PMCCPMCCPMCC

Search tips
Search criteria 

Advanced

 
Logo of genesMDPIHomeThis articleThis journalInstructions for authorsSubscribegenes
 
Genes (Basel). 2017 February; 8(2): 54.
Published online 2017 January 27. doi:  10.3390/genes8020054
PMCID: PMC5333043

Non-Canonical Replication Initiation: You’re Fired!

Eishi Noguchi, Academic Editor

Abstract

The division of prokaryotic and eukaryotic cells produces two cells that inherit a perfect copy of the genetic material originally derived from the mother cell. The initiation of canonical DNA replication must be coordinated to the cell cycle to ensure the accuracy of genome duplication. Controlled replication initiation depends on a complex interplay of cis-acting DNA sequences, the so-called origins of replication (ori), with trans-acting factors involved in the onset of DNA synthesis. The interplay of cis-acting elements and trans-acting factors ensures that cells initiate replication at sequence-specific sites only once, and in a timely order, to avoid chromosomal endoreplication. However, chromosome breakage and excessive RNA:DNA hybrid formation can cause break-induced (BIR) or transcription-initiated replication (TIR), respectively. These non-canonical replication events are expected to affect eukaryotic genome function and maintenance, and could be important for genome evolution and disease development. In this review, we describe the difference between canonical and non-canonical DNA replication, and focus on mechanistic differences and common features between BIR and TIR. Finally, we discuss open issues on the factors and molecular mechanisms involved in TIR.

Keywords: replication control, RNA:DNA hybrid, transcription-initiated replication

1. Origin-Dependent Replication

1.1. Chromosomal DNA Replication Initiation in Escherichia coli and Saccharomyces cerevisiae

Replication initiation at a single origin (ori) in the bacteria Escherichia coli has been the first, and until present, best-described mechanism of a classical replication initiation (see Figure 1; for reviews, see References [1,2,3,4,5]). Within the circular E. coli chromosome [6], a single origin called oriC provides a platform for protein recognition, local double-stranded DNA (dsDNA) opening, and access of the replication machinery [1]. OriC contains multiple repeats of the DnaA-box consensus sequence, and an AT-rich DNA-unwinding element (DUE) adjacent to the DnaA box [7] for the ATP-driven binding of the initiator protein DnaA [1]. OriC activation is coupled with bacterial growth rate [8], to efficiently initiate replication at the appropriate time and to avoid replication initiation at particular origins more than once [9,10,11,12,13]. DnaA binds to oriC and facilitates binding of the helicase loader-helicase DnaC–DnaB complex to form the pre-priming complex [4,14]. The DnaB helicase then stably interacts with the DnaG primase until RNA primer synthesis is accomplished [15]. Probably, RNA primer synthesis induces conformational changes that release DnaB from DnaG, because primer synthesis is coordinated with or followed by translocation of DnaB to the junction of the replication fork (reviewed in [16]). Subsequently, primer elongation by the DNA polymerase III (DNA Pol III) holoenzyme marks the switch from replication initiation to elongation [17,18]. In contrast to the single origin found in E. coli, the budding yeast Saccharomyces cerevisiae contains about 400 replication origins. The number of origins per genome is related to the genome size, explaining why eukaryotic genomes require more replication origins for their timely genome duplication [19]. Yeast continues to be one of the most advantageous model systems to study the basis of eukaryotic replication, but in contrast to prokaryotic cells, yeast chromosomes are packaged into nucleosomes. Dependent on their activation timing, replication origins can be separated into early and late replicating origins ([20,21,22], reviewed in [23]). In general, origin-dependent replication initiation requires the following conditions to be fulfilled: recognition of origins, pre-replicative complex (pre-RC) assembly during G1 phase (origin-licensing), and activation of the pre-RC at G1/S-phase (origin-firing; see Figure 1 and Table 1). S. cerevisiae origins are defined by a specific consensus sequence, known as autonomously replicating sequence (ARS) [24,25,26]. The AT-rich ARS consensus sequence (ACS) itself is not sufficient for replication initiation [27] but is required for the loading of the pre-RC during G1 phase ([28,29]). The pre-RC is composed of the origin recognition complex proteins Orc1–6 (ORC), Cdc6, Cdt1, and an inactive form of the replicative helicase Mcm2–7 complex ([30,31,32], reviewed in [33]). At G1/S-phase, the Dbf4-dependent kinase (DDK) and S-phase-dependent cyclin-dependent kinases (S-CDKs) phosphorylate Mcm4, Sld2, and Sld3 ([34,35]), prior to the stepwise recruitment of replication factors Cdc45/Sld3/Sld7 and Sld2/Dpb11/Mcm10/GINS/DNA Pol-ε ([36,37,38,39], see [40] for a review). Building up of the active Cdc45/Mcm2–7/GINS (CMG) helicase complex completes the replisome formation [41] and, consequently, DNA synthesis by the DNA Pol-α-primase complex is initiated [42]. Replication initiation is completed by the loading of the proliferating cell nuclear antigen (PCNA) onto the DNA Pol-α synthesized primer to switch to processive DNA synthesis by DNA Pol-ε and Pol-δ (see [43]).

Figure 1
Schematic outline of origin-dependent initiation of chromosomal and mitochondrial DNA replication. cis-acting origin DNA sequences (dotted lines), RNA (green), newly synthesized DNA (red), and helicases (green circle) are indicated. Note that chromosomal ...
Table 1
Factors required for origin-dependent DNA replication initiation in Escherichia coli and Saccharomyces cerevisiae.

Yeast has developed sophisticated mechanisms to avoid endoreplication events caused by replication re-initiation of already replicated origins. B-type CDKs prevent re-initiation through multiple overlapping mechanisms, including phosphorylation of ORC factors [44], nuclear exclusion of the Mcm2–7 complex and Cdc6 [45,46], transcriptional downregulation, polyubiquitination, and degradation of phosphorylated Cdc6 ([47,48,49]). Under certain conditions, traces of non-phosphorylatable Cdc6 [50] or mutations in components of the pre-replicative complex (origin recognition complex, Cdc6, and MCM proteins are sufficient to re-initiate DNA replication in G2/M cells. In the latter case, a Mec1 and Mre11-Rad50-Xrs2 (MRX) complex-dependent DNA damage signaling pathway is activated to restrain the extent of re-replication and to promote survival when origin-localized replication control pathways are abrogated [51]. Genome-wide analysis suggests that replication re-initiation in G2/M phase primarily occurs at a subset of both active and latent origins, but is independent of chromosomal determinants that specify the use and timing of these origins in S phase [52]. Moreover, the frequency and locations of re-replication events differ from the S to the G2/M phase, illustrating the dynamic nature of DNA replication controls [52]. Additional mechanisms may exist to prevent chromosomal re-replication in metazoans [53]. Interestingly, a recent study identified 42 uncharacterized human genes that are required to prevent either DNA re-replication or unscheduled endoreplication [54].

1.2. Mitochondrial DNA Replication Initiation

The variation in mitochondrial DNA (mtDNA) copy number reflects the fact that its replication cycle is not coupled with S phase-restricted, chromosomal DNA replication. Replication of mtDNA is connected with mtDNA transcription through the formation of a RNA:DNA hybrid that has been first detected by electron microscopy as a short three-stranded DNA region [55]. During transcription, the nascent transcript behind an elongating RNA polymerase (RNAP) can invade the double stranded DNA duplex and hybridize with the complementary DNA template strand. The formation of an RNA:DNA hybrid, opposite to an unpaired non-template DNA strand, results in a so-called R-loop structure (for a review see [56]). RNA:DNA hybrids are also the onset of Okazaki fragments, which serve as primers during DNA lagging-strand replication (for a review see [57]; see Figure 1 and Table 1). In the case of mtDNA replication, an R-loop is required for replication priming [58] at the mtDNA heavy-strand replication origin (OriH) and light-strand replication origin (OriL) [59]. OriH and OriL consist of a promoter and downstream conserved sequences with a high GC content, and are conserved from S. cerevisiae to humans [60]. Budding yeast contains about eight OriH-like regions (ori1–8; [60]) of which ori1–3 and ori5 represent bona fide origins of replication (see [61,62]). The OriH region of many organisms includes three conserved sequence blocks called CSB1, CSB2, and CSB3 [58], and transition from RNA to DNA synthesis is thought to happen at CSB2 [63]. Yeast mitochondrial RNA polymerase Rpo41, the helicase Irc3, and the single-stranded DNA (ssDNA)-binding protein Rim1 are the main factors involved in DNA strand separation during mtDNA replication [64,65,66]. After processing by RNase H1, the RNA molecule is used as a primer for DNA synthesis by the MIP1 encoded mitochondrial DNA polymerase γ (DNA Pol-γ) in budding yeasts [59]. Interestingly, in the absence of RNase H1, primer retention at OriL provides an obstacle for DNA Pol-γ [67], leading to mtDNA depletion and embryonic lethality in mice [68].

Apart from DNA Pol-γ, in metazoans the replicative mtDNA helicase Twinkle and the mitochondrial single-stranded DNA-binding protein (mtSSB) play key roles mtDNA replication fork progression (reviewed in [69,70]). The mechanism of mtDNA replication is not fully understood, and various possible mechanisms have been proposed ([71], reviewed in [72]). Currently, there are three main models of mtDNA replication. One is the initial “strand-displacement model”, proposing that leading strand DNA synthesis begins at a specific site and advances approximately two-thirds of the way around the molecule before DNA synthesis is initiated on the lagging strand [73]. A second “strand-coupled model” refers to a strand-asynchronous, unidirectional replication mode [74]. A third “RITOLS model” (RNA incorporation throughout the lagging strand) proposes that replication initiates in the major noncoding region at OriH, while OriL is a major initiation site of lagging-strand DNA synthesis but the lagging strand is laid down initially as RNA [75]. The idea of transcription-dependent mtDNA replication initiation has been unanimously accepted. However, by taking advantage of mutants devoid of the mitochondrial RNA polymerase Rpo41, Fangman et. al. suggested that replication priming by transcription is not the only mechanism for mtDNA replication initiation in yeast [76,77,78]. Alternatively, the mitochondrial ori5 has been shown to initiate mtDNA amplification by a rolling circle mechanism [79]. These kinds of replication events are linked to increased mtDNA damage and breaks by oxidative stress, and can be modulated by nuclease and recombinase activities carried out by Din7 and Mhr1, respectively [80].

Collectively, these findings demonstrate that mtDNA replication initiation is capable of adapting to stress situations, and that the stress-dependent, mitochondrial import of nuclear-encoded proteins such as Din7 and Mhr1 could provide another layer of mtDNA replication control. Interestingly, all other proteins involved in replication initiation are nuclear-encoded, and some genes, such as RNH1, encode both nuclear and mitochondrial protein isoforms [81]. It will be exciting to see if new players in mtDNA replication initiation may appear in response to different endogenous or exogenous stimuli. To date, little is known about how nuclear and mitochondrial replication checkpoints are interconnected, and how they control mtDNA replication initiation. Interestingly, a recent study showed that the DNA damage response protein kinase Rad53 (hChk2) is essential for an mtDNA inheritance checkpoint [82]. In mtDNA-depleted rho° cells, the DNA helicase Pif1 (petite integration frequency 1) undergoes Rad53-dependent phosphorylation. Pif1 is a highly conservative helicase localized to both nucleus and mitochondria in yeast and human cells [83] and promotes DNA replication through interaction with G-quadruplex DNA sequences ([84], reviewed in [85]). Thus, loss of mtDNA activates a nuclear checkpoint kinase that inhibits G1- to S-phase progression [82]. Pif1 is only one example of nuclear DNA helicases to protect mtDNA but, notably [86], it also has an essential role in recombination-dependent replication (as discussed subsequently). Future research may lead to the identification of other factors involved in the crosstalk between nuclear and mitochondrial genome duplication, and even improve our understanding of how the control of mitochondrial replication initiation is related to genome stability, aging, and mitochondrial diseases.

2. Origin-Independent Replication

2.1. Break-Induced Replication

A classic example of the initiation of origin-independent DNA replication events is recombination-dependent DNA replication, often called break-induced replication (BIR; see Figure 2 and Table 2, and [87] for a review). Kogoma and colleagues originally designated BIR in bacteria as DNA damage-inducible DNA replication, termed inducible stable DNA replication ((iSDR) [88,89], and reviewed in [90]). Double-strand end repair is initiated by break recognition and loading of the RecBCD helicase/nuclease complex. DNA unwinding by RecBCD leads to subsequent binding of RecA to ssDNA. Then, the strand exchange reaction between two recombining DNA double helices was proposed to as the mechanism by which DNA replication is primed [91,92]. DnaA is essential for helicase loading at oriC, whereas PriA, PriB, PriC, and DnaT appear to load DnaB into the forming replisome to promote replication fork assembly at a recombinational D-loop structure ([93], see [94] for a review). Finally, the branch migration and Holliday-junction resolving activities of the RuvABC complex are involved in the resolution of converging replication intermediates generated during iSDR [95].

Figure 2
Schematic representation of possible mechanism involved in origin-independent replication initiation by inducible stable DNA replication/break-induced replication (iSDR/BIR) or constitutive stable DNA replication/transcription-initiated replication (cSDR/TIR). ...
Table 2
Factors required for origin-independent DNA replication by iSDR/BIR or cSDR/TIR.

BIR was later found to occur in yeast upon transformation of yeast with linearized DNA fragments [96,97]. BIR turned out to promote DNA replication restart at broken replication forks and telomeres ([98,99], and reviewed in [87,100,101]) being an error-prone recombination-dependent DNA repair process that occurs in G2/M when only one end of a double-strand break (DSB) is available for recombination [102]. BIR can be Rad51-dependent or independent [102,103]. Rad51 is homologous to the bacterial ssDNA-binding protein RecA, and mainly involved in the search for homology and strand-pairing stages of homologous recombination [104]. Rad51-independent BIR at a one-ended break can occur when long-range strand invasion is not required. It primarily operates during intramolecular recombination; however, intermolecular events mostly rely on Rad51-dependent strand invasion [98,105]. More than 95% of BIR events in S. cerevisiae are reported to be Rad51-dependent and do not require either Rad50 or Rad59 [98,106], thus we discuss the Rad51-dependent pathway in more detail. During Rad51-dependent BIR, a DSB end is resected to produce a 3′-ended single-stranded DNA tail, subsequently coated by Rad51 nucleoprotein filaments [102]. This Rad51 filament then invades a homologous sequence and a D-loop is created, followed by an extension of the invading strand by new DNA synthesis using the paired homologous sequence as a template [107]. BIR is known to be a multistep process in which strand invasion occurs rapidly; by contrast, new DNA synthesis does not initiate until 3–4 h after strand invasion [99,102,108]. Once initiated, DNA synthesis may be very processive and continue to the end of the donor chromosome (reviewed in [109]).

Yeast proteins taking part in BIR also play a role in recombination. Recombination proteins Rad51, Rad52, Rad54, Rad55, and Rad57 initiate BIR by promoting strand invasion and D-loop formation [88,98]. BIR requires leading- and lagging-strand DNA synthesis and all essential DNA replication factors, including Pol-α-primase,Cdc7,Cdt1, Mcm10, Ctf4 and CMG helicase complex (except Cdc6 and ORC proteins), specific for pre-RC assembly and specifically needed for origin-dependent DNA replication [99,110]. It still remains to be determined how MCMs are recruited to the D-loop, but it is important to note that BIR occurs at the G2/M phase and normally depends on the Pif1 helicase. BIR may initiate in the absence of Pif1, but Pif1 appears to be required for long-range synthesis during BIR that proceeds by asynchronous synthesis of leading and lagging strands and leads to conservative inheritance of the new genetic material [111,112]. Analysis of BIR-dependent replication intermediates by 2D-agarose gels [113] revealed bubble arc-like migrating structures suggesting the accumulation of ssDNA at unrepaired DNA lesions within the template strand [112,114]. Investigation of BIR in yeast diploid cells led to observation of frequent switches of BIR between two homologous DNA templates, leading to the proposal that BIR is initiated via an unstable replication fork [115]. It was proposed that BIR could occur by several rounds of strand invasion, even at dispersed repeated sequences [115], leading to chromosome rearrangements [116]. However, the specific mechanisms of multiple strand invasions, D-loop displacement, and transition to a stable replication fork remain unknown.

Pol32, a nonessential subunit of Pol-δ, is another key player in BIR [111]. Pol32’s role in BIR is not unequivocally clear, but it has been reported to be essential for Rad51-dependent BIR [99] and required for replication fork processivity [111]. Interestingly, it has been recently shown that theMus81 endonuclease is required to limit BIR-associated template switching during Pol32-dependent DNA synthesis [117]. The involvement of structure-specific nucleases in BIR, such as Mus81-Mms4, Slx1-Slx4, and Yen1, suggests that these nucleases are needed for the processing or resolution of various types of BIR-dependent replication intermediates [118].

The establishment of a replication fork appears to be the slowest step in BIR. In bacteria, the normal initiation role of the DnaA and DnaC proteins in loading DnaB helicase at origins is replaced by the PriA complex (reviewed in [119,120]). PriA is implicated in loading DnaB onto replication fork structures other than replisomes, thus making PriA indispensable for the completion of any replication fork repair [121]. There is no obvious PriA homologue in eukaryotes, but it has been speculated that such a protein must exist. In yeast, the DnaB helicase function is provided by the Mcm2–7complex, which is conserved in all eukaryotes. The Cdc7–Dbf4 protein kinase promotes assembly of a stable Cdc45–MCM complex exclusively on chromatin in S phase [37], and, interestingly, BIR also requires the cell cycle-dependent kinase Cdc7 to initiate BIR [110]. As Rad51-dependent BIR occurs efficiently in G2-arrested yeast cells [102], either a subset of replication-competent MCM helicases remain bound to already replicated DNA, or DNA damage signaling leads to MCM-complex loading and Cdc7-dependent BIR activation in G2 phase. Recent studies show that SUMOylation and polyubiquitylation of MCM proteins have a role in replication initiation and termination, respectively [122,123,124]. It still remains to be determined if these post-translational MCM modifications affect BIR and if other helicases can drive BIR in the absence of MCM proteins. Pif1 may do so, as it already has a known role in BIR [111]. Pif1 is phosphorylated in response to DNA breaks by the Mec1/Rad53 DNA damage pathway in order to block the activity of telomerase at DNA breaks but not at chromosome ends [125], and its phosphorylation is required for BIR-mediated telomere replication in yeast [126]. Although this is pure speculation, it is conceivable that Pif1 might also be prone to Cdc7-dependent phosphorylation in order to fulfill its function in recombination-coupled DNA synthesis.

2.2. Transcription-Initiated Replication

R-loops have been shown to have roles in T4 bacteriophage, E. coli ColE1 plasmid, and mtDNA replication as well as B-cell immunoglobulin class switch recombination. R-loops are abundant structures, however, unscheduled R-loop formation challenges genome dynamics and function [127,128], and is related to neurological diseases and cancer (reviewed in [129,130,131,132,133]).

The role of R-loops in replication initiation was first demonstrated in E. coli ColE1 plasmid [134,135,136] and bacteriophage T4 replication (reviewed in [137]). Another legacy of Tokio Kogoma and colleagues was the discovery of oriC-independent DNA replication events ([138,139,140], reviewed in [90]). This type of replication was named constitutive stable DNA replication (cSDR) and, surprisingly, E. coli cells can stay alive exclusively on these origin-independent initiation events. One mutation that conferred this phenotype was found to inactivate the rnhA gene encoding RNase H1, an RNase specific to RNA in the RNA:DNA hybrid form [141,142]. cSDR was thought to originate from chromosomal sites named oriK, and only recently have specific candidate locations for oriK been mapped [143]. Moreover, it has been shown that origin-independent DNA synthesis arises in E. coli cells lacking the RecG helicase and results in chromosome duplication [144]. In contrast to RNase H1, RecG deals with replication fork fusion intermediates [145,146]; hence, origin-independent synthesis is initiated in different ways, but in both cases a fraction of forks will proceed in an orientation opposite to normal [144]. Drolet et al. [147] provided first evidence that R-loops can accumulate incells lacking topA, which encodes a type 1A topoisomerase that relieves negative supercoiling behind the RNAP, by showing that overexpression of rnhA partially compensates for the lack of topA. Notably, E. coli possesses two type 1A enzymes, Top1 (topA-encoded) and Top3 (topB-encoded), but only cells lacking Top1 are prone to cSDR [148]. Apart from transcription, cSDR requires RecA, and the primosome-complex including PriA, PriB, DnaT, and DNA Pol I [90,149,150]. RecA may also participate in cSDR by binding to ssDNA to stabilize an R-loop, or facilitate an inverse strand exchange reaction performed by RecA ([151,152], see Figure 2). In cSDR, DNA Pol I is thought to extend the RNA of the R-loop and to provide a substrate for PriA binding, as well as DnaB and DNA Pol III loading [90]. Interestingly, cSDR uses the same replicative helicase (DnaB) and replisome components (DNA Pol III) to initiate replication from oriC, but uses the PriA-dependent primosome for replicative helicase loading [90], as is the case for replication restart of disassembled replisomes [94]. Improperly regulated DNA replication may lead to various consequences related to genome instability. Interestingly, evidence that R-loop-dependent replication leads to DNA breakage and genome instability in non-growing E. coli cells has been presented [153], and mutations reducing replication from R-loops suppress the defects of growth, chromosome segregation, and DNA supercoiling in cells lacking Top1 and RNase H1 activity [154].

Transcription-linked replication initiation in eukaryotic cells was thought to be an exclusive feature of mtDNA replication. Yet, some highly transcribed DNA regions, such as RNAPI-transcribed ribosomal DNA (rDNA) or RNAP III-transcribed genes, were shown to be hot spots for R-loop formation in yeast mutants lacking RNases H [155,156]. In addition, mutants lacking an RNA/DNA helicase Sen1 [157,158] or the yeast Pab1-binding protein Pbp1 (hAtaxin-2) had been found to increase R-loop formation [159]. The absence of RNase H and Top1 activities causes synthetic lethality in yeast, suggesting that persistent R-loop formation could constrain cell viability [160,161]. Accordingly, persistent R-loop formation could be induced by treatment of RNase H mutants with the Top1 inhibitor camptothecin (CPT) leading to the detection of unscheduled transcription-initiated replication (TIR) events in yeast ([161], see Figure 2). TIR initiation intermediates were observed within the rDNA region, but were not linked to a defined replication origin; moreover, they were observed in the late S/G2 phase of the cell cycle, when replication termination and completion was expected to take place [161]. TIR was RNAPI transcription-dependent and led to replication fork pausing sites at sites of protein–DNA interaction. Taken together, these results suggest that R-loops could mediate origin-independent replication initiation events that constitute a non-canonical replisome, lacking the factors required to bypass replication constrains.

The factors and mechanisms participating in transcription-initiated replication events still remain to be elucidated. Various nonexclusive mechanisms could cooperate to trigger TIR events (summarized in Figure 2). These include strand invasion-dependent replication events that might be stimulated by the presence of single-stranded DNA within R-loops. In the absence of RNase H and Top1 activities, the rDNA locus turns into a hotspot for DSBs [161], thus it is conceivable that these DSBs drive recombination-dependent replication such as BIR. Other possibilities include that R-loops cause replication fork collapse and TIR is the result of replication restart of a replisome–RNAP complex [162,163]. An interesting possibility would be de novo replisome assembly at an R-loop. The RNA present within the R-loop could prime leading-strand synthesis and provoke assembly of replication-competent replicases at S/G2 phase [164]. Apparently, ssDNA opposite an RNA:DNA hybrid could activate Mec1-mediated checkpoint activation and binding of the replication protein A (RPA) complex, which has been shown to be involved in replication initiation as well as DNA repair by interacting with both the DNA Pol-α-primase complex and with DNA Pol-δ [164,165]. An R-loop may promote DNA replication restart by Pol-α-driven DNA synthesis, since the essential DNA Pol-α-primase subunit Pol12 remains active and phosphorylated in S/G2 and is inactivated while cells exit mitosis [44,161,166]. Moreover, a recent work by Symington and coworkers suggests that BIR occurs by a conservative mode of DNA synthesis [107]. Thus, it will be interesting to determine whether the same is true for TIR, or if TIR pursues a semiconservative replication mode. It is striking that in E. coli, many factors involved in iSDR are also needed for cSDR. These findings suggest that in yeast, many factors involved in BIR might be required for TIR. These factors include proteins involved in homologous recombination, DNA end-processing, helicases, primases, DNA polymerases, and, finally, structure-specific endonucleases (as listed in Table 2). Nevertheless, genetic interactions in yeast cells between RNase H deficiencies and proteins involved in BIR still remain to be determined.

Yet-to-be determined questions include whether TIR is limited to rDNA, and whether TIR can be observed in other RNA/DNA helicases mutants, including Sen1 [156,157,158] or the yeast ataxin-2 protein Pbp1 [159]. Recently, it has been shown that replication initiates, albeit very infrequently, within the telomeric repeats [167]. A long noncoding telomeric repeat-containing RNA (TERRA) has been implicated in telomere maintenance during replicative senescence and cancer [168,169]. TERRA accumulates specifically at short telomeres and may promote replication-fork restarting by recruiting homology-directed repair (HDR) mediators or even by directly priming replication in an origin-independent manner [167], similar to what was reported by Stuckey et al. [161]. This proposal might be supported by the fact that the cell cycle regulation of TERRA becomes perturbed at telomeres that are maintained by HDR, and that TERRA remains telomere-associated at G2/M in cells that use the alternative lengthening of telomeres (ALT) mechanism [170]. Interestingly, loss of ATP-dependent helicase ATRX that is frequently mutated in ALT-positive cancers, leads to persistent association of RPA with telomeres after DNA replication [170]. ATRX is involved in establishing transcriptionally silenced heterochromatin, and one hypothesis is that ATRX helicase and ATPase activity resolves G4 DNA secondary structures formed opposite of a TERRA-containing R-loop ([169,171], reviewed in [167]).

3. Conclusions

Since the detection of recombination-dependent replication of the E. coli chromosome by Lark and Kogoma about 50 years ago [172], we have learned a lot about mechanisms that can lead to non-canonical replication initiation in prokaryotic and eukaryotic cells. It is generally accepted that recombination serves to rescue broken chromosomes and stalled replication forks, however, we are far away from the complete picture on how cells manage to bypass the need for origin-dependent replication initiation. The mechanistic models and enzymatic steps leading to iSDR and cSDR in E. coli can be considered as a blueprint for BIR and TIR events in eukaryotic systems. Interestingly, all known features of BIR and TIR can participate in mtDNA replication events. Nevertheless, an important difference is noted by the fact that nuclear BIR and TIR events happen in a chromatin context with eukaryotic replication, starting with nucleosome packaging.

Many aspects of non-canonical DNA replication in eukaryotes still remain unknown and deserve to be addressed in the future; in particular, the factors driving replication fork progression and the mode of TIR-dependent DNA synthesis need to be characterized. Special attention should be given to the identification of key replication factors involved in TIR, such as DNA polymerases and helicases, but also to otherwise auxiliary replication proteins such as Pol32. R-loops are essential for the onset of TIR, and this might not be the only difference between TIR and BIR events. As outlined in Figure 2, the question remains if TIR is driven by strand invasion of the R-loop. TIR has been characterized only in repetitive ribosomal DNA sequences, raising the question of whether it is sister-chromatid-dependent, or if it uses non-sister chromatids as a template for DNA synthesis. In either case, strand invasion could be Rad51-dependent or independent. However, the role of Rad51 in TIR still needs to be determined. Genetic screens might help to shed light on factors required for TIR initiation and provide more insight to the differences between TIR and BIR.

The other model proposed in Figure 2 includes de novo assembly of a replication fork at an R-loop. In this case, which replication factors would be assembled at an R-loop, and would this kind of non-canonical replication restart be S phase-dependent? Would conservative or semiconservative replication account for the newly synthesized DNA? Could an R-loop even contribute to the activation of less defined replication origins in higher eukaryotes? Unrevealed functions of R-loops in higher eukaryotes may include a role the epigenetic regulation of origin-dependent replication initiation [173,174]. Interestingly, a nuclease-resistant G-quadruplex hybrid structure involving both RNA and DNA is present at the mtDNA replication initiation site [65]. G-rich RNA mediates Epstein-Barr virus nuclear antigen 1 EBNA1 and ORC interaction [175], thus it is conceivable that that transcription-related RNA structures might replace the need for specific origin-recognition sequences. By using a high-resolution PCR strategy to localize replication origins directly on total unfractionated human DNA, over-replicated regions were found to overlap with transcription initiation sites of CpG island promoters [176] and, recently, active transcription was proposed to be a driving force for the human parasite Leishmania major spatial and the temporal program of DNA replication [177]. Last but not least, TIR could be considered as an ancient mechanism to promote gene amplification events linked to nuclear differentiation and evolution. In order to resolve these questions, future studies should include higher eukaryotic model systems to see if TIR has a role in genome stability connected to various human diseases, including cancer.

Acknowledgments

This work was supported in part by the Spanish Ministry of Science and Innovation (MINECO; BFU2015-69183-P) and the European Union (FEDER; R.E.W.) and an ERASMUS+ fellowship of the European Union (B.R.). We would like to thank Hélène Gaillard and Daniel Fitzgerald for critical reading of the manuscript, and Zoe Cooper for style correction.

Author Contributions

Author Contributions

R.E.W. organized and wrote the paper. B.R. contributed to writing the paper and designing the figures.

Conflicts of Interest

Conflicts of Interest

The authors declare no conflict of interest. The founding sponsors had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, and in the decision to publish the results.

References

1. Fuller R.S., Funnell B.E., Kornberg A. The DnaA protein complex with the E. coli chromosomal replication origin (oriC) and other DNA sites. Cell. 1984;38:889–900. doi: 10.1016/0092-8674(84)90284-8. [PubMed] [Cross Ref]
2. Hwang D.S., Kornberg A. Opening of the replication origin of Escherichia coli by DnaA protein with protein HU or IHF. J. Biol. Chem. 1992;267:23083–23086. [PubMed]
3. Boye E., Lobner-Olesen A., Skarstad K. Limiting DNA replication to once and only once. EMBO Rep. 2000;1:479–483. doi: 10.1093/embo-reports/kvd116. [PubMed] [Cross Ref]
4. Mott M.L., Berger J.M. DNA replication initiation: Mechanisms and regulation in bacteria. Nat. Rev. Microbiol. 2007;5:343–354. doi: 10.1038/nrmicro1640. [PubMed] [Cross Ref]
5. Wolanski M., Donczew R., Zawilak-Pawlik A., Zakrzewska-Czerwinska J. OriC-encoded instructions for the initiation of bacterial chromosome replication. Front. Microbiol. 2014;5:735. [PMC free article] [PubMed]
6. Kohara Y., Akiyama K., Isono K. The physical map of the whole E. coli chromosome: Application of a new strategy for rapid analysis and sorting of a large genomic library. Cell. 1987;50:495–508. doi: 10.1016/0092-8674(87)90503-4. [PubMed] [Cross Ref]
7. Speck C., Messer W. Mechanism of origin unwinding: Sequential binding of DnaA to double- and single-stranded DNA. EMBO J. 2001;20:1469–1476. doi: 10.1093/emboj/20.6.1469. [PubMed] [Cross Ref]
8. Wold S., Skarstad K., Steen H.B., Stokke T., Boye E. The initiation mass for DNA replication in Escherichia coli K-12 is dependent on growth rate. EMBO J. 1994;13:2097–2102. [PubMed]
9. Yamaki H., Ohtsubo E., Nagai K., Maeda Y. The oriC unwinding by dam methylation in Escherichia coli. Nucleic Acids Res. 1988;16:5067–5073. doi: 10.1093/nar/16.11.5067. [PMC free article] [PubMed] [Cross Ref]
10. Campbell J.L., Kleckner N. E. coli oriC and the dnaA gene promoter are sequestered from dam methyltransferase following the passage of the chromosomal replication fork. Cell. 1990;62:967–979. doi: 10.1016/0092-8674(90)90271-F. [PubMed] [Cross Ref]
11. Boye E., Stokke T., Kleckner N., Skarstad K. Coordinating DNA replication initiation with cell growth: Differential roles for DnaA and SeqA proteins. Proc. Natl. Acad. Sci. USA. 1996;93:12206–12211. doi: 10.1073/pnas.93.22.12206. [PubMed] [Cross Ref]
12. Torheim N.K., Boye E., Løbner-Olesen A., Stokke T., Skarstad K. The Escherichia coli SeqA protein destabilizes mutant DnaA204 protein. Mol. Microbiol. 2000;37:629–638. doi: 10.1046/j.1365-2958.2000.02031.x. [PubMed] [Cross Ref]
13. Fujimitsu K., Senriuchi T., Katayama T. Specific genomic sequences of E. coli promote replicational initiation by directly reactivating ADP-DnaA. Genes Dev. 2009;23:1221–1233. doi: 10.1101/gad.1775809. [PubMed] [Cross Ref]
14. Bramhill D., Kornberg A. Duplex opening by DnaA protein at novel sequences in initiation of replication at the origin of the E. coli chromosome. Cell. 1988;52:743–755. doi: 10.1016/0092-8674(88)90412-6. [PubMed] [Cross Ref]
15. Chang P., Marians K.J. Identification of a region of Escherichia coli DnaB required for functional interaction with DnaG at the replication fork. J. Biol. Chem. 2000;275:26187–26195. doi: 10.1074/jbc.M001800200. [PubMed] [Cross Ref]
16. Chodavarapu S., Kaguni J.M. The Enzymes. Volume 39. Academic Press; New York, NY, USA: 2016. Replication initiation in bacteria; pp. 1–30. Chapter 1. [PubMed]
17. O’Donnell M.E., Kornberg A. Complete replication of templates by Escherichia coli DNA polymerase III holoenzyme. J. Biol. Chem. 1985;260:12884–12889. [PubMed]
18. Lewis J.S., Jergic S., Dixon N.E. The Enzymes. Volume 39. Academic Press; New York, NY, USA: 2016. The E. coli DNA replication fork; pp. 31–88. Chapter 2. [PubMed]
19. Gilbert D.M. Replication origins in yeast versus metazoa: Separation of the haves and the have nots. Curr. Opin. Genet. Dev. 1998;8:194–199. doi: 10.1016/S0959-437X(98)80141-X. [PubMed] [Cross Ref]
20. Yamashita M., Hori Y., Shinomiya T., Obuse C., Tsurimoto T., Yoshikawa H., Shirahige K. The efficiency and timing of initiation of replication of multiple replicons of Saccharomyces cerevisiae chromosome VI. Genes Cells. 1997;2:655–665. doi: 10.1046/j.1365-2443.1997.1530351.x. [PubMed] [Cross Ref]
21. Das S.P., Borrman T., Liu V.W., Yang S.C., Bechhoefer J., Rhind N. Replication timing is regulated by the number of MCMs loaded at origins. Genome Res. 2015;25:1886–1892. doi: 10.1101/gr.195305.115. [PubMed] [Cross Ref]
22. Peace J.M., Ter-Zakarian A., Aparicio O.M. Rif1 regulates initiation timing of late replication origins throughout the S. cerevisiae genome. PLoS ONE. 2014;9:e98501 doi: 10.1371/journal.pone.0098501. [PMC free article] [PubMed] [Cross Ref]
23. Goren A., Cedar H. Replicating by the clock. Nat. Rev. Mol. Cell Biol. 2003;4:25–32. doi: 10.1038/nrm1008. [PubMed] [Cross Ref]
24. Stinchcomb D.T., Struhl K., Davis R.W. Isolation and characterisation of a yeast chromosomal replicator. Nature. 1979;282:39–43. doi: 10.1038/282039a0. [PubMed] [Cross Ref]
25. Bell S.P., Stillman B. ATP-dependent recognition of eukaryotic origins of DNA replication by a multiprotein complex. Nature. 1992;357:128–134. doi: 10.1038/357128a0. [PubMed] [Cross Ref]
26. Marahrens Y., Stillman B. A yeast chromosomal origin of DNA replication defined by multiple functional elements. Science. 1992;255:817–823. doi: 10.1126/science.1536007. [PubMed] [Cross Ref]
27. Nieduszynski C.A., Knox Y., Donaldson A.D. Genome-wide identification of replication origins in yeast by comparative genomics. Genes Dev. 2006;20:1874–1879. doi: 10.1101/gad.385306. [PubMed] [Cross Ref]
28. Rao H., Stillman B. The origin recognition complex interacts with a bipartite DNA binding site within yeast replicators. Proc. Natl. Acad. Sci. USA. 1995;92:2224–2228. doi: 10.1073/pnas.92.6.2224. [PubMed] [Cross Ref]
29. Speck C., Chen Z., Li H., Stillman B. ATPase-dependent cooperative binding of ORC and Cdc6 to origin DNA. Nat. Struct. Mol. Biol. 2005;12:965–971. doi: 10.1038/nsmb1002. [PMC free article] [PubMed] [Cross Ref]
30. Aparicio O.M., Weinstein D.M., Bell S.P. Components and dynamics of DNA replication complexes in S. cerevisiae: Redistribution of MCM proteins and Cdc45p during S phase. Cell. 1997;91:59–69. doi: 10.1016/S0092-8674(01)80009-X. [PubMed] [Cross Ref]
31. Kawasaki Y., Kim H.D., Kojima A., Seki T., Sugino A. Reconstitution of Saccharomyces cerevisiae prereplicative complex assembly in vitro. Genes Cells. 2006;11:745–756. doi: 10.1111/j.1365-2443.2006.00975.x. [PubMed] [Cross Ref]
32. Tanaka S., Diffley J.F. Interdependent nuclear accumulation of budding yeast Cdt1 and Mcm2–7 during G1 phase. Nat. Cell Biol. 2002;4:198–207. doi: 10.1038/ncb757. [PubMed] [Cross Ref]
33. Chesnokov I.N. Multiple functions of the origin recognition complex. Int. Rev. Cytol. 2007;256:69–109. [PubMed]
34. Tanaka S., Umemori T., Hirai K., Muramatsu S., Kamimura Y., Araki H. CDK-dependent phosphorylation of Sld2 and Sld3 initiates DNA replication in budding yeast. Nature. 2007;445:328–332. doi: 10.1038/nature05465. [PubMed] [Cross Ref]
35. Sheu Y.J., Stillman B. Cdc7-Dbf4 phosphorylates MCM proteins via a docking site-mediated mechanism to promote S phase progression. Mol. Cell. 2006;24:101–113. doi: 10.1016/j.molcel.2006.07.033. [PMC free article] [PubMed] [Cross Ref]
36. Zou L., Stillman B. Formation of a preinitiation complex by S-phase cyclin CDK-dependent loading of Cdc45p onto chromatin. Science. 1998;280:593–596. doi: 10.1126/science.280.5363.593. [PubMed] [Cross Ref]
37. Muramatsu S., Hirai K., Tak Y.S., Kamimura Y., Araki H. CDK-dependent complex formation between replication proteins Dpb11, Sld2, Polε, and GINS in budding yeast. Genes Dev. 2010;24:602–612. doi: 10.1101/gad.1883410. [PubMed] [Cross Ref]
38. Homesley L., Lei M., Kawasaki Y., Sawyer S., Christensen T., Tye B.K. Mcm10 and the Mcm2–7 complex interact to initiate DNA synthesis and to release replication factors from origins. Genes Dev. 2000;14:913–926. [PubMed]
39. Gambus A., Jones R.C., Sanchez-Diaz A., Kanemaki M., van Deursen F., Edmondson R.D., Labib K. GINS maintains association of Cdc45 with MCM in replisome progression complexes at eukaryotic DNA replication forks. Nat. Cell Biol. 2006;8:358–366. doi: 10.1038/ncb1382. [PubMed] [Cross Ref]
40. Tanaka S., Araki H. Helicase activation and establishment of replication forks at chromosomal origins of replication. Cold Spring Harb. Perspect. Biol. 2013;5:a010371. doi: 10.1101/cshperspect.a010371. [PMC free article] [PubMed] [Cross Ref]
41. Moyer S.E., Lewis P.W., Botchan M.R. Isolation of the Cdc45/Mcm2–7/GINS (CMG) complex, a candidate for the eukaryotic DNA replication fork helicase. Proc. Natl. Acad. Sci. USA. 2006;103:10236–10241. doi: 10.1073/pnas.0602400103. [PubMed] [Cross Ref]
42. Foiani M., Marini F., Gamba D., Lucchini G., Plevani P. The B subunit of the DNA polymerase alpha-primase complex in Saccharomyces cerevisiae executes an essential function at the initial stage of DNA replication. Mol. Cell. Biol. 1994;14:923–933. doi: 10.1128/MCB.14.2.923. [PMC free article] [PubMed] [Cross Ref]
43. Garg P., Burgers P.M. DNA polymerases that propagate the eukaryotic DNA replication fork. Crit. Rev. Biochem. Mol. Biol. 2005;40:115–128. doi: 10.1080/10409230590935433. [PubMed] [Cross Ref]
44. Weinreich M., Liang C., Chen H.H., Stillman B. Binding of cyclin-dependent kinases to ORC and Cdc6p regulates the chromosome replication cycle. Proc. Natl. Acad. Sci. USA. 2001;98:11211–11217. doi: 10.1073/pnas.201387198. [PubMed] [Cross Ref]
45. Labib K., Diffley J.F., Kearsey S.E. G1-phase and B-type cyclins exclude the DNA-replication factor Mcm4 from the nucleus. Nat. Cell Biol. 1999;1:415–422. [PubMed]
46. Nguyen V.Q., Co C., Li J.J. Cyclin-dependent kinases prevent DNA re-replication through multiple mechanisms. Nature. 2001;411:1068–1073. doi: 10.1038/35082600. [PubMed] [Cross Ref]
47. Moll T., Tebb G., Surana U., Robitsch H., Nasmyth K. The role of phosphorylation and the CDC28 protein kinase in cell cycle-regulated nuclear import of the S. cerevisiae transcription factor SWI5. Cell. 1991;66:743–758. doi: 10.1016/0092-8674(91)90118-I. [PubMed] [Cross Ref]
48. Drury L.S., Perkins G., Diffley J.F. The Cdc4/34/53 pathway targets Cdc6p for proteolysis in budding yeast. EMBO J. 1997;16:5966–5976. doi: 10.1093/emboj/16.19.5966. [PubMed] [Cross Ref]
49. Drury L.S., Perkins G., Diffley J.F. The cyclin-dependent kinase Cdc28p regulates distinct modes of Cdc6p proteolysis during the budding yeast cell cycle. Curr. Biol. 2000;10:231–240. doi: 10.1016/S0960-9822(00)00355-9. [PubMed] [Cross Ref]
50. Honey S., Futcher B. Roles of the CDK Phosphorylation Sites of Yeast Cdc6 in Chromatin Binding and Rereplication. Mol. Biol. Cell. 2007;18:1324–1336. doi: 10.1091/mbc.E06-06-0544. [PMC free article] [PubMed] [Cross Ref]
51. Archambault V., Ikui A.E., Drapkin B.J., Cross F.R. Disruption of mechanisms that prevent rereplication triggers a DNA damage response. Mol. Cell. Biol. 2005;25:6707–6721. doi: 10.1128/MCB.25.15.6707-6721.2005. [PMC free article] [PubMed] [Cross Ref]
52. Green B.M., Morreale R.J., Ozaydin B., Derisi J.L., Li J.J. Genome-wide mapping of DNA synthesis in Saccharomyces cerevisiae reveals that mechanisms preventing reinitiation of DNA replication are not redundant. Mol. Biol. Cell. 2006;17:2401–2414. doi: 10.1091/mbc.E05-11-1043. [PMC free article] [PubMed] [Cross Ref]
53. Blow J.J., Dutta A. Preventing re-replication of chromosomal DNA. Nat. Rev. Mol. Cell Biol. 2005;6:476–486. doi: 10.1038/nrm1663. [PMC free article] [PubMed] [Cross Ref]
54. Vassilev A., Lee C.Y., Vassilev B., Zhu W., Ormanoglu P., Martin S.E., DePamphilis M.L. Identification of genes that are essential to restrict genome duplication to once per cell division. Oncotarget. 2016;7:34956–34976. doi: 10.18632/oncotarget.9008. [PMC free article] [PubMed] [Cross Ref]
55. Kasamatsu H., Robberson D.L., Vinograd J. A novel closed-circular mitochondrial DNA with properties of a replicating intermediate. Proc. Natl. Acad. Sci. USA. 1971;68:2252–2257. doi: 10.1073/pnas.68.9.2252. [PubMed] [Cross Ref]
56. Aguilera A., Garcia-Muse T. R loops: From transcription byproducts to threats to genome stability. Mol. Cell. 2012;46:115–124. doi: 10.1016/j.molcel.2012.04.009. [PubMed] [Cross Ref]
57. Lujan S.A., Williams J.S., Kunkel T.A. DNA polymerases divide the labor of genome replication. Trends Cell Biol. 2016;26:640–654. doi: 10.1016/j.tcb.2016.04.012. [PubMed] [Cross Ref]
58. Xu B., Clayton D.A. RNA-DNA hybrid formation at the human mitochondrial heavy-strand origin ceases at replication start sites: An implication for RNA-DNA hybrids serving as primers. EMBO J. 1996;15:3135–3143. [PubMed]
59. Nicholls T.J., Minczuk M. In D-loop: 40 years of mitochondrial 7S DNA. Exp. Gerontol. 2014;56:175–181. doi: 10.1016/j.exger.2014.03.027. [PubMed] [Cross Ref]
60. Baldacci G., Cherif-Zahar B., Bernardi G. The initiation of DNA replication in the mitochondrial genome of yeast. EMBO J. 1984;3:2115–2120. [PubMed]
61. Shadel G.S. Yeast as a model for human mtDNA replication. Am. J. Hum. Genet. 1999;65:1230–1237. doi: 10.1086/302630. [PubMed] [Cross Ref]
62. Williamson D. The curious history of yeast mitochondrial DNA. Nat. Rev. Genet. 2002;3:475–481. [PubMed]
63. Wanrooij P.H., Uhler J.P., Shi Y., Westerlund F., Falkenberg M., Gustafsson C.M. A hybrid G-quadruplex structure formed between RNA and DNA explains the extraordinary stability of the mitochondrial R-loop. Nucleic Acids Res. 2012;40:10334–10344. doi: 10.1093/nar/gks802. [PMC free article] [PubMed] [Cross Ref]
64. Sanchez-Sandoval E., Diaz-Quezada C., Velazquez G., Arroyo-Navarro L.F., Almanza-Martinez N., Trasvina-Arenas C.H., Brieba L.G. Yeast mitochondrial RNA polymerase primes mitochondrial DNA polymerase at origins of replication and promoter sequences. Mitochondrion. 2015;24:22–31. doi: 10.1016/j.mito.2015.06.004. [PubMed] [Cross Ref]
65. Sedman T., Gaidutsik I., Villemson K., Hou Y., Sedman J. Double-stranded DNA-dependent ATPase Irc3p is directly involved in mitochondrial genome maintenance. Nucleic Acids Res. 2014;42:13214–13227. doi: 10.1093/nar/gku1148. [PMC free article] [PubMed] [Cross Ref]
66. Van Dyck E., Foury F., Stillman B., Brill S.J. A single-stranded DNA binding protein required for mitochondrial DNA replication in S. cerevisiae is homologous to E. coli SSB. EMBO J. 1992;11:3421–3430. [PubMed]
67. Holmes J.B., Akman G., Wood S.R., Sakhuja K., Cerritelli S.M., Moss C., Bowmaker M.R., Jacobs H.T., Crouch R.J., Holt I.J. Primer retention owing to the absence of RNase H1 is catastrophic for mitochondrial DNA replication. Proc. Natl. Acad. Sci. USA. 2015;112:9334–9339. doi: 10.1073/pnas.1503653112. [PubMed] [Cross Ref]
68. Cerritelli S.M., Frolova E.G., Feng C., Grinberg A., Love P.E., Crouch R.J. Failure to produce mitochondrial DNA results in embryonic lethality in Rnaseh1 null mice. Mol. Cell. 2003;11:807–815. doi: 10.1016/S1097-2765(03)00088-1. [PubMed] [Cross Ref]
69. Holt I.J. Mitochondrial DNA replication and repair: All a flap. Trends Biochem. Sci. 2009;34:358–365. doi: 10.1016/j.tibs.2009.03.007. [PubMed] [Cross Ref]
70. Ciesielski G.L., Oliveira M.T., Kaguni L.S. The Enzymes. Volume 39. Academic Press; New York, NY, USA: 2016. Animal mitochondrial DNA replication; pp. 255–292. Chapter 8. [PMC free article] [PubMed]
71. Gerhold J.M., Aun A., Sedman T., Joers P., Sedman J. Strand invasion structures in the inverted repeat of Candida albicans mitochondrial DNA reveal a role for homologous recombination in replication. Mol. Cell. 2010;39:851–861. doi: 10.1016/j.molcel.2010.09.002. [PubMed] [Cross Ref]
72. Gustafsson C.M., Falkenberg M., Larsson N.G. Maintenance and expression of mammalian mitochondrial DNA. Annu. Rev. Biochem. 2016;85:133–160. doi: 10.1146/annurev-biochem-060815-014402. [PubMed] [Cross Ref]
73. Clayton D.A. Replication of animal mitochondrial DNA. Cell. 1982;28:693–705. doi: 10.1016/0092-8674(82)90049-6. [PubMed] [Cross Ref]
74. Holt I.J., Lorimer H.E., Jacobs H.T. Coupled leading- and lagging-strand synthesis of mammalian mitochondrial DNA. Cell. 2000;100:515–524. doi: 10.1016/S0092-8674(00)80688-1. [PubMed] [Cross Ref]
75. Yasukawa T., Reyes A., Cluett T.J., Yang M.Y., Bowmaker M., Jacobs H.T., Holt I.J. Replication of vertebrate mitochondrial DNA entails transient ribonucleotide incorporation throughout the lagging strand. EMBO J. 2006;25:5358–5371. doi: 10.1038/sj.emboj.7601392. [PubMed] [Cross Ref]
76. Fangman W.L., Henly J.W., Churchill G., Brewer B.J. Stable maintenance of a 35-base-pair yeast mitochondrial genome. Mol. Cell. Biol. 1989;9:1917–1921. doi: 10.1128/MCB.9.5.1917. [PMC free article] [PubMed] [Cross Ref]
77. Fangman W.L., Henly J.W., Brewer B.J. RPO41-independent maintenance of [rho-] mitochondrial DNA in Saccharomyces cerevisiae. Mol. Cell. Biol. 1990;10:10–15. doi: 10.1128/MCB.10.1.10. [PMC free article] [PubMed] [Cross Ref]
78. Lorimer H.E., Brewer B.J., Fangman W.L. A test of the transcription model for biased inheritance of yeast mitochondrial DNA. Mol. Cell. Biol. 1995;15:4803–4809. doi: 10.1128/MCB.15.9.4803. [PMC free article] [PubMed] [Cross Ref]
79. Ling F., Hori A., Shibata T. DNA recombination-initiation plays a role in the extremely biased inheritance of yeast [rho-] mitochondrial DNA that contains the replication origin ori5. Mol. Cell. Biol. 2007;27:1133–1145. doi: 10.1128/MCB.00770-06. [PMC free article] [PubMed] [Cross Ref]
80. Ling F., Hori A., Yoshitani A., Niu R., Yoshida M., Shibata T. Din7 and Mhr1 expression levels regulate double-strand-break-induced replication and recombination of mtDNA at ori5 in yeast. Nucleic Acids Res. 2013;41:5799–5816. doi: 10.1093/nar/gkt273. [PMC free article] [PubMed] [Cross Ref]
81. Engel M.L., Hines J.C., Ray D.S. The Crithidia fasciculata RNH1 gene encodes both nuclear and mitochondrial isoforms of RNase H. Nucleic Acids Res. 2001;29:725–731. doi: 10.1093/nar/29.3.725. [PMC free article] [PubMed] [Cross Ref]
82. Crider D.G., Garcia-Rodriguez L.J., Srivastava P., Peraza-Reyes L., Upadhyaya K., Boldogh I.R., Pon L.A. Rad53 is essential for a mitochondrial DNA inheritance checkpoint regulating G1 to S progression. J. Cell Biol. 2012;198:793–798. doi: 10.1083/jcb.201205193. [PMC free article] [PubMed] [Cross Ref]
83. Futami K., Shimamoto A., Furuichi Y. Mitochondrial and nuclear localization of human Pif1 helicase. Biol. Pharm. Bull. 2007;30:1685–1692. doi: 10.1248/bpb.30.1685. [PubMed] [Cross Ref]
84. Sanders C.M. Human Pif1 helicase is a G-quadruplex DNA-binding protein with G-quadruplex DNA-unwinding activity. Biochem. J. 2010;430:119–128. doi: 10.1042/BJ20100612. [PubMed] [Cross Ref]
85. Bochman M.L., Sabouri N., Zakian V.A. Unwinding the functions of the Pif1 family helicases. DNA Repair (Amst) 2010;9:237–249. doi: 10.1016/j.dnarep.2010.01.008. [PMC free article] [PubMed] [Cross Ref]
86. Ding L., Liu Y. Borrowing nuclear DNA helicases to protect mitochondrial DNA. Int. J. Mol. Sci. 2015;16:10870–10887. doi: 10.3390/ijms160510870. [PMC free article] [PubMed] [Cross Ref]
87. Anand R.P., Lovett S.T., Haber J.E. Break-induced DNA replication. Cold Spring Harb. Perspect. Biol. 2013;5:a010397. doi: 10.1101/cshperspect.a010397. [PMC free article] [PubMed] [Cross Ref]
88. Asai T., Sommer S., Bailone A., Kogoma T. Homologous recombination-dependent initiation of DNA replication from DNA damage-inducible origins in Escherichia coli. EMBO J. 1993;12:3287–3295. [PubMed]
89. Asai T., Bates D.B., Kogoma T. DNA replication triggered by double-stranded breaks in E. coli: Dependence on homologous recombination functions. Cell. 1994;78:1051–1061. doi: 10.1016/0092-8674(94)90279-8. [PMC free article] [PubMed] [Cross Ref]
90. Kogoma T. Stable DNA replication: Interplay between DNA replication, homologous recombination, and transcription. Microbiol. Mol. Biol. Rev. 1997;61:212–238. [PMC free article] [PubMed]
91. Kuzminov A., Stahl F.W. Double-strand end repair via the RecBC pathway in Escherichia coli primes DNA replication. Genes Dev. 1999;13:345–356. doi: 10.1101/gad.13.3.345. [PubMed] [Cross Ref]
92. Magee T.R., Kogoma T. Requirement of RecBC enzyme and an elevated level of activated RecA for induced stable DNA replication in Escherichia coli. J. Bacteriol. 1990;172:1834–1839. doi: 10.1128/jb.172.4.1834-1839.1990. [PMC free article] [PubMed] [Cross Ref]
93. Liu J., Xu L., Sandler S.J., Marians K.J. Replication fork assembly at recombination intermediates is required for bacterial growth. Proc. Natl. Acad. Sci. USA. 1999;96:3552–3555. doi: 10.1073/pnas.96.7.3552. [PubMed] [Cross Ref]
94. Gabbai C.B., Marians K.J. Recruitment to stalled replication forks of the PriA DNA helicase and replisome-loading activities is essential for survival. DNA Repair (Amst) 2010;9:202–209. doi: 10.1016/j.dnarep.2009.12.009. [PMC free article] [PubMed] [Cross Ref]
95. Asai T., Imai M., Kogoma T. DNA damage-inducible replication of the Escherichia coli chromosome is initiated at separable sites within the minimal oriC. J. Mol. Biol. 1994;235:1459–1469. doi: 10.1006/jmbi.1994.1101. [PubMed] [Cross Ref]
96. Voelkel-Meiman K., Roeder G.S. Gene conversion tracts stimulated by HOT1-promoted transcription are long and continuous. Genetics. 1990;126:851–867. [PubMed]
97. Morrow D.M., Connelly C., Hieter P. “Break copy” duplication: A model for chromosome fragment formation in Saccharomyces cerevisiae. Genetics. 1997;147:371–382. [PubMed]
98. Davis A.P., Symington L.S. RAD51-dependent break-induced replication in yeast. Mol. Cell. Biol. 2004;24:2344–2351. doi: 10.1128/MCB.24.6.2344-2351.2004. [PMC free article] [PubMed] [Cross Ref]
99. Lydeard J.R., Jain S., Yamaguchi M., Haber J.E. Break-induced replication and telomerase-independent telomere maintenance require Pol32. Nature. 2007;448:820–823. doi: 10.1038/nature06047. [PubMed] [Cross Ref]
100. Kraus E., Leung W.Y., Haber J.E. Break-induced replication: A review and an example in budding yeast. Proc. Natl. Acad. Sci. USA. 2001;98:8255–8262. doi: 10.1073/pnas.151008198. [PubMed] [Cross Ref]
101. Malkova A., Ira G. Break-induced replication: Functions and molecular mechanism. Curr. Opin. Genet. Dev. 2013;23:271–279. doi: 10.1016/j.gde.2013.05.007. [PMC free article] [PubMed] [Cross Ref]
102. Malkova A., Naylor M.L., Yamaguchi M., Ira G., Haber J.E. RAD51-dependent break-induced replication differs in kinetics and checkpoint responses from RAD51-mediated gene conversion. Mol. Cell. Biol. 2005;25:933–944. doi: 10.1128/MCB.25.3.933-944.2005. [PMC free article] [PubMed] [Cross Ref]
103. Malkova A., Signon L., Schaefer C.B., Naylor M.L., Theis J.F., Newlon C.S., Haber J.E. RAD51-independent break-induced replication to repair a broken chromosome depends on a distant enhancer site. Genes Dev. 2001;15:1055–1060. doi: 10.1101/gad.875901. [PubMed] [Cross Ref]
104. Godin S.K., Sullivan M.R., Bernstein K.A. Novel insights into RAD51 activity and regulation during homologous recombination and DNA replication. Biochem. Cell Biol. 2016;94:407–418. doi: 10.1139/bcb-2016-0012. [PMC free article] [PubMed] [Cross Ref]
105. Ira G., Haber J.E. Characterization of RAD51-independent break-induced replication that acts preferentially with short homologous sequences. Mol. Cell. Biol. 2002;22:6384–6392. doi: 10.1128/MCB.22.18.6384-6392.2002. [PMC free article] [PubMed] [Cross Ref]
106. Verma P., Greenberg R.A. Noncanonical views of homology-directed DNA repair. Genes Dev. 2016;30:1138–1154. doi: 10.1101/gad.280545.116. [PubMed] [Cross Ref]
107. Donnianni R.A., Symington L.S. Break-induced replication occurs by conservative DNA synthesis. Proc. Natl. Acad. Sci. USA. 2013;110:13475–13480. doi: 10.1073/pnas.1309800110. [PubMed] [Cross Ref]
108. Jain S., Sugawara N., Lydeard J., Vaze M., Tanguy Le Gac N., Haber J.E. A recombination execution checkpoint regulates the choice of homologous recombination pathway during DNA double-strand break repair. Genes Dev. 2009;23:291–303. doi: 10.1101/gad.1751209. [PubMed] [Cross Ref]
109. McEachern M.J., Haber J.E. Break-induced replication and recombinational telomere elongation in yeast. Annu. Rev. Biochem. 2006;75:111–135. doi: 10.1146/annurev.biochem.74.082803.133234. [PubMed] [Cross Ref]
110. Lydeard J.R., Lipkin-Moore Z., Sheu Y.J., Stillman B., Burgers P.M., Haber J.E. Break-induced replication requires all essential DNA replication factors except those specific for pre-RC assembly. Genes Dev. 2010;24:1133–1144. doi: 10.1101/gad.1922610. [PubMed] [Cross Ref]
111. Wilson M.A., Kwon Y., Xu Y., Chung W.H., Chi P., Niu H., Mayle R., Chen X., Malkova A., Sung P., et al. Pif1 helicase and Polδ promote recombination-coupled DNA synthesis via bubble migration. Nature. 2013;502:393–396. doi: 10.1038/nature12585. [PMC free article] [PubMed] [Cross Ref]
112. Saini N., Ramakrishnan S., Elango R., Ayyar S., Zhang Y., Deem A., Ira G., Haber J.E., Lobachev K.S., Malkova A. Migrating bubble during break-induced replication drives conservative DNA synthesis. Nature. 2013;502:389–392. doi: 10.1038/nature12584. [PMC free article] [PubMed] [Cross Ref]
113. Fangman W.L., Brewer B.J. Activation of replication origins within yeast chromosomes. Annu. Rev. Cell Biol. 1991;7:375–402. doi: 10.1146/annurev.cb.07.110191.002111. [PubMed] [Cross Ref]
114. Yang Y., Sterling J., Storici F., Resnick M.A., Gordenin D.A. Hypermutability of damaged single-strand DNA formed at double-strand breaks and uncapped telomeres in yeast Saccharomyces cerevisiae. PLoS Genet. 2008;4:e1000264 doi: 10.1371/journal.pgen.1000264. [PMC free article] [PubMed] [Cross Ref]
115. Smith C.E., Llorente B., Symington L.S. Template switching during break-induced replication. Nature. 2007;447:102–105. doi: 10.1038/nature05723. [PubMed] [Cross Ref]
116. Llorente B., Smith C.E., Symington L.S. Break-induced replication: What is it and what is it for? Cell Cycle. 2008;7:859–864. doi: 10.4161/cc.7.7.5613. [PubMed] [Cross Ref]
117. Mayle R., Campbell I.M., Beck C.R., Yu Y., Wilson M., Shaw C.A., Bjergbaek L., Lupski J.R., Ira G. DNA REPAIR. Mus81 and converging forks limit the mutagenicity of replication fork breakage. Science. 2015;349:742–747. doi: 10.1126/science.aaa8391. [PMC free article] [PubMed] [Cross Ref]
118. Pardo B., Aguilera A. Complex chromosomal rearrangements mediated by break-induced replication involve structure-selective endonucleases. PLoS Genet. 2012;8:e1002979 doi: 10.1371/journal.pgen.1002979. [PMC free article] [PubMed] [Cross Ref]
119. Lovett S.T. Connecting replication and recombination. Mol. Cell. 2003;11:554–556. doi: 10.1016/S1097-2765(03)00110-2. [PubMed] [Cross Ref]
120. Heller R.C., Marians K.J. Replisome assembly and the direct restart of stalled replication forks. Nat. Rev. Mol. Cell Biol. 2006;7:932–943. doi: 10.1038/nrm2058. [PubMed] [Cross Ref]
121. Sandler S.J., Marians K.J. Role of PriA in replication fork reactivation in Escherichia coli. J. Bacteriol. 2000;182:9–13. doi: 10.1128/JB.182.1.9-13.2000. [PMC free article] [PubMed] [Cross Ref]
122. Wei L., Zhao X. A new MCM modification cycle regulates DNA replication initiation. Nat. Struct. Mol. Biol. 2016;23:209–216. doi: 10.1038/nsmb.3173. [PMC free article] [PubMed] [Cross Ref]
123. Maric M., Maculins T., De Piccoli G., Labib K. Cdc48 and a ubiquitin ligase drive disassembly of the CMG helicase at the end of DNA replication. Science. 2014;346:1253596. doi: 10.1126/science.1253596. [PMC free article] [PubMed] [Cross Ref]
124. Moreno S.P., Bailey R., Campion N., Herron S., Gambus A. Polyubiquitylation drives replisome disassembly at the termination of DNA replication. Science. 2014;346:477–481. doi: 10.1126/science.1253585. [PubMed] [Cross Ref]
125. Makovets S., Blackburn E.H. DNA damage signalling prevents deleterious telomere addition at DNA breaks. Nat. Cell Biol. 2009;11:1383–1386. doi: 10.1038/ncb1985. [PMC free article] [PubMed] [Cross Ref]
126. Vasianovich Y., Harrington L.A., Makovets S. Break-induced replication requires DNA damage-induced phosphorylation of Pif1 and leads to telomere lengthening. PLoS Genet. 2014;10:e1004679 doi: 10.1371/journal.pgen.1004679. [PMC free article] [PubMed] [Cross Ref]
127. Aguilera A., Garcia-Muse T. Causes of genome instability. Annu. Rev. Genet. 2013;47:1–32. doi: 10.1146/annurev-genet-111212-133232. [PubMed] [Cross Ref]
128. Costantino L., Koshland D. The Yin and Yang of R-loop biology. Curr. Opin. Cell Biol. 2015;34:39–45. doi: 10.1016/j.ceb.2015.04.008. [PMC free article] [PubMed] [Cross Ref]
129. Hamperl S., Cimprich K.A. The contribution of co-transcriptional RNA:DNA hybrid structures to DNA damage and genome instability. DNA Repair (Amst) 2014;19:84–94. doi: 10.1016/j.dnarep.2014.03.023. [PMC free article] [PubMed] [Cross Ref]
130. Groh M., Gromak N. Out of balance: R-loops in human disease. PLoS Genet. 2014;10:e1004630 doi: 10.1371/journal.pgen.1004630. [PMC free article] [PubMed] [Cross Ref]
131. Santos-Pereira J.M., Aguilera A. R loops: New modulators of genome dynamics and function. Nat. Rev. Genet. 2015;16:583–597. doi: 10.1038/nrg3961. [PubMed] [Cross Ref]
132. Skourti-Stathaki K., Kamieniarz-Gdula K., Proudfoot N.J. R-loops induce repressive chromatin marks over mammalian gene terminators. Nature. 2014;516:436–439. doi: 10.1038/nature13787. [PMC free article] [PubMed] [Cross Ref]
133. Gaillard H., Aguilera A. Transcription as a threat to genome integrity. Annu. Rev. Biochem. 2016;85:291–317. doi: 10.1146/annurev-biochem-060815-014908. [PubMed] [Cross Ref]
134. Dasgupta S., Masukata H., Tomizawa J. Multiple mechanisms for initiation of ColE1 DNA replication: DNA synthesis in the presence and absence of ribonuclease H. Cell. 1987;51:1113–1122. doi: 10.1016/0092-8674(87)90597-6. [PubMed] [Cross Ref]
135. Masukata H., Dasgupta S., Tomizawa J. Transcriptional activation of ColE1 DNA synthesis by displacement of the nontranscribed strand. Cell. 1987;51:1123–1130. doi: 10.1016/0092-8674(87)90598-8. [PubMed] [Cross Ref]
136. Marians K.J. Prokaryotic DNA replication. Annu. Rev. Biochem. 1992;61:673–719. doi: 10.1146/annurev.bi.61.070192.003325. [PubMed] [Cross Ref]
137. Mosig G. The essential role of recombination in phage T4 growth. Annu. Rev. Genet. 1987;21:347–371. doi: 10.1146/annurev.ge.21.120187.002023. [PubMed] [Cross Ref]
138. de Massy B., Fayet O., Kogoma T. Multiple origin usage for DNA replication in sdrA(rnh) mutants of Escherichia coli K-12. Initiation in the absence of oriC. J. Mol. Biol. 1984;178:227–236. doi: 10.1016/0022-2836(84)90141-4. [PubMed] [Cross Ref]
139. Kogoma T. Absence of RNase H allows replication of pBR322 in Escherichia coli mutants lacking DNA polymerase I. Proc. Natl. Acad. Sci. USA. 1984;81:7845–7849. doi: 10.1073/pnas.81.24.7845. [PubMed] [Cross Ref]
140. Von Meyenburg K., Boye E., Skarstad K., Koppes L., Kogoma T. Mode of initiation of constitutive stable DNA replication in RNase H-defective mutants of Escherichia coli K-12. J. Bacteriol. 1987;169:2650–2658. doi: 10.1128/jb.169.6.2650-2658.1987. [PMC free article] [PubMed] [Cross Ref]
141. Horiuchi T., Maki H., Sekiguchi M. RNase H-defective mutants of Escherichia coli: A possible discriminatory role of RNase H in initiation of DNA replication. Mol. Gen. Genet. 1984;195:17–22. doi: 10.1007/BF00332717. [PubMed] [Cross Ref]
142. Ogawa T., Pickett G.G., Kogoma T., Kornberg A. RNase H confers specificity in the DnaA-dependent initiation of replication at the unique origin of the Escherichia coli chromosome in vivo and in vitro. Proc. Natl. Acad. Sci. USA. 1984;81:1040–1044. doi: 10.1073/pnas.81.4.1040. [PubMed] [Cross Ref]
143. Maduike N.Z., Tehranchi A.K., Wang J.D., Kreuzer K.N. Replication of the Escherichia coli chromosome in RNase HI-deficient cells: Multiple initiation regions and fork dynamics. Mol. Microbiol. 2014;91:39–56. doi: 10.1111/mmi.12440. [PMC free article] [PubMed] [Cross Ref]
144. Dimude J.U., Stockum A., Midgley-Smith S.L., Upton A.L., Foster H.A., Khan A., Saunders N.J., Retkute R., Rudolph C.J. The consequences of replicating in the wrong orientation: Bacterial chromosome duplication without an active replication origin. MBio. 2015;6:e01294-15. doi: 10.1128/mBio.01294-15. [PMC free article] [PubMed] [Cross Ref]
145. Hong X., Cadwell G.W., Kogoma T. Escherichia coli RecG and RecA proteins in R-loop formation. EMBO J. 1995;14:2385–2392. [PubMed]
146. Hong X., Cadwell G.W., Kogoma T. Activation of stable DNA replication in rapidly growing Escherichia coli at the time of entry to stationary phase. Mol. Microbiol. 1996;21:953–961. doi: 10.1046/j.1365-2958.1996.591419.x. [PubMed] [Cross Ref]
147. Drolet M., Phoenix P., Menzel R., Masse E., Liu L.F., Crouch R.J. Overexpression of RNase H partially complements the growth defect of an Escherichia coli delta topA mutant: R-loop formation is a major problem in the absence of DNA topoisomerase I. Proc. Natl. Acad. Sci. USA. 1995;92:3526–3530. doi: 10.1073/pnas.92.8.3526. [PubMed] [Cross Ref]
148. Martel M., Balleydier A., Sauriol A., Drolet M. Constitutive stable DNA replication in Escherichia coli cells lacking type 1A topoisomerase activity. DNA Repair (Amst) 2015;35:37–47. doi: 10.1016/j.dnarep.2015.08.004. [PubMed] [Cross Ref]
149. Sandler S.J. Requirements for replication restart proteins during constitutive stable DNA replication in Escherichia coli K-12. Genetics. 2005;169:1799–1806. doi: 10.1534/genetics.104.036962. [PubMed] [Cross Ref]
150. Usongo V., Drolet M. Roles of type 1A topoisomerases in genome maintenance in Escherichia coli. PLoS Genet. 2014;10:e1004543 doi: 10.1371/journal.pgen.1004543. [PMC free article] [PubMed] [Cross Ref]
151. Kasahara M., Clikeman J.A., Bates D.B., Kogoma T. RecA protein-dependent R-loop formation in vitro. Genes Dev. 2000;14:360–365. [PubMed]
152. Zaitsev E.N., Kowalczykowski S.C. A novel pairing process promoted by Escherichia coli RecA protein: Inverse DNA and RNA strand exchange. Genes Dev. 2000;14:740–749. [PubMed]
153. Wimberly H., Shee C., Thornton P.C., Sivaramakrishnan P., Rosenberg S.M., Hastings P.J. R-loops and nicks initiate DNA breakage and genome instability in non-growing Escherichia coli. Nat. Commun. 2013;4:2115. doi: 10.1038/ncomms3115. [PMC free article] [PubMed] [Cross Ref]
154. Usongo V., Martel M., Balleydier A., Drolet M. Mutations reducing replication from R-loops suppress the defects of growth, chromosome segregation and DNA supercoiling in cells lacking topoisomerase I and RNase HI activity. DNA Repair (Amst) 2016;40:1–17. doi: 10.1016/j.dnarep.2016.02.001. [PubMed] [Cross Ref]
155. El Hage A., Webb S., Kerr A., Tollervey D. Genome-wide distribution of RNA-DNA hybrids identifies RNase H targets in tRNA genes, retrotransposons and mitochondria. PLoS Genet. 2014;10:e1004716 doi: 10.1371/journal.pgen.1004716. [PMC free article] [PubMed] [Cross Ref]
156. Chan Y.A., Aristizabal M.J., Lu P.Y., Luo Z., Hamza A., Kobor M.S., Stirling P.C., Hieter P. Genome-wide profiling of yeast DNA:RNA hybrid prone sites with DRIP-chip. PLoS Genet. 2014;10:e1004288 doi: 10.1371/journal.pgen.1004288. [PMC free article] [PubMed] [Cross Ref]
157. Mischo H.E., Gomez-Gonzalez B., Grzechnik P., Rondon A.G., Wei W., Steinmetz L., Aguilera A., Proudfoot N.J. Yeast Sen1 helicase protects the genome from transcription-associated instability. Mol. Cell. 2011;41:21–32. doi: 10.1016/j.molcel.2010.12.007. [PMC free article] [PubMed] [Cross Ref]
158. Alzu A., Bermejo R., Begnis M., Lucca C., Piccini D., Carotenuto W., Saponaro M., Brambati A., Cocito A., Foiani M., et al. Senataxin associates with replication forks to protect fork integrity across RNA-polymerase-II-transcribed genes. Cell. 2012;151:835–846. doi: 10.1016/j.cell.2012.09.041. [PMC free article] [PubMed] [Cross Ref]
159. Salvi J.S., Chan J.N., Szafranski K., Liu T.T., Wu J.D., Olsen J.B., Khanam N., Poon B.P., Emili A., Mekhail K. Roles for Pbp1 and caloric restriction in genome and lifespan maintenance via suppression of RNA-DNA hybrids. Dev. Cell. 2014;30:177–191. doi: 10.1016/j.devcel.2014.05.013. [PubMed] [Cross Ref]
160. El Hage A., French S.L., Beyer A.L., Tollervey D. Loss of Topoisomerase I leads to R-loop-mediated transcriptional blocks during ribosomal RNA synthesis. Genes Dev. 2010;24:1546–1558. doi: 10.1101/gad.573310. [PubMed] [Cross Ref]
161. Stuckey R., Garcia-Rodriguez N., Aguilera A., Wellinger R.E. Role for RNA:DNA hybrids in origin-independent replication priming in a eukaryotic system. Proc. Natl. Acad. Sci. USA. 2015;112:5779–5784. doi: 10.1073/pnas.1501769112. [PubMed] [Cross Ref]
162. Pomerantz R.T., O’Donnell M. The replisome uses mRNA as a primer after colliding with RNA polymerase. Nature. 2008;456:762–766. doi: 10.1038/nature07527. [PMC free article] [PubMed] [Cross Ref]
163. Pomerantz R.T., O’Donnell M. Direct restart of a replication fork stalled by a head-on RNA polymerase. Science. 2010;327:590–592. doi: 10.1126/science.1179595. [PMC free article] [PubMed] [Cross Ref]
164. Longhese M.P., Plevani P., Lucchini G. Replication factor A is required in vivo for DNA replication, repair, and recombination. Mol. Cell. Biol. 1994;14:7884–7890. doi: 10.1128/MCB.14.12.7884. [PMC free article] [PubMed] [Cross Ref]
165. Bartrand A.J., Iyasu D., Brush G.S. DNA stimulates Mec1-mediated phosphorylation of replication protein A. J. Biol. Chem. 2004;279:26762–26767. doi: 10.1074/jbc.M312353200. [PubMed] [Cross Ref]
166. Foiani M., Liberi G., Lucchini G., Plevani P. Cell cycle-dependent phosphorylation and dephosphorylation of the yeast DNA polymerase alpha-primase B subunit. Mol. Cell. Biol. 1995;15:883–891. doi: 10.1128/MCB.15.2.883. [PMC free article] [PubMed] [Cross Ref]
167. Rippe K., Luke B. TERRA and the state of the telomere. Nat. Struct. Mol. Biol. 2015;22:853–858. doi: 10.1038/nsmb.3078. [PubMed] [Cross Ref]
168. Balk B., Maicher A., Dees M., Klermund J., Luke-Glaser S., Bender K., Luke B. Telomeric RNA-DNA hybrids affect telomere-length dynamics and senescence. Nat. Struct. Mol. Biol. 2013;20:1199–1205. doi: 10.1038/nsmb.2662. [PubMed] [Cross Ref]
169. Arora R., Lee Y., Wischnewski H., Brun C.M., Schwarz T., Azzalin C.M. RNase H1 regulates TERRA-telomeric DNA hybrids and telomere maintenance in ALT tumour cells. Nat. Commun. 2014;5:5220. doi: 10.1038/ncomms6220. [PMC free article] [PubMed] [Cross Ref]
170. Flynn R.L., Cox K.E., Jeitany M., Wakimoto H., Bryll A.R., Ganem N.J., Bersani F., Pineda J.R., Suva M.L., Benes C.H., et al. Alternative lengthening of telomeres renders cancer cells hypersensitive to ATR inhibitors. Science. 2015;347:273–277. doi: 10.1126/science.1257216. [PMC free article] [PubMed] [Cross Ref]
171. Law M.J., Lower K.M., Voon H.P., Hughes J.R., Garrick D., Viprakasit V., Mitson M., De Gobbi M., Marra M., Morris A., et al. ATR-X syndrome protein targets tandem repeats and influences allele-specific expression in a size-dependent manner. Cell. 2010;143:367–378. doi: 10.1016/j.cell.2010.09.023. [PubMed] [Cross Ref]
172. Kogoma T., Lark K.G. Characterization of the replication of Escherichia coli DNA in the absence of protein synthesis: Stable DNA replication. J. Mol. Biol. 1975;94:243–256. doi: 10.1016/0022-2836(75)90081-9. [PubMed] [Cross Ref]
173. Lombrana R., Almeida R., Alvarez A., Gomez M. R-loops and initiation of DNA replication in human cells: A missing link? Front. Genet. 2015;6:158. doi: 10.3389/fgene.2015.00158. [PMC free article] [PubMed] [Cross Ref]
174. Leonard A.C., Mechali M. DNA replication origins. Cold Spring Harb. Perspect. Biol. 2013;5:a010116. doi: 10.1101/cshperspect.a010116. [PMC free article] [PubMed] [Cross Ref]
175. Norseen J., Thomae A., Sridharan V., Aiyar A., Schepers A., Lieberman P.M. RNA-dependent recruitment of the origin recognition complex. EMBO J. 2008;27:3024–3035. doi: 10.1038/emboj.2008.221. [PubMed] [Cross Ref]
176. Gomez M., Antequera F. Overreplication of short DNA regions during S phase in human cells. Genes Dev. 2008;22:375–385. doi: 10.1101/gad.445608. [PubMed] [Cross Ref]
177. Lombrana R., Alvarez A., Fernandez-Justel J.M., Almeida R., Poza-Carrion C., Gomes F., Calzada A., Requena J.M., Gomez M. Transcriptionally driven DNA replication program of the human parasite Leishmania major. Cell Rep. 2016;16:1774–1786. doi: 10.1016/j.celrep.2016.07.007. [PubMed] [Cross Ref]

Articles from Genes are provided here courtesy of Multidisciplinary Digital Publishing Institute (MDPI)