAP III.
• The lagging strand is synthesized discontinuously in the 5′to 3′ direction through the formation of Okazaki fragments.
• DNAP I remove the RNA primers and replace the existing gap with the appropriate deoxynucleotides.
• DNA ligase seals the breaks between the Okazaki fragments as well as around the primers to form continuous strands.

Proofreading:

• In bacteria, all three DNA polymerases (I, II and III) have the ability to proofread, using 3’ → 5’ exonuclease activity.
• When an incorrect base pair is recognized, DNA polymerase reverses its direction by one base pair of DNA and excises the mismatched base.
• Following base excision, the polymerase can re-insert the correct base and replication can continue.

Significance

• DNA replication is a fundamental genetic process that is essential for cell growth and division. 
• DNA replication involve the generation of a new molecule of nucleic acid, DNA, crucial for life.
• DNA replication is important for properly regulating the growth and division of cells. 
• It conserves the entire genome for the next generation.
• David Hames and Nigel Hooper (2005). Biochemistry. Third ed. Taylor & Francis Group: New York.
• Bailey, W. R., Scott, E. G., Finegold, S. M., & Baron, E. J. (1986). Bailey and Scott’s Diagnostic microbiology. St. Louis: Mosby.
• Madigan, M. T., Martinko, J. M., Bender, K. S., Buckley, D. H., & Stahl, D. A. (2015). Brock biology of microorganisms (Fourteenth edition.). Boston: Pearson.
• https://en.wikipedia.org/wiki/Proofreading_(biology)
• https://sciencing.com/comparing-contrasting-dna-replication-prokaryotes-eukaryotes-13739.html

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Sagar Aryal

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• Published: 24 December 2007

Mechanisms and functions of DNA mismatch repair

• Guo-Min Li 1  

Cell Research volume  18 ,  pages 85–98 ( 2008 ) Cite this article

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DNA mismatch repair (MMR) is a highly conserved biological pathway that plays a key role in maintaining genomic stability. The specificity of MMR is primarily for base-base mismatches and insertion/deletion mispairs generated during DNA replication and recombination. MMR also suppresses homeologous recombination and was recently shown to play a role in DNA damage signaling in eukaryotic cells. Escherichia coli MutS and MutL and their eukaryotic homologs, MutSα and MutLα, respectively, are key players in MMR-associated genome maintenance. Many other protein components that participate in various DNA metabolic pathways, such as PCNA and RPA, are also essential for MMR. Defects in MMR are associated with genome-wide instability, predisposition to certain types of cancer including hereditary non-polyposis colorectal cancer, resistance to certain chemotherapeutic agents, and abnormalities in meiosis and sterility in mammalian systems.

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Rad27 and Exo1 function in different excision pathways for mismatch repair in Saccharomyces cerevisiae

Bacterial DNA excision repair pathways

Distinct DNA repair pathways cause genomic instability at alternative DNA structures

Introduction.

DNA damage accumulates in cells over time as a result of exposure to exogenous chemicals and physical agents (i.e., benzo[ a ]pyrene, polychlorinated biphenyls, dioxin, cigarette smoke, asbestos, ultraviolet light, radon), as well as endogenous reactive metabolites including reactive oxygen and nitrogen species (ROS and NOS). Another source of DNA damage is errors that occur during normal DNA metabolism or aberrant DNA processing reactions, including DNA replication, recombination, and repair. Nucleotide misincorporation generates DNA base-base mismatches during DNA synthesis at variable rates, depending on many factors, including the specific DNA polymerases. In general, the replicative DNA polymerases have relatively high replication fidelity (see McCulloch and Kunkel, this issue), while translesion DNA polymerases, which specifically bypass sites of DNA damage, have lower replication fidelity (see Andersen et al. and Gan et al. in this issue). DNA damage, if unrepaired, has the potential to generate mutations in somatic or germline cells, which can alter cellular phenotype and cause dysfunction and disease. To prevent such deleterious effects and safeguard the integrity of the genome, cells possess multiple mechanisms to repair DNA damage and thus prevent mutations. One such system is the critical pathway known as DNA mismatch repair (MMR).

MMR corrects DNA mismatches generated during DNA replication, thereby preventing mutations from becoming permanent in dividing cells 1 , 2 , 3 . Because MMR reduces the number of replication-associated errors, defects in MMR increase the spontaneous mutation rate 4 . Inactivation of MMR in human cells is associated with hereditary and sporadic human cancers 1 , 3 , 5 , and the MMR system is required for cell cycle arrest and/or programmed cell death in response to certain types of DNA damage 6 , 7 . Thus, MMR plays a role in the DNA damage response pathway that eliminates severely damaged cells and prevents both mutagenesis in the short term and tumorigenesis in the long term.

The prototypical Escherichia coli MMR pathway has been extensively studied and is well characterized both biochemically and genetically. Thus, E. coli MMR is a useful and important framework for understanding eukaryotic MMR. In this review, the biochemistry and genetics of E. coli MMR will be described briefly by way of introduction, and the remainder of the discussion will focus on the cellular functions of MMR and their roles in cancer avoidance in mammalian cells. For areas of research on human MMR not discussed in this paper or an additional discussion of MMR in other species, readers are referred to the following excellent reviews: 1 , 2 , 3 , 8 , 9 , 10 , 11 .

Mechanism of mismatch correction

Dna mmr in e. coli.

E. coli MMR requires the following protein components: MutS, MutL, MutH, DNA helicase II (MutU/UvrD), four exonucleases (ExoI, ExoVII, ExoX, and RecJ), single-stranded DNA binding protein (SSB), DNA polymerase III holoenzyme, and DNA ligase 12 , 13 . MutS, MutL, and MutH initiate MMR and play specialized biological roles in MMR in E. coli .

MutS recognizes base-base mismatches and small nucleotide insertion/deletion (ID) mispairs, and thus MutS has been called the “mismatch recognition” protein 3 . MutS possesses intrinsic ATPase activity. High-resolution structures of MutS bound to DNA have been determined by X-ray crystallography 14 , 15 . These structures revealed that MutS binds to a mismatch as a homodimer. Interestingly, the mismatch-binding site is comprised of sequence-wise identical but structurally and functionally different domains from the two subunits, indicating asymmetry in the protein-DNA complex. Hence, the MutS homodimer acts as a virtual heterodimer when bound to a DNA mismatch. This characteristic is mimicked by eukaryotic MutS homologs (MSH), which function as heterodimers instead of homodimers (see below). MMR in E. coli is ATP-dependent, and requires the functional MutS ATPase.

MutL interacts physically with MutS, enhances mismatch recognition, and recruits and activates MutH. Defects in MutL completely inhibit MMR in E. coli . Despite the fact that a functional human MutL homolog, MutLα, possesses an endonuclease activity that is essential for mammalian MMR 16 , no hydrolytic activity has been detected in MutL. However, MutL may play a role as a molecular matchmaker that facilitates assembly of a functional MMR complex 3 , 17 , because it stimulates the loading and the processivity of helicase II (or UvrD) at the MMR initiation site 18 , 19 . Like MutS, MutL functions as a homodimer and possesses ATPase activity 20 . Mutations in the ATP-binding domain lead to a dominant negative mutator phenotype 21 . MutL mutants that are defective in ATP hydrolysis but proficient in ATP binding can activate MutH but cannot stimulate MutH in response to a mismatch or MutS, suggesting that ATP hydrolysis by MutL is essential for mediating the activation of MutH by MutS 22 . Recent studies show that MutL interacts physically with the clamp loader subunits of DNA polymerase III 23 , 24 , suggesting that MutL may promote binding of DNA polymerase III to MMR intermediates. These observations suggest that MMR is coupled with DNA replication.

In E. coli , DNA is methylated at the N6 position of adenine in dGATC sequences. In replicating DNA, the daughter strand is transiently unmethylated, and it is the presence of hemimethylated dGATC sequences that molecularly distinguishes the newly synthesized daughter strand from the parental DNA strand. In MMR, hemi-methylated dGATC sites determine the strand specificity of repair. MutH, which recognizes hemimethylated dGATC sequences, functions as a monomer and belongs to a family of type-II restriction endonucleases 25 , 26 . Upon its recruitment and activation by MutS and MutL in the presence of ATP, MutH specifically incises the unmethylated daughter strand of hemimethylated dGATC 3 , 22 , and this strand-specific nick provides the initiation site for mismatch-provoked excision.

The first step of the MMR pathway is binding of a MutS homodimer to the mismatch. Subsequently, a hemi-methylated dGATC site 5′ or 3′ to the mismatch is located and cleaved by the concerted action of MutS, MutL, MutH, and ATP. Three models have been proposed to address how mismatch binding by MutS leads to cleavage of the hemimethylated dGATC site (see the section of Unsolved Fundamental Problems in MMR for details). The strand-specific nick generated by MutH at hemimethylated dGATC is a starting point for excision of the mispaired base. In the presence of MutL, helicase II (UvrD) loads at the nick and unwinds the duplex from the nick towards the mismatch 18 , generating single-strand DNA, which is rapidly bound by single-stranded DNA-binding protein (SSB) and protected from nuclease attack 27 . Depending on the position of the strand break relative to the mismatch, ExoI or ExoX (3′→5′ exonuclease), or ExoVII or RecJ (5′→3′ exonuclease) excises the nicked strand from the nicked site (the dGATC site) up to and slightly past the mismatch. The resulting single-stranded gap undergoes repair DNA resynthesis and ligation by DNA polymerase III holoenzyme, SSB, and DNA ligase 3 .

These early studies on E. coli MMR demonstrate three key features of this important pathway: first, repair is strand specific (i.e., restricted to the newly synthesized DNA strand); second, repair is bi-directional, proceeding 5′→3′ or 3′→5′ from the nick to the site of the mismatch; and third, MMR has broad substrate specificity including base-base mismatches and small ID mispairs. All of these properties require functional MutS, MutL, and MutH. Because the mechanism of MMR is highly conserved throughout evolution, E. coli MMR is an excellent model for MMR in eukaryotic cells.

MMR in human cells

MMR is a highly conserved biological pathway with strong similarities between human MMR and prototypical E. coli MMR 2 , 3 . These similarities include substrate specificity, bidirectionality, and nick-directed strand specificity. The role of hemi-methylated dGATC sites as a signal for strand discrimination is not conserved from E. coli MMR to human MMR, but because the hemi-methylated dGATC site directs MutH-dependent nicking, and because human MMR is presumed to be nick-directed in vivo , both systems are thought to discriminate daughter and template strands using a strand-specific nick.

Several human MMR proteins have been identified based on their homology to E. coli MMR proteins ( Table 1 ). These include human homologs of MutS 28 , 29 , 30 , 31 , 32 , MutL 33 , 34 , 35 , 36 , EXO1 37 , 38 , 39 , single-strand DNA-binding protein RPA 27 , 40 , proliferating cellular nuclear antigen (PCNA) 41 , 42 , 43 , DNA polymerase δ (pol δ) 44 , and DNA ligase I 45 . Although E. coli MutS and MutL proteins are homodimers, human MutS and MutL homologues are heterodimers 32 , 34 , 46 . hMSH2 heterodimerizes with hMSH6 or hMSH3 to form hMutSα or hMutSβ, respectively, both of which are ATPases that play a critical role in mismatch recognition and initiation of repair 2 . hMutSα preferentially recognizes base-base mismatches and ID mispairs of 1 or 2 nucleotides, while hMutSβ preferentially recognizes larger ID mispairs. At least 4 human MutL homologs (hMLH1, hMLH3, hPMS1, and hPMS2) have been identified 33 , 35 , 36 , 47 . hMLH1 heterodimerizes with hPMS2, hPMS1, or hMLH3 to form hMutLα, hMutLβ, or hMutLγ, respectively 2 . hMutLα is required for MMR and hMutLγ plays a role in meiosis, but no specific biological role has been identified for hMutLβ 2 . hMutLα possesses an ATPase activity and defects in this activity inactivate MMR in human cells. In a reconstituted human MMR system, hMutLα regulates termination of mismatch-provoked excision 45 . Recent studies show that MutLα possesses a PCNA/replication factor C (RFC)-dependent endonuclease activity which plays a critical role in 3′ nick-directed MMR involving EXO1 16 .

PCNA interacts with MSH2 and MLH1 and is thought to play roles in the initiation and DNA resynthesis steps of MMR 41 , 43 . PCNA also interacts with MSH6 and MSH3 48 , 49 , 50 , 51 via a conserved PCNA interaction motif termed the PIP box 52 . It has been proposed that PCNA may help localize MutSα and MutSβ to mispairs in newly replicated DNA 53 , 54 . Although PCNA is absolutely required during 3′ nick-directed MMR, it is not essential during 5′ nick-directed MMR 55 . This observation might be explained by the fact that EXO1, a 5′→3′ exonuclease, is involved in both 5′ and 3′ directed MMR. Like PCNA, EXO1 also interacts with MSH2 and MLH1 37 , 38 , 39 , 56 , 57 , 58 . While EXO1 can readily carry out 5′ directed mismatch excision in the presence of MutSα or MutSβ and RPA 45 , 59 , its role in catalyzing 3′ nick-directed excision requires the MutLα endonuclease, which is activated by PCNA and RFC 16 , 60 . Although it has been suggested that EXO1 possesses a cryptic 3′→5′ exonuclease activity 60 , 61 , current data do not support that hypothesis. Instead, recent studies suggest the following steps for EXO1-catalyzed 3′ nick-directed repair: (1) after recognition of the 3′ nick and the mismatch, MutLα endonuclease makes an incision 5′ to the mismatch in a manner dependent on PCNA and RFC; and (2) EXO1 performs 5′→3′ excision from the MutLα-incision site through and beyond the site of the mismatch 16 . However, exo1 null mutants in mice and yeast have a weak mutator phenotype 56 , 62 ; thus, it is likely that additional as yet unidentified exonucleases are involved in eukaryotic MMR.

Other protein components involved in human MMR include single-strand DNA-binding protein RPA, RFC, high mobility group box 1 protein (HMGB1), and DNA pol δ. RPA seems to be involved in all stages of MMR: it binds to nicked heteroduplex DNA before MutSα and MutLα, stimulates mismatch-provoked excision, protects the ssDNA gapped region generated during excision, and facilitates DNA resynthesis 27 , 45 , 60 , 63 . Furthermore, RPA is phosphorylated after pol δ is recruited to the gapped DNA substrate. Recent studies indicate that phosphorylation reduces the affinity of RPA for DNA, that unphosphorylated RPA stimulates mismatch-provoked DNA excision more efficiently than phosphorylated RPA, and that phosphorylated RPA facilitates MMR-associated DNA resynthesis more efficiently than unphosphorylated RPA 63 . These results are consistent with the fact that a high-affinity RPA-DNA complex might be required to protect nascent ssDNA and to displace DNA-bound MutSα/MutLα 27 , 45 , while a lower-affinity RPA-DNA complex might facilitate DNA resynthesis by pol δ 63 . HMGB1 is a mismatch-binding protein and has a DNA-unwinding activity 64 , 65 , 66 . It interacts with MSH2 and MSH6 in vitro 67 . Recent studies show that HMGB1 can substitute for RPA in an in vitro reconstituted MMR system 45 . Additional studies are needed to precisely define the function of HMGB1 in MMR.

Unsolved fundamental problems in MMR

Despite great progress in identifying MMR proteins and genes and application of state-of-the-art biochemical and genetic approaches to analyze the mechanism of MMR in prokaryotic and eukaryotic cells, several key questions about this pathway remain unanswered. One of these questions concerns the mechanism by which MMR proteins facilitate the communication between two physically distant DNA sites: the mismatch and the strand discrimination signal. It is generally agreed that the strand discrimination signal is a strand-specific nick in both prokaryotic and eukaryotic cells (see above), although the source of the nicking activity, at least for the leading strand, is not known in eukaryotic cells. Previous studies have proposed several alternative models for this process, which can be classified into “ cis -” or “moving” and “ trans -” or “stationary” models ( Figure 1 ). The “stationary” model ( Figure 1 , right) proposes that interactions among MMR proteins induce DNA bending or looping that brings the two distant sites together, while MutS (or the MSH heterodimers, i.e. MutSα and MutSβ) remains bound at the mismatch 19 , 22 . In this model, the MutS (or MSH heterodimers) ATPase activity acts in a proofreading role to verify mismatch binding and authorize the downstream excision 22 . Support for the stationary model came from the following experiments. Junop et al. showed that recognition of a mismatch by MutS on a DNA molecule activated MutH cleavage of a GATC site located on a separate DNA molecule without a mismatch 22 . Consistent with this observation, a second study demonstrated that mismatch-provoked excision could be initiated when a biotin-streptavidin blockade was placed between the mismatch and pre-existing nick 68 . The “ cis ” or “moving” models suggest that MutS-MutL (or MutSα/β-MutLα) complexes load at a mismatch site and then move away from the site to search for the strand break, where exonucleases can be recruited to initiate excision.

Models for signaling downstream MMR events following mismatch recognition. A schematic diagram for signaling between the mismatch and the strand discrimination signal is shown. Here, a 5′ nick is the strand discrimination signal. Similar models apply for 3′ nick-directed MMR. The “stationary” or “ trans ” model (right) emphasizes that MutS or its homolog (MSH) proteins remain bound at the mismatch. It is the protein-protein interactions that induce DNA bending or looping that brings the two distant sites together. The two DNA sites can cooperate in a “ trans ” configuration. In two “ cis ” or “moving” models, one called the “translocation” model (left) and the other called the “molecular switch” or “sliding clamp” model (middle), it is hypothesized that the MSH proteins bind to the mismatch and then move away from the site to search for the strand discrimination signal. The translocation model suggests that ATP hydrolysis drives unidirectional movement of the MSH proteins, resulting in the formation of an α-like loop. In the molecular switch model (center), binding of an MSH protein (in its ADP-bound state) to the mismatch triggers an ADP to ATP exchange that promotes bi-directional sliding of the protein away from the mismatch, thereby emptying the mismatch site for an incoming MSH protein. Mismatch excision begins when an MSH protein reaches the strand break.

There are two moving models, one called the “translocation” model and the other called the “molecular switch” or “sliding clamp” model ( Figure 1 ). In the translocation model 69 , ATP reduces the mismatch-binding affinity of MutS or the MSH heterodimers, and ATP hydrolysis drives unidirectional translocation of MutS proteins along the DNA helix. DNA is threaded through the protein complex until the latter reaches a strand discrimination signal in either orientation, a process that forms a DNA loop ( Figure 1 , left). In the molecular switch model, MutS or the MSH heterodimer binds to mismatched DNA in an ADP-bound state. The mismatch binding by MutS or the MSH heterodimer triggers a conformational change that allows an ADP to ATP exchange, which promotes a second conformational change that allows MutS or the MSH heterodimers to form a sliding clamp 70 , 71 , 72 , 73 . In this model ( Figure 1 , middle), it is the binding of ATP, not ATP hydrolysis, that signals downstream events including formation of ternary complex with MutL (or MLH heterodimers) and sliding of the ternary complex from the mismatch to the strand break 70 , 71 , 72 , 73 .

Recent studies by Pluciennik and Modrich 74 argue in favor of a moving rather than a stationary mechanism, because their data demonstrate that a dsDNA break 75 or a protein “roadblock” between the mismatch and the nick inhibits in vitro MMR with recombinant E. coli proteins. It is not clear why two “roadblock” experiments 68 , 74 obtained distinct results. In a reconstituted human MMR reaction, Zhang et al. 45 show that multiple MutSα-MutLα complexes are essential for processing a single mismatch, providing evidence to support the molecular switch model. Additional studies are needed to address these unresolved questions about the molecular mechanism of MMR.

MMR mediates DNA damage signaling

Mmr deficiency and drug resistance.

DNA-damaging agents such as the alkylating agents N -methyl- N′ -nitro- N -nitrosoguanidine (MNNG), temozolomide, or procarbazine are cytotoxic agents that kill most of the replicating cells. Many cancer therapeutics are genotoxic and cytotoxic agents that induce apoptotic cell death. Interestingly, many cells that acquire resistance to such agents are deficient in MMR. For example, the human lymphoblastoid cell line MTI, which has a defect in hMSH6 , was derived by culturing TK6 cells in the presence of a high concentration of MNNG. The resulting MNNG-resistant MT1 cells are defective in strand-specific MMR 76 . Many human colorectal cancer cell lines are also resistant to alkylating agents and have associated defects in MMR. The causal relationship between drug resistance and MMR is demonstrated by the fact that hMLH1 -defective MNNG-resistant cells lose drug resistance when the hMLH1 defect is genetically complemented with wild-type hMLH1 on chromosome 3 77 . It has also been observed that defects in MSH2 and PMS2 confer resistance to alkylating agents (reviewed in 78 ). The mechanism by which MMR influences drug cytotoxicity is discussed further below.

Resistance to methotrexate (MTX) has also been associated with phenotypic changes in MMR in human cells. This occurs by the unusual mechanism of co-amplification of the human chromosomal region that encodes dihydrofolate reductase (DHFR, the target of MTX) and hMSH3 79 , 80 . Amplification of DHFR lowers sensitivity to MTX by overexpressing the target of the drug. However, overexpression of hMSH3 sequesters hMSH2 in the hMutSβ heterodimer, effectively preventing formation of the hMutSα (hMSH2/hMSH6) heterodimer, which leads to degradation of uncomplexed hMSH6, significant dysregulation of MMR and hypermutability 81 , 82 . Overexpression of DHFR combined with genome-wide hypermutability and defective MMR are likely responsible for the MTX resistance of HL60 and other tumor cells.

MMR proteins promote DNA damage-induced cell cycle arrest and apoptosis

Cell cycle arrest is an important mechanism for preventing DNA damage-induced genomic instability. A large number of studies have characterized the so-called G2 or S phase checkpoints, and identified proteins required for cell cycle arrest, including ATM, ATR, p53, p73, Chk1, and Chk2. However, it was a somewhat unexpected finding that hMutSα- and hMutLα-deficient cells are defective in cell cycle arrest in response to multiple types of DNA damaging agents 6 , 7 , 83 . While the molecular basis of this effect is not precisely known, it has been reported that MMR-deficient cells fail to phosphorylate p53 and p73 in response to DNA damage 84 , 85 . This implicates ATM, ATR, and/or c-Abl, because these kinases phosphorylate p53 and p73 during the response to DNA damage 85 , 86 . In support of this, it has been reported that hMutSα and hMutLα interact physically with ATM, ATR-ARTIP, c-Abl, and p73 in cells treated with DNA damaging agents/drugs 83 , 87 , 88 , 89 . These observations implicate hMutSα and hMutLα in a signaling cascade that leads from DNA damage to cell cycle arrest and/or apoptosis. They also at least in part explain the fact that drug-induced cytotoxicity is lost in MMR-deficient cells, as discussed above 6 . Very recently, EXO1 has been shown to be essential for upstream induction of DNA damage response, possibly by reducing ssDNA formation and recruiting RPA and ATR to the damage site 90 . It remains to be seen if MutSα and/or MutLα act to recruit EXO1 in DNA damage response as they do in MMR.

Two models have been proposed to describe the role of MMR in DNA damage signaling. The “futile DNA repair cycle” model ( Figure 2 , left) proposes that strand-specific MMR, which targets only newly replicated DNA, engages in a futile DNA repair cycle when it encounters DNA lesions in the template strand, and this futile cycling activates DNA damage signaling pathways to induce cell cycle arrest and apoptosis 6 . Support for this model came from both in vivo and in vitro experiments. Stojic et al. 86 showed that exposure to MNNG induces DNA breaks/gaps, cell cycle arrest, and persistent nuclear foci at sites of DNA damage. The DNA damage-associated repair foci contain both damage signaling and DNA repair proteins, including ATR, γ-H2AX, and RPA. York and Modrich 91 showed that nicked circular heteroduplex plasmid DNA containing a single O 6 -methylguanine (O 6 -me-G)-thymine (T) mispair cannot be repaired by the MMR system when the lesion (O 6 -me-G) and the nick are on opposite strands; this suggests a futile repair process. An alternative model, referred to as the direct signaling model ( Figure 2 , right), argues that hMutSα/hMutLα directly trigger DNA damage signaling by recruiting ATM or ATR/ARTIP to the lesion, which activates a checkpoint response. This model is supported by an elegant study from the Hsieh laboratory showing that ATR and ATRIP form a complex with MutSα/MutLα in the presence of O 6 -me-G/T, which activates the ATR kinase and phosphorylates Chk1 89 . Because mammalian MMR proteins interact with a broad spectrum of DNA lesions 6 , this model is consistent with the notion that MutSα/MutLα acts as a sensor for DNA damage in mammalian cells. Both models provide a reasonable explanation for decreased DNA damage-induced apoptotic signaling and increased drug resistance in MMR-deficient cells.

Models for MMR-dependent DNA damage signaling. The “futile DNA repair cycle” model (left) suggests that DNA adducts (solid black circle) induce misincorporation, which triggers the strand-specific MMR reaction. Since MMR only targets the newly synthesized strand for repair, the offending adduct in the template strand cannot be removed, and will provoke a new cycle of MMR upon repair resynthesis. Such a futile repair cycle persists and activates the ATR and/or ATM damage signaling network to promote cell cycle arrest and/or programmed cell death. The direct signaling model proposes that recognition of DNA adducts by MSH-MLH complexes allows the proteins to recruit ATR and/or ATM to the site, activating the downstream damage signaling.

Role of MMR in other DNA metabolic pathways

MMR proteins have also been implicated in homeologous recombination, immunoglobulin class switching, somatic hypermutation, interstrand-crosslink repair, and trinucleotide repeat (TNR) expansion. Homeologous recombination (recombination between related but non-identical DNA sequences) generates mispairs/heteroduplexes, and induces genomic instability via chromosomal translocations, deletions, or inversions 92 , 93 . The frequency of homeologous recombination is much lower than that of homologous recombination in normal cells, but the frequency of homeologous recombination is dramatically elevated in MMR-deficient cells, suggesting that MMR suppresses homeologous recombination 3 , 94 . MutS and MutL inhibit DNA strand exchange between divergent sequences in vitro , most likely by binding to the mismatches generated during strand exchange 95 . Recent studies in yeast reveal that suppression of homeologous recombination is mediated by MutSα and a RecQ family helicase, SGS1 96 , 97 . Consistent with this notion, yeast strains defective in sgs1 fail to suppress homeologous recombination 97 , 98 . It has been postulated that MutSα recruits SGS1 to DNA mismatches, where it unwinds the heteroduplex and blocks homeologous recombination 96 . Although suppression of homeologous recombination by MMR proteins in human cells is less well understood, two human SGS1 homologous proteins, BLM and RECQ1, interact with MutSα 99 , 100 . This suggests that a similar mechanism is used to suppress homeologous recombination in yeast and human cells.

Recent studies also implicate MMR in repair of inter-strand crosslinks (ICLs), in a process that involves protein components from homologous recombination, double-strand break repair, and nucleotide excision repair 101 , 102 , 103 . The precise nature of this involvement is not yet clear, and the specific MMR proteins that participate remain somewhat controversial. However, hMutSβ appears to directly bind ICLs in vitro 102 , and hMutLα interacts specifically with the helicase domain of Fanconi Anemia protein FANC-J to facilitate ICL repair 101 .

The studies discussed above suggest that MMR promotes genomic stability. However, during immunoglobulin class switching and somatic hypermutation, it appears that MMR proteins play a highly specialized role in promoting genetic variation. Immunoglobulin class switching and somatic hypermutation are mechanisms for increasing antibody diversity during antigen-stimulated B-cell differentiation. During this process, activation-induced cytidine deaminase (AID) deaminates cytosine residues to uracil, generating G:U mispairs, which can be recognized and processed by MMR 104 , 105 . However, during repair resynthesis, the high-fidelity replicative pol δ and ɛ are thought to be replaced by the translesion polymerases, which are error-prone and crucially introduce base substitutions and frameshift mutations 106 . Additionally, MMR proteins play an important role in class switch recombination, an event where the IgM constant region is substituted by downstream constant sequences. In this capacity, MMR proteins utilize strand breaks generated by uracil DNA glycosylase to repair the AID-induced G:U mispairs in a strand-indiscriminate manner, leading to double-strand DNA breaks. It is these breaks that stimulate class switch recombination 107 . B cells from mice deficient in MMR genes ( MSH2 , MSH6 , MLH1 , PMS2 , or EXO1 ) display a low level of somatic hypermutation and reduced class switch recombination 8 , 62 , 108 , 109 .

MMR proteins also promote TNR expansion, a phenomenon associated with a number of neurological disorders in humans, including Huntington's disease, myotonic dystrophy, and fragile X syndrome 110 (also see Kovtun and McMurray in this issue). TNRs such as (CAG) n form single-stranded DNA loop/hairpin structures in vitro 111 . Surprisingly, transgenic CAG repeats undergo expansion in wild-type, but not in knockout mice defective in MMR genes MSH2 and MSH3 112 , 113 , suggesting that the expansion mutations of the transgenic CAG repeats require functional MMR proteins MSH2 and MSH3. In vitro biochemical studies indeed show that MutSβ (the MSH2-MSH3 heterodimer) specifically binds to the (CAG) n hairpin structure 113 . One model proposes that MutSβ inhibits repair or resolution of the (CAG) n hairpin, thus stimulating (CAG) n expansion 113 . However, many of these studies were conducted using transgenic mouse models for TNR expansion. Thus, the role of MMR in human neurological diseases involving TNR expansion is at present unclear, and additional genetic and biochemical studies are needed to define the mechanism of TNR expansion in human cells.

MMR deficiency leads to cancer development

Mmr defects in hereditary non-polyposis colorectal cancer (hnpcc) and other cancers.

In the early 1990s, it was shown that HNPCC and some cases of sporadic colon cancer are caused by defects in human MMR 3 . For HNPCC, Kolodner and co-workers, and Vogelstein and co-workers independently identified germ-line mutations in hMSH2 at chromosome 2p16-p21 in HNPCC families 29 , 30 . Genetic analyses of HNPCC kindreds revealed a large increase in frequency of insertion and deletion mutations in simple repeat (microsatellite) sequences, a phenomenon known as microsatellite instability (MSI) 114 . MSI was also observed at lower incidence in sporadic colon cancers 114 , 115 , 116 . Additional studies showed 100- to 700-fold decreased stability of poly(GT) tracts in yeast strains with single or double knockouts in MSH2 , MLH1 , or PMS1 117 . Biochemical studies by Modrich and co-workers 118 and Kunkel and co-workers 119 also showed that extracts of MSI-positive tumor cells were severely defective in repair of base-base and ID mispairs. Thus, genetic and biochemical evidence converged to support the hypothesis that defective MMR plays a causal role in carcinogenesis leading to HNPCC, and strongly implicated such defects in some sporadic human cancers such as colorectal cancer.

The second locus linked to HNPCC was hMLH1 at 3p21-p23 33 , 36 , 120 . Furthermore, HNPCC has also been linked to mutations in two additional MutL homolog genes hPMS1 and hPMS2 , on 2q and 7p, respectively 35 . Defects in hMLH1 represent the majority of all HNPCC cases 35 , 36 , with mutations in hMSH2 accounting for a large fraction of all the remaining HNPCC cases for which a genetic defect has been identified. In contrast, germ-line mutations in hMSH3 have not yet been linked to HNPCC, and mutations in hPMS1 , hPMS2 , and hMSH6 are relatively rare in HNPCC patients 1 , 2 , 3 . These observations are consistent with the fact that hMSH2 and hMLH1 are essential MMR components, while hPMS1 , hPMS2 , hMSH6 , and hMSH3 play important but partially redundant and/or dispensable roles in MMR.

Genetic evidence described above shows that defects in MMR correlate with HNPCC, and biochemical studies provide compelling additional evidence for this hypothesis. In particular, cell lines from HNPCC patients and sporadic tumors with MSI are defective in strand-specific MMR, and these cells can be divided into at least two complementation groups, corresponding to hMutLα 34 and hMutSα 32 . Importantly, purified hMutLα or hMutSα specifically complements the biochemical defect in these cells. Similar evidence was obtained from chromosome or gene transfer experiments: expressions of the exogenous MMR genes complement the biochemical defect and stabilize simple repetitive sequences in the transfected cells (reviewed in 121 ).

These genetic and biochemical complementation studies essentially prove the causal role of MMR defects in HNPCC. However, many questions remain, including whether or not mutations in hMLH3 or hEXOI also cause HNPCC, a possibility that remains controversial 47 , 122 , 123 . Mice defective in EXOI exhibit a mild defect in genomic stability, are partially defective in strand-specific MMR, and have increased rates of some cancers 62 . Thus, it seems possible that germ-line mutations in hEXOI could potentially have similar phenotypic effects in humans. Additional studies are needed to clarify this point.

Although MSI was first correlated with MMR defects in tumors from HNPCC patients, MSI is also associated with a wide variety of non-HNPCC and non-colonic tumors (reviewed in Ref. 124 ). These include endometrial, ovarian, gastric, cervical, breast, skin, lung, prostate, and bladder tumors as well as glioma, leukemia, and lymphoma. Biochemical studies confirmed that MSI cell lines from sporadic leukemia, endometrial, ovarian, prostate, and bladder cancers are defective in strand-specific MMR 32 , 125 , 126 . Interestingly, MSI in sporadic non-colonic tumors is often associated with hypermethylation of the promoter of hMLH1 (see below for details), and few mutations in MMR genes have been identified in these cells. These findings suggest that MMR defects are a likely cause of MSI in non-colonic sporadic cancers, although other mechanisms may also be involved in causing the MSI mutator phenotype.

Mouse models for MMR demonstrate roles in cancer and meiosis

Knockout mouse models have been developed for MSH2 , MSH3 , MSH6 , MLH1 , MLH3 , PMS1 , PMS2 , and EXO1 (reviewed in 8 , 62 , 127 , 128 ) and their phenotypes have been somewhat informative. Most of the knockout mice have a mutator phenotype, are MSI-positive, and are cancer-prone. However, the primary cancer susceptibility of MSH2 , MLH1 , and PMS2 knockout mice is lymphoma, not colorectal cancer as in humans, and secondary cancer susceptibilities are to gastrointestinal tumors, skin neoplasms, and/or sarcomas ( Table 2 ).

MSH2 −/− deficient mice are fertile 129 , 130 , are MSI-positive, develop lymphoma within 1 year of age, and have a significantly shorter lifespan than wild-type mice (i.e., 50% mortality by 6 months of age). The phenotype of MSH6 −/− deficient mice is similar to that of MSH2 −/− deficient mice, but lacking MSI 129 , 131 , a phenotype resembling that of atypical HNPCC with an hMSH6 defect as the tumors in these MSH6 -defective individuals have longer latency and low MSI 132 . Cells from MSH3 −/− mice are defective in repair of ID mispairs but can repair base-base mismatches. MSH3 −/− mice are either tumor free 133 or develop tumors at a very late age 134 , essentially consistent with the fact that no MSH3 mutations have been identified in HNPCC patients. However, in MSH3 −/− and MSH6 −/− double deficient mice, the tumor predisposition phenotype is indistinguishable from MSH2 −/− or MLH1 −/− mice 133 , 134 , suggesting that MSH3 cooperates with MSH6 in tumor suppression.

Sterility is a characteristic feature of MLH mutant mice (except PMS1 ) 135 , 136 , 137 . These animals are also susceptible to cancer and display genomic instability, reflecting defective MMR. However, male and female MLH1 and MLH3 knockout mice 135 , 136 , 137 , and male PMS2 knockout mice are completely sterile 138 . PMS1 knockout mice are exceptional, because they are fertile, they lack cancer susceptibility, and, apart from a very small increase in mutations in mononucleotide repeats, they appear to be MSI-negative 127 . EXO1 defective mice are also sterile 62 . It is clear that the loss of fertility in these knockout mice is caused by abnormal meiosis 62 , 128 , 135 , 136 , 137 .

The characteristics of all MMR knockout mouse models are summarized in Table 2 . These data strongly support the ideas that MMR is a basic genome surveillance mechanism and that defects in MMR can promote cancer development. The effects of MMR defects on carcinogenesis appear to be tissue- and species-specific, in a manner that is poorly understood. The effects of MMR defects on meiosis in humans remain poorly characterized. However, MMR clearly plays a critical role during meiosis and/or gamete formation in mice.

Epigenetic silencing of MMR gene expression leads to cancers

Mutations in MMR genes cause genomic instability and MSI in HNPCC and in a subset of sporadic colorectal cancers. However, in a significant fraction of MSI-positive sporadic colon tumors that have an MMR defect, mutations have not been identified in MMR genes. Epigenetic silencing of hMLH1 via promoter hypermethylation strongly down-regulates MMR in many of these cases 139 , 140 . In contrast, hypermethylation of the hMSH2 gene is rarely observed in tumors with MSI. In fact, it has been reported that more than 95% of MSI-H sporadic tumors demonstrate mutation and/or epigenetic silencing of hMLH1 141 . While most studies demonstrate epigenetic silencing of the hMLH1 promoter in sporadic tumors, hypermethylation of the hMLH1 promoter was also recently demonstrated in an HNPCC patient who does not have a germ-line mutation in any MMR gene 142 . Interestingly, recent studies suggest that this effect may be heritable 143 , 144 , 145 . Direct evidence that hMLH1 promoter hypermethylation down-regulates hMLH1 gene expression was obtained by treating cells with 5-aza-deoxycytidine, which reversed promoter hypermethylation, and restored hMLH1 gene expression and normal MMR capacity 146 , 147 .

MMR deficiency and mutations in coding repeat sequences

Previous studies demonstrate that defects in MMR increase the mutation rate in genes containing a simple repeat sequence in coding regions, often referred to as target genes 148 . Thus, defects in MMR confer a mutator phenotype. It is presumed that such a mutator phenotype has genome-wide consequences and could increase the frequency of additional genome-destabilizing and cancer-promoting mutations; however, this is a difficult hypothesis to test experimentally. One approach is to selectively analyze the stability of di- and tri-nucleotide tracts within coding regions. For example, Markowitz et al. 149 reported two mutation “hotspots” in the type II transforming growth factor-β receptor ( TGF-β RII ) gene in MMR-deficient tumor cells from a patient with sporadic colorectal cancer. One of these mutational hotspots fell within a 6-bp GT dinucleotide repeat and the other fell within an (A) 10 mononucleotide repeat 149 . Both of these hotspots were sites of frequent frameshift mutations that truncated the TGF-β RII gene product. Similar observations have been made in other colorectal tumor cells and many other MSI-positive tumor cells including glioma, gastric, uterine, cervical and squamous head and neck tumors, as well as ulcerative colitis-associated cancer and cecum cancer. Furthermore, in some of these tumor cells, somatic frameshift mutations have been documented in many other genes including Bax , insulin-like growth factor 2 receptor ( IGF2-R ), transcription factor E2F-4 , APC , PTEN , hMSH3 , hMSH6 , Mre11 , MBD4/MED , ACTRII , AIM2 , APAF-1 , AXIN-2 , BCL-10 , BLM , Caspase-5 , CDX-2 , CHK-1 , FAS , GRB-14 , cell cycle protein hG4-1 , KIAA0977 , ubiquinone oxidoreductase gene NADH , OGT , Rad50 , RHAMM , RIZ , SEC63 , SLC23AT , TCF-4 , and WISP-3 (reviewed in Ref. 150 ). These data are consistent with the idea that similar mutations occur on a genome-wide basis and at a much higher rate in MMR-deficient cells than in wild-type cells. Because the genes noted above play critical roles in regulating cell growth or genomic stability, loss-of-function mutations in these genes may be crucial steps in the multi-step pathway of carcinogenesis.

Conclusion and perspectives

The discovery that defects in MMR play a causal role in HNPCC and many MSI-positive sporadic cancers brought immediate clinical relevance to research in the field of eukaryotic MMR. This discovery led to intensive research on and better understanding of the biological roles of MMR in eukaryotic cells, which relate to cancer prevention and therapy. Although the primary role of MMR is to improve replication fidelity by correcting replication-associated base-base mismatches or ID mispairs, important secondary roles are to modulate DNA recombination and facilitate DNA damage signaling. Thus, it is abundantly clear that defects in MMR are “permissive” for carcinogenesis.

The identification that MMR-deficient cells are resistant to certain chemotherapeutic drugs such as temozolomide, procarbazine, or cisplatin has significant impacts on cancer treatments, especially for patients with tumors defective in MMR. It is also known that MMR deficiency can be acquired during chemotherapy by selective mutations in MMR genes 6 . Therefore, the risks for chemotherapy are two-fold. First, for patients with MMR-deficient tumors, chemotherapeutic treatments could selectively kill patients' MMR-proficient cells (e.g., blood cells) that undergo proliferation, thereby leading to rapid deaths of the cancer patients. Second, if a patient's tumor is not caused by loss of MMR function, chemotherapeutic treatments may kill the tumor cells; at the same time, the treatment may induce or select for a mutation in MMR genes, which could lead to a secondary cancer. Therefore, novel chemotherapeutic or alternative approaches are needed for cancer patients with or without MSI-positive tumors. Such approaches might include targeted gene therapy, which could selectively restore drug sensitivity in tumor cells defective in MMR or treatment with agents that stimulate apoptosis downstream of MMR in tumor cells. Additional research on mechanisms that selectively kill MMR-deficient cells is also warranted. Such efforts should also include more basic research on the molecular mechanisms of eukaryotic MMR. Understanding these mechanisms will support efforts for developing new therapeutic approaches for patients with HNPCC or other MSI-positive MMR-deficient tumors.

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Acknowledgements

The author acknowledges research support from the National Institutes of Health (GM072756 and CA115942) and the Kentucky Lung Cancer Research Program, USA. The author regrets the lack of citations for many important observations mentioned in the text, but their omission is made necessary by restrictions in the preparation of this review. The author holds the James-Gardner Endowed Chair in Cancer Research.

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Li, GM. Mechanisms and functions of DNA mismatch repair. Cell Res 18 , 85–98 (2008). https://doi.org/10.1038/cr.2007.115

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Published : 24 December 2007

Issue Date : January 2008

DOI : https://doi.org/10.1038/cr.2007.115

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Control of Eukaryotic DNA Replication Initiation—Mechanisms to Ensure Smooth Transitions

DNA replication differs from most other processes in biology in that any error will irreversibly change the nature of the cellular progeny. DNA replication initiation, therefore, is exquisitely controlled. Deregulation of this control can result in over-replication characterized by repeated initiation events at the same replication origin. Over-replication induces DNA damage and causes genomic instability. The principal mechanism counteracting over-replication in eukaryotes is a division of replication initiation into two steps—licensing and firing—which are temporally separated and occur at distinct cell cycle phases. Here, we review this temporal replication control with a specific focus on mechanisms ensuring the faultless transition between licensing and firing phases.

1. Introduction

DNA replication control occurs with exceptional accuracy to keep genetic information stable over as many as 10 16 cell divisions (estimations based on [ 1 ]) during, for example, an average human lifespan. A fundamental part of the DNA replication control system is dedicated to ensure that the genome is replicated exactly once per cell cycle. If this control falters, deregulated replication initiation occurs, which leads to parts of the genome becoming replicated more than once per cell cycle (reviewed in [ 2 , 3 , 4 ]). This over-replication (also referred to as rereplication) is accompanied by DNA damage, for example, in the form of double-strand breaks and subsequent genome rearrangements [ 5 , 6 , 7 , 8 ]. Notably, deregulation of DNA replication is increasingly recognized as a critical factor for genome instability during cancer development [ 9 , 10 , 11 , 12 , 13 ] and many oncogenes act by abrogating the G1/S cell cycle transition, thereby influencing DNA replication control [ 14 , 15 ]. Therefore, it seems plausible that over-replication could be a source of the genome rearrangements observed in tumour cells [ 16 ], but whether over-replication is a major driver of carcinogenesis is a question for future research.

Previous research has given us a detailed picture of the principal regulation of DNA replication initiation (see below) in the yeast Saccharomyces cerevisiae and also in metazoan systems. Replication initiation occurs in two interconnected steps—licensing and firing—which are coupled to separate phases of the cell cycle. Experimental systems to entirely abolish this separation cause widespread over-replication, a highly toxic condition. It is still a matter of active research as to how mutual exclusivity of licensing and firing is maintained at cell cycle transitions and, thus, how cells are protected from sporadic over-replication at these transitions. With this review, we aim to highlight established and also putative mechanisms that might act to ensure robust separation of licensing and firing and thus robustly block over-replication. We refer readers to the following excellent reviews for a detailed overview of the mechanism of replication initiation [ 2 , 17 , 18 ], elongation [ 18 , 19 ], and termination [ 18 , 20 , 21 ], as well as replication fork stalling [ 22 , 23 , 24 ].

2. DNA Replication Initiation in Eukaryotes

In eukaryotes, DNA replication initiates at many sites within the genome (replication origins) in parallel to allow fast duplication of large genomes. This brings about a need for tight control of initiation in order to ensure that each part of the genome is replicated exactly once per cell cycle. Cells achieve “once-per-cell-cycle replication initiation” by dividing the replication initiation process into two temporally separate phases—licensing and firing [ 2 , 3 ]. In mechanistic terms, licensing corresponds to the loading of inactive precursors of the Mcm2–7 helicase at replication origins by the pre-replicative complex ([ 25 , 26 , 27 , 28 , 29 ], Figure 1 A, upper panel), while firing corresponds to activation of the replicative helicase by association of additional accessory subunits ([ 30 , 31 , 32 , 33 , 34 , 35 , 36 ], Figure 1 A, lower panel). Previous studies have revealed the essential licensing and firing factors of budding yeast, and an in vitro reconstitution of origin-dependent initiation of replication has been achieved using the corresponding set of purified proteins [ 30 , 37 , 38 , 39 , 40 ]. In brief, licensing involves the licensing factors ORC (origin recognition complex Orc1–6), Cdc6, and Mcm2–7/Cdt1 and achieves origin recognition and ATP-dependent loading of the Mcm2–7 helicase core in the form of an inactive double hexamer, which encircles double-stranded DNA and is positioned in a head-to-head orientation, thus establishing bidirectionality of DNA replication ( Figure 1 A, [ 25 , 26 , 27 , 28 , 29 , 41 , 42 , 43 , 44 , 45 , 46 , 47 ]). Firing involves the helicase accessory subunits Cdc45 and GINS; the firing factors Sld2, Sld3, and Dpb11, as well as DNA polymerase ε and Mcm10 and achieves association of Cdc45 and GINS with Mcm2–7 and, thereby, activation of the replicative CMG helicase ( C dc45 M cm2–7 G INS), remodeling of the helicase to encircle single-stranded DNA (the leading strand template), and initial DNA unwinding [ 36 , 37 , 48 , 49 , 50 , 51 , 52 , 53 , 54 , 55 , 56 ]. After this committed step of initiation, multiple replication factors such as DNA polymerases associate with the replicative CMG helicase to catalyze chromosome replication [ 18 , 19 ]. Notably, firing and licensing factors are conserved from yeast to human [ 57 ], suggesting that not only the principal mechanism of replication initiation is highly conserved during evolution, but also that these conserved factors will most likely be essential targets of control.

Two-step mechanism of DNA replication initiation. ( A ) Inactive helicase precursors are loaded during origin licensing (upper panel); CDK and DDK promote activation of these precursors to form active CMG helicases during origin firing (lower panel). In addition to the depicted factors, origin firing and helicase activation involve Sld7, DNA polymerase ε, and Mcm10, which are indicated as additional factors. ( B ) Changing activity of CDK and DDK couples licensing and firing strictly to distinct phases of the cell cycle.

2.1. DNA Replication Initiation Control in Budding Yeast

Eukaryotic DNA replication initiates at multiple origins spread across the genome in order to allow a fast S phase despite large genomes. Features that define replication origins differ between species and have been comprehensively reviewed elsewhere [ 58 ]. Usage of multiple initiation sites inevitably brings with it the need for coordination. In particular, eukaryotic DNA replication control serves the purpose of generating a complete copy of the genome while avoiding any form of over-replication. Therefore, the two steps of initiation are interconnected (firing requiring prior licensing) but coupled to separate cell cycle phases, ensuring that every origin initiates at maximum once per cell cycle. Moreover, Mcm2–7 helicase precursors (the product of the licensing reaction) are removed from an origin when this origin is passively replicated [ 25 , 59 , 60 ], ensuring that origin firing cannot occur on post-replicative chromatin.

Temporal separation of licensing and firing, therefore, is key for ensuring that DNA replication at a given origin occurs only once per cell cycle. Indeed, when licensing and firing are experimentally induced to occur simultaneously, successive rounds of licensing and firing reactions trigger over-replication [ 5 ]. Temporal separation of licensing and firing is achieved by coupling them to specific phases of the cell cycle. Licensing generally occurs from late M phase to the G1/S transition [ 28 , 61 , 62 ]. Firing occurs in S phase, but the cellular firing potential (at least of budding yeast cells) remains high even until the metaphase-to-anaphase transition [ 31 ]. The next paragraphs focus on the well-understood cell cycle regulation of replication initiation in budding yeast, which is facilitated by the two major cell cycle kinases cyclin-dependent kinase 1 (Cdk1, also Cdc28, in the following referred to as CDK) and Dbf4-dependent kinase (DDK). Both kinases are regulated by the controlled expression and, importantly, destruction of regulatory subunits (cyclins and Dbf4, respectively) and are active from the G1/S transition to the metaphase-to-anaphase transition [ 63 , 64 , 65 , 66 , 67 , 68 , 69 , 70 , 71 ].

Firing essentially requires the activity of both CDK and DDK [ 72 , 73 , 74 ] and, indeed, many replication factors have been shown to be substrates of CDK and/or DDK phosphorylation [ 75 , 76 , 77 , 78 , 79 , 80 ]. Most importantly, the essential CDK and DDK substrates in replication initiation of budding yeast have been identified and specific mutants have been generated which bypass the CDK- and DDK-dependent steps, thus giving critical mechanistic insights into firing activation [ 48 , 49 , 51 , 81 , 82 , 83 ]. CDK phosphorylates Sld2 and Sld3, and this phosphorylation generates binding sites for Dpb11 [ 48 , 49 , 51 , 84 , 85 ]. Notably, Dpb11 contains two separate phosphoprotein binding sites [ 86 , 87 , 88 , 89 , 90 , 91 , 92 , 93 ], enabling it to simultaneously interact with Sld2 and Sld3 [ 48 , 49 , 84 ]. Although we currently lack insights into the precise architecture of the assembly of firing factors with the Mcm2–7 helicase, it is clear that the CDK-mediated interactions of Sld2, Sld3, and Dpb11 are critical to facilitate the association of the helicase accessory factors Cdc45 and GINS with Mcm2–7 and, thereby, the formation of the CMG helicase [ 84 , 94 , 95 ]. In particular, Cdc45 was suggested to be recruited in the form of a Sld3–Cdc45 complex [ 94 , 95 , 96 , 97 ], while GINS was suggested to be reeled in via Sld2 [ 84 , 98 ]. DDK, on the other hand, targets Mcm2–7 itself. DDK phosphorylation appears to not only relieve the Mcm2–7 complex from autoinhibition [ 81 , 82 , 83 ], it also appears to generate a Sld3-binding site on the Mcm2–7 complex [ 97 ]. Overall, the dependency on CDK and DDK therefore strictly links firing to S, G2, and early M phases [ 31 ].

Licensing, on the other hand, is inhibited by CDK phosphorylation of licensing factors. CDK phosphorylates ORC (Orc2 and Orc6 in particular), and this phosphorylation leads to inhibition of ORC [ 5 , 99 ]. CDK phosphorylates Cdc6 and the phosphorylation marks generate a degron on Cdc6, which facilitates destruction of Cdc6 by the ubiquitin–proteasome system [ 100 , 101 , 102 , 103 ]. CDK also phosphorylates the Mcm2–7 complex (in particular, Mcm3), which leads to the nuclear export of the soluble Mcm2–7/Cdt1 complex [ 104 ], with nuclear degradation of Mcm3 seemingly playing a backup role [ 105 ].

Furthermore, CDK also inhibits licensing in phosphorylation-independent ways. In particular, CDK was shown to engage in inhibitory protein–protein interactions with both Cdc6 (Clb2–Cdk1) and ORC [ 106 , 107 , 108 ]. The inhibition of the licensing reaction by CDK therefore restricts licensing to the cellular state of CDK inactivity from late mitosis to the G1/S transition.

2.2. Additional DNA Replication Initiation Control Mechanisms in Metazoa

The principles of replication control are highly similar in different eukaryotes and, at this point, it appears certain that division of replication initiation into separate licensing and firing phases occurring in G1 and S is a universal feature of eukaryotes. However, the specific mechanisms restricting licensing and firing to different phases of the cell cycle appear to be much less conserved compared to the core mechanisms of DNA replication itself [ 57 , 109 ]. As such, it is not surprising that metazoa—while apparently still using the CDK- and DDK-dependent control of licensing and firing [ 110 , 111 , 112 , 113 , 114 ]—have developed additional mechanisms which are independent of phosphorylation events. Interestingly, at least two of these mechanisms impinge on the licensing factor Cdt1, making it a focal point of regulation.

The first mechanism involves the licensing inhibitor geminin, orthologs of which can be found in worms, insects, and vertebrates [ 115 , 116 , 117 , 118 , 119 ]. Geminin from Xenopus laevis binds to the licensing factor Cdt1, thereby sequestering it from its licensing role to promote Mcm2–7 loading [ 115 , 116 ]. Geminin is, in principle, CDK-independent but nonetheless under strict cell cycle control, giving cell cycle phase specificity to this mechanism of licensing inhibition. Geminin is an ubiquitylation substrate of APC Cdh1 and is degraded in late M and G1 [ 117 ]. Conversely, the presence of geminin leads to sequestration of Cdt1 from S to early M and to geminin-dependent suppression of licensing during this time. Therefore, this mechanism contributes to restricting licensing to late M and G1 as well as ensuring mutual exclusivity of licensing and firing.

A second example of licensing inhibition is Cdt1 degradation by the Cul4–Cdt2 ubiquitin ligase complex [ 120 , 121 , 122 , 123 , 124 ]. This mechanism is found in metazoa [ 120 , 121 ] as well as Schizosaccharomyces pombe [ 125 , 126 ] and is indirectly coupled to the cell cycle, as it is dependent on ongoing DNA replication. Specifically, Cul4–Cdt2 requires association with the replication elongation factor PCNA [ 122 , 123 , 124 ], which thereby functions as a replication-coupled platform for Cdt1 destruction and, thereby, licensing inhibition. Per definition, this mechanism will only be active when DNA replication elongation is ongoing. As such, it is suited to keep Cdt1 levels low during S phase, thereby contributing to licensing and firing separation, but it is insufficient to establish separation of licensing and firing on its own.

2.3. Deregulation of DNA Replication Initiation—Over-Replication and Genome Instability

The principles of replication control are highly similar in different eukaryotes, and temporal separation of replication initiation into two phases appears to be a universal feature of eukaryotic replication. In order to understand the consequences of deregulated replication initiation, several experimental systems have been developed. In particular, budding yeast cells as well as Xenopus laevis egg extracts have been insightful models and have collectively shown that a temporal overlap of licensing and firing leads to over-replication (see below). Over-replication, in turn, will cause DNA damage in the form of double-strand breaks (DSBs) and induce genome instability [ 7 , 8 , 127 , 128 ].

In budding yeast, the best-understood system to induce widespread over-replication utilizes mutants that bypass CDK-dependent controls of licensing, which otherwise block this process in S, G2, and early M. Specifically, CDK-dependent inhibition of licensing is abolished by (i) mutation of the CDK phosphorylation sites on Orc2 and Orc6, (ii) overexpression of a degradation-resistant, truncated version of Cdc6, and (iii) expression of Mcm7 with a constitutive nuclear localization signal [ 5 ]. Continued expression of these inhibition-resistant licensing factors is lethal for cells, underlining the importance of temporal separation of licensing and firing, but conditional systems have allowed studying over-replication by inducing licensing and over-replication in M or S phase cells, respectively [ 5 , 129 ]. Specific induction of over-replication in a single cell cycle using this system or derivatives, where only a subset of factors has been deregulated, has allowed initial insights into the consequences of over-replication [ 6 , 129 , 130 , 131 , 132 ]. From this work, we can conclude that over-replication is a potent inducer of genome instability, which manifests as chromosome rearrangements, gene amplifications, chromosomal instability, and even aneuploidy [ 6 , 8 , 130 , 132 , 133 ]. A major source of this genome instability appear to be DSBs in the over-replicated region which, given the inherent overamplification of genomic material, can only be repaired in a manner that involves rearrangements and/or duplications of chromosomal sequences. Currently, we cannot, however, say by which specific mechanism these DSBs are induced and whether other DNA lesions or structures contribute significantly to genome instability after over-replication, and we refer readers to excellent reviews dedicated to this topic [ 134 , 135 ].

Deregulation of licensing has also been the key to study the consequences of widespread over-replication in metazoan systems, in particular using the cell-free Xenopus laevis egg extract system. Strategies have involved depletion of geminin and inhibition of Cdt1 proteolysis [ 120 , 121 , 136 , 137 ] and exogenous addition of Cdt1 at high levels [ 136 , 138 ] to replicating extracts, as well as depletion of geminin [ 139 , 140 , 141 ] or the mitotic regulator Emi1 [ 128 , 142 ] in cultured cells. All systems lead to over-replication and also to the induction of DSBs, which trigger a cell cycle arrest that is dependent on the DNA damage checkpoint [ 128 , 141 , 143 ]. Collectively, these studies have also put forward the idea that the frequency of over-replication initiation events may be an important factor influencing the number of induced DSBs, but perhaps also the mechanism by which they are induced.

2.4. Partial Deregulation of DNA Replication Initiation—Sporadic Over-Replication

While systems to induce widespread over-replication have been immensely informative, it can be argued for at least two reasons that sporadic over-replication is perhaps an even more relevant scenario in the context of cellular physiology: (i) Sporadic over-replication will not be as toxic as widespread over-replication. Not only will fewer over-replication events generate less DNA damage, less DNA damage will also not be as efficient in inducing the DNA damage checkpoint and its associated protective functions. Therefore, sporadic over-replication might manifest primarily in the form of genome instability and could be a contributing factor during tumorigenesis, when cells face deregulation of critical G1/S cell cycle controls [ 9 , 11 , 14 , 15 ]. (ii) A remarkable feature of DNA replication control is its multi-layered character; for example, CDK independently inhibits several licensing factors in budding yeast [ 5 ]. It is conceivable that such synergizing pathways confer overall robustness and, indeed, it has been suggested that overlapping pathways are required to give each replication origin the immense accuracy of replication initiation, which is statistically required for genome replication to be overall accurate [ 144 ]. Disabling just one layer of regulation will therefore generally only result in sporadic over-replication, but mutations disabling just one layer of control will be much more frequent compared to mutations disabling the entire control system.

While experimental systems to induce sporadic over-replication by partial deregulation of licensing [ 6 ] or firing factors [ 96 , 145 ] have been explored, our understanding of sporadic over-replication is limited by the lack of sensitive methods to analyze such events. While genetic and sequencing-based analyses of genome rearrangements are extremely sensitive, genome rearrangements can have many sources and are not uniquely linked to over-replication [ 146 ]. In contrast, commonly used physical assays are unable to visualize sporadic over-replication [ 5 , 129 , 147 ], and development of more sensitive methods will be a key future task for the field. Most importantly, our inability to specifically measure sporadic over-replication also means that we are currently unable to predict whether the principal set of replication control mechanisms has been identified or not. In the following section, we explore what kind of additional mechanisms ensuring a temporal separation of licensing and firing exist on top of the principal CDK/DDK control of replication initiation. In particular, we focus on the transitions between licensing and firing phases and on mechanisms that act at these cell cycle transitions in order to ensure robust separation of licensing and firing and, thereby, once-per-cell-cycle replication.

3. DNA Replication Control at Cell Cycle Transitions

In order to prevent sporadic over-replication, cells need to avoid any scenario in which even sub-pools of licensing and firing factors are simultaneously active. As such, it is interesting to consider how such a scenario is avoided at cell cycle transitions and whether the principal cell cycle control mechanisms at these transitions are sufficient to guarantee mutual exclusivity or whether additional replication-specific mechanisms are employed. Importantly, the transitions between the two replication phases coincide with the two major cell cycle transitions, where cells switch between CDK-on and CDK-off states.

3.1. Bistable Switches—The Fundament of DNA Replication Control at Cell Cycle Transitions

The problem of accurately separating events of one cell cycle phase from those in the next cell cycle phase is by no means a problem unique to DNA replication control. Indeed, sharp (ultrasensitive) and unidirectional cell cycle transitions are the very fundament of the cell cycle [ 148 ]. In a simplified model of the cell cycle, cells exist in one of two stable states—CDK-on and CDK-off—and the transitions between these two states are extremely rapid (switch-like). Progress in quantitative measurements of cell cycle factors combined with mathematical modeling shows that both the G1/S transition, during which cells switch to the CDK-on state and activate firing, as well as the metaphase-to-anaphase transition, during which cells switch to the CDK-off state and activate licensing, take the form of bistable switches [ 149 , 150 , 151 ]. These cell cycle transitions are also characterized by hysteresis and, thereby, are unidirectional and irreversible. On a molecular level, critical mechanisms that cause bistability and hysteresis are (i) positive feedback (specifically, double-negative feedback loops involving inhibitors of CDK (G1/S) or the APC (metaphase to anaphase) [ 152 , 153 , 154 , 155 , 156 ], (ii) complex, irreversible molecular processes such as protein degradation or translation, and (iii) post-translational modifications of key proteins occurring at multiple sites [ 157 ].

It is therefore logical to assume that the sharp and irreversible rise (G1/S) and drop (metaphase to anaphase) of CDK activity is the principal mechanism that causes rapid activation/inactivation of licensing and firing factors and thereby counteracts over-replication at cell cycle transitions. Indeed, a shallower rise of CDK activity at the G1/S transition (in budding yeast sic1–cpd mutants) coincides with enhanced chromosome loss [ 157 ], which can be interpreted to be a consequence of sporadic over-replication at the G1/S transition. Notwithstanding these initial insights, further experimental systems to manipulate the nature of cell cycle transitions as well as mathematical modeling are required to verify that DNA replication control critically requires sharp cell cycle transitions.

3.2. Temporal Order of Licensing/Firing Activation/Inactivation at Cell Cycle Transitions

With cell cycle regulators undergoing switch-like transitions, one can speculate that the activation and inactivation of licensing and firing factors would also be switch-like ( Figure 2 A). With switch-like licensing–firing transitions, it would, for example, be possible to simultaneously activate firing and inactivate licensing at G1/S without risking over-replication. At this point, we can, however, not exclude the possibility that the regulation of replication control factors will be more gradual, depending, for example, on slow turnover of proteins or their modifications ( Figure 2 B). With gradual licensing–firing transitions, a simultaneous activation/inactivation could potentially involve licensing and firing factors being (partially) active at the same time ( Figure 2 B), therefore potentially allowing sporadic over-replication.

Separation of licensing and firing at cell cycle phase transitions. Three different models for transitions between replication phases are depicted here for the example of the G1/S transition. Note the temporal overlap of licensing (grey) and firing (blue) activity in the different models, indicating the potential for sporadic over-replication.

Currently, there is little evidence supporting either of the two models. There is, however, mounting evidence that activation/inactivation of licensing/firing factors does not occur simultaneously, but that it is ordered ( Figure 2 C). This order appears to follow the general “inactivate first, activate second” rule, which means that at the G1/S transition licensing is turned off before firing is activated and that at the metaphase-to-anaphase transition firing is turned off before licensing is activated. This control will therefore result in temporal gaps between licensing and firing phases, which could be a mechanism to provide a robust block to over-replication. In the following section, we summarize our knowledge of mechanisms ( Figure 3 ) potentially generating such temporal order. It needs to be pointed out, however, that at this stage, we have only isolated pieces of evidence for individual mechanisms, primarily from studies in budding yeast and the Xenopus system. Therefore, while a general picture is emerging, we cannot yet comment on the conservation of individual mechanisms.

Mechanisms preventing over-replication. Summary of molecular mechanisms inhibiting licensing or firing at specific cell cycle phases (CDK-off or CDK-on) and specifically at the G1/S transition and the metaphase-to-anaphase transition. These inhibitory mechanisms are likely candidates to generate a temporal order in the activation/inactivation of licensing and firing factors and to thereby counteract sporadic over-replication at cell cycle transitions and limit replication to once per cell cycle.

3.3. Intrinsic Temporal Order by CDK–Substrate Interactions

Differential affinities of CDK to its substrates involved in licensing and firing could lead to those substrates becoming phosphorylated at different kinase concentrations. With rising CDK activity, such a mechanism could lead to a temporal offset in the inactivation of licensing factors and the activation of firing factors. Notably, both licensing and firing factors are also subject to multisite phosphorylation [ 5 , 48 , 49 , 77 , 78 , 85 ]. It is therefore easily conceivable that different thresholds are generated dependent on (i) the number of sites needed for activation/inactivation, (ii) the affinity of CDK for the individual sites, and (iii) the overall affinity for the substrate. Indeed, a mathematical model of replication initiation in budding yeast suggests that multisite phosphorylation of Sld2 (together with differential G1-CDK and S-CDK phosphorylation, see below) could cause licensing to become inactivated before firing becomes activated, thereby generating a temporal gap between the two processes [ 158 ].

An additional feature of CDK control that is utilized to generate order in the replication program is the fact that CDK is not a single entity, but that different cyclin–CDK complexes are active (to quantitatively different degrees) at different points throughout the cell cycle. Indeed, some budding yeast licensing factors are already targeted and inactivated by G1/S-CDK [ 101 , 102 , 104 ], while firing factors are only targeted and activated by S-CDK and not by G1/S-CDK [ 48 , 49 , 51 , 145 ]. G1/S-CDK activation precedes and is required for S-CDK activation, and mathematical modeling therefore suggests that the differential substrate specificity of G1/S-CDK and S-CDK for licensing and firing factors (together with effects by multisite phosphorylation, see above) will induce a delay in firing activation with respect to licensing inactivation [ 158 ]. Overall, these data suggest a temporal order at the G1/S transition in budding yeast, whereby licensing factors are inactivated first and firing factors are activated second.

Differential specificity of different CDK complexes could also play a role at the metaphase-to-anaphase transition. While in budding yeast most licensing factors are better substrates for S-CDK (Clb5–Cdk1) than for M-CDK (Clb2–Cdk1), Mcm3 seems not to follow this general trend and to be very efficiently phosphorylated also by M-CDK [ 159 ]. Firing could therefore become inactivated as soon as levels of S-CDK drop in M phase, while licensing inhibition (via Mcm3 phosphorylation and Orc6 phosphorylation (see next paragraph)) is maintained as long as M-CDK is active, consistent with temporal order and the “inactivate first, activate second” model.

3.4. Temporal Order by Phosphatase–Substrate Interactions

At the metaphase-to-anaphase transition, replication initiation factors become dephosphorylated, which for firing factors corresponds to inactivation and for licensing factors corresponds to activation. While dephosphorylation could in principle also be achieved by protein turnover (see below), most replication factors appear to be actively dephosphorylated by protein phosphatases. It is therefore conceivable that phosphatases play a role in determining a temporal order of firing inactivation and licensing activation. One mitotic phosphatase that was shown to act on replication factors (Orc2, Orc6, Mcm3, Sld2, and Sld3) is budding yeast Cdc14 [ 145 , 160 , 161 ], which is a specific antagonist of CDK phosphorylation function during mitotic exit [ 162 ]. Notably, it was shown that Cdc14 has differential catalytic efficiency for different substrates, which results in temporally ordered dephosphorylation [ 163 ]. Interestingly, Orc6 dephosphorylation occurred late compared to other Cdc14 substrates [ 163 ], apparently generating a window of opportunity for prior dephosphorylation of firing factors. It needs to be noted that even after Cdc14 inactivation, the temporal order of dephosphorylation of firing factors (first) and licensing factors (second) at the metaphase-to-anaphase transition remained largely intact [ 145 ]. While Cdc14 may therefore be a factor contributing to a temporal order of dephosphorylation at the metaphase-to-anaphase transition, it is not the only mechanism. Other phosphatases acting redundantly with Cdc14 or degradative mechanisms (see below) are likely candidates.

3.5. Temporal Order by Degradation

Cell cycle-regulated degradation is a commonly used mechanism to inactivate licensing factors (e.g., budding yeast Cdc6 [ 100 , 101 , 102 , 103 ] and Cdt1 in metazoa and fission yeast [ 120 , 121 , 122 , 123 , 124 , 125 , 126 , 164 ], see above) during the firing phase or to inactivate licensing inhibitors (e.g., metazoan geminin [ 117 ], see above) during the licensing phase. Additionally, cell cycle phase-specific degradation can be used to generate temporal order at cell cycle transitions.

A mechanism of early inactivation by degradation may be active at the G1/S transition in budding yeast. The licensing factor Cdc6, which is strongly regulated by degradation, including most importantly efficient inactivation by SCF–Cdc4 in S phase [ 100 , 101 , 102 ], is even a target for degradation in G1. Notably, with the exception of a possible involvement of the ubiquitin ligases Tom1 and Dia2, we currently have little knowledge of the mechanism and function of Cdc6 degradation in G1 (so called “mode 1” [ 101 , 103 ]). It is conceivable that enhanced turnover of Cdc6 prior to the G1/S transition may help Cdc6 inactivation.

Interestingly, a similar mechanism (early degradation) is active on the firing factor Sld2 before the metaphase-to-anaphase transition in budding yeast [ 145 ] and was also shown to be involved in meiotic DNA replication control [ 165 ]. Sld2 degradation at the metaphase-to-anaphase transition depends on CDK phosphorylation of Sld2 but additionally requires cell cycle kinases DDK, Cdc5, and Mck1, which phosphorylate Sld2 at a phospho-degron motif, which is read out by the ubiquitin ligases Dma1 and Dma2 [ 145 ]. Dependency on these four kinases generates a specific temporal window of Sld2 degradation in early M phase before the metaphase-to-anaphase transition. Enhanced Sld2 turnover at the metaphase-to-anaphase transition is apparently used to generate a temporal order of “inactivate firing first and activate licensing second”, as mutation of the Sld2 degron interferes with this order and shortens the gap between firing inactivation and licensing activation [ 145 ]. Notably, this mechanism is likely involved in preventing sporadic over-replication, as strains deficient in Sld2 degradation show an origin-dependent chromosomal rearrangement phenotype when combined with mutants that constitutively activate Sld3 [ 145 ]. Overall, both mechanisms leading to early degradation of Cdc6 and Sld2 can be envisioned to establish ordered inactivation of licensing/firing factors before activation of firing/licensing factors.

Notably, mammalian Cdc6 was also found to become degraded in G1-arrested cells by APC Cdh1 [ 166 ]. It seems, however, questionable whether this mechanism is involved in promoting early inactivation of Cdc6 prior to the G1/S transition, as Cdc6 becomes phosphorylated right at the G1/S transition by G1/S-CDK (cyclin E-Cdk2) [ 167 ]. This phosphorylation protects Cdc6 from APC-dependent degradation and induces a wave of licensing in late G1 [ 167 ]. While G1/S-CDK phosphorylation activates Cdc6, S-CDK (cyclin A-Cdk2) phosphorylation inhibits Cdc6 by promoting nuclear export [ 168 , 169 , 170 ]. Moreover, Cdc6 becomes degraded in S phase in a phosphorylation-independent manner [ 171 ]. With activating and inactivating phosphorylation marks and different modes of degradation, it is therefore at this point unclear how Cdc6 inactivation relates to the activation of firing factors.

Lastly, it is interesting to note that even substrates of the same ubiquitin ligase can display temporal order in the degradation. As such, it was shown for the APC that its substrates become degraded with temporal order ([ 172 , 173 ], see below). Similarly, the human CRL4–Cdt2 ubiquitin ligase, which specifically degrades “G1 factors” such as Cdt1 in early S phase in a manner that depends on the replication factor PCNA [ 120 , 122 , 123 , 164 ], targets different substrates to become degraded with different kinetics [ 174 ]. Interestingly, the licensing factor Cdt1 was shown to be degraded prior to p21, the critical S-CDK and G1/S-CDK inhibitor [ 174 ]. Importantly, according to our current understanding, degradation of both factors requires DNA-bound PCNA and therefore occurs after the first origins have already fired [ 122 , 123 , 164 ]. Nonetheless, these data suggest that licensing is inhibited due to Cdt1 degradation before S-CDK is fully active due to degradation of p21 [ 174 ]. This suggests that temporal ordering of degradation of different substrates by a single ubiquitin ligase may contribute to the overall temporal order of DNA replication control.

3.6. Temporal Order by a Two-Kinase System

An apparently universal feature of eukaryotic DNA replication is the dual control by CDK and DDK. The underlying reason for this two-kinase system, however, remains obscure. While essential roles for two independent kinases acting in the temporal program of replication or the response to replication perturbation seem likely [ 96 , 175 , 176 , 177 ], we would like to speculate that such a two-kinase system may also be suited to generate temporal order at cell cycle transitions.

Specifically, it needs to be noted that licensing inhibition has so far exclusively been attributed to CDK [ 5 , 99 , 100 , 101 , 102 , 104 ], while firing activation essentially requires both CDK and DDK [ 48 , 49 , 51 , 74 , 81 , 82 , 83 , 85 ]. Therefore, if CDK was activated before DDK at the G1/S transition, licensing would be inhibited before firing was activated, thus minimizing the risk of over-replication. Indeed, DDK activity is cell cycle regulated and kept low in G1 by APC-mediated degradation of Dbf4 [ 63 , 66 , 178 ]. However, when compared to CDK, overall changes in global DDK activity are not as pronounced [ 172 ]. Rather, the DDK activity profile appears to be sharpened locally by DDK inhibition at replication origins via the antagonizing Rif1–PP1 phosphatase complex [ 177 , 179 , 180 ] and via antagonizing Mcm2–7 modification with SUMO [ 181 ]. Currently, it is therefore unclear whether DDK is indeed activated after S-CDK.

However, genetic arguments from studies in budding yeast support that DDK might limit the timing of firing: (i) While previous studies have indicated that in the order of molecular events during firing the DDK-dependent step precedes the CDK-dependent steps [ 31 , 37 , 97 ], it was derived from conditional mutations in CDK or DDK that DDK action requires prior CDK activity [ 96 , 182 ]. (ii) DDK was found to be a limiting factor for origin firing [ 175 ] and it seems reasonable that the rather low amounts of DDK kinase will phosphorylate replication factors intrinsically more slowly than abundant CDK. In contrast, data from in vitro reconstitution using extracts or purified proteins suggests that DDK- and CDK-dependent steps are not necessarily separate [ 31 , 37 ]. Indeed, firing can be experimentally induced no matter whether replication proteins are first phosphorylated with CDK or DDK, respectively [ 37 ]. Nonetheless, these data would still be consistent with a temporal order of rising DDK and CDK activity at the G1/S transition in vivo.

Lastly, is the converse true in M phase? Is DDK inactivated prior to CDK at the metaphase-to-anaphase transition? Indeed, a recent study in budding yeast suggests that the APC mediates degradation of its key substrates (the anaphase inhibitor securin, S-phase cyclin Clb5, M-phase cyclin Clb2 and Dbf4) with a specific order and, indeed, Dbf4 was found to become degraded prior to Clb2, suggesting that at the metaphase-to-anaphase transition, DDK activity will drop prior to CDK activity [ 172 ]. This finding is therefore consistent with firing inactivation occurring prior to licensing reactivation in M phase and suggests that an overall purpose of the two-kinase control of replication might be temporal separation of licensing and firing.

4. Conclusions and Outlook

The principal control ensuring temporal separation of the two phases of replication initiation and thereby “once-per-cell-cycle replication” are well understood in several eukaryotic models. On top of this control exist several auxiliary mechanisms which appear to act specifically at cell cycle transitions between the two phases of replication initiation. Their sheer existence suggests that additional control is needed to equip DNA replication with the formidable accuracy that eukaryotes require to propagate a stable genome over generations. We propose that a specific purpose of these mechanisms is to generate temporal order in the inactivation and activation of licensing and firing factors at cell cycle transitions in order to provide a robust block against sporadic over-replication events which, otherwise, may manifest in genome instability [ 6 , 8 , 127 , 128 , 130 , 145 , 147 ]. Detailed studies of these mechanisms are needed to fill the gaps in our knowledge on how cells provide smooth transitions from licensing to firing phases and vice versa. In particular, we need to address scenarios of deregulated replication initiation leading to sporadic over-replication in order to reveal whether and to what degree single cells and, in particular, organisms can tolerate sporadic over-replication and whether sporadic over-replication could be a driving force in cancer.

Acknowledgments

We apologize to all our colleagues whose work could not be cited due to space limitations. We thank Lorenzo Galanti for critically reading the manuscript and for insightful discussion.

Research in the Pfander lab is funded by German Research Council and the Max Planck Society.

Conflicts of Interest

The authors declare that they do not have any competing interests.