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Genes 2017, 8(4), 112; https://doi.org/10.3390/genes8040112

Review
DNA Replication Origins and Fork Progression at Mammalian Telomeres
Department of Cellular Biochemistry, Graduate School of Pharmaceutical Sciences, Kyushu University, 3-1-1 Maidashi, Higashi-ku, Fukuoka 812-8582, Japan
*
Authors to whom correspondence should be addressed.
Academic Editor: Eishi Noguchi
Received: 7 February 2017 / Accepted: 24 March 2017 / Published: 28 March 2017

Abstract

:
Telomeres are essential chromosomal regions that prevent critical shortening of linear chromosomes and genomic instability in eukaryotic cells. The bulk of telomeric DNA is replicated by semi-conservative DNA replication in the same way as the rest of the genome. However, recent findings revealed that replication of telomeric repeats is a potential cause of chromosomal instability, because DNA replication through telomeres is challenged by the repetitive telomeric sequences and specific structures that hamper the replication fork. In this review, we summarize current understanding of the mechanisms by which telomeres are faithfully and safely replicated in mammalian cells. Various telomere-associated proteins ensure efficient telomere replication at different steps, such as licensing of replication origins, passage of replication forks, proper fork restart after replication stress, and dissolution of post-replicative structures. In particular, shelterin proteins have central roles in the control of telomere replication. Through physical interactions, accessory proteins are recruited to maintain telomere integrity during DNA replication. Dormant replication origins and/or homology-directed repair may rescue inappropriate fork stalling or collapse that can cause defects in telomere structure and functions.
Keywords:
DNA replication; genome integrity; telomere; shelterin; G-quadruplex; RecQ-like helicase; fragile telomere; replication fork barrier; dormant origin

1. Introduction

In eukaryotic cells, protection of the ends of linear chromosomes depends on specialized nucleoprotein structures known as telomeres, which function as buffers for the shortening of linear chromosomes during each round of semi-conservative DNA replication and prevent activation of DNA damage responses, such as the ATM and ATR checkpoint signaling, classical and alternative non-homologous end joining pathways, and homologous recombination repair [1,2,3,4]. Vertebrate telomeric DNA consists of thousands of tandem 5′-TTAGGG-3′ repeats [5]. In contrast to the small telomeres of yeasts that consist of several hundred base pairs, human telomeres are typically 10–15 kb in length, and those of mice are 20–50 kb [1]. The telomeric repeat array is bound by the shelterin protein complex that is composed of telomeric repeat-binding factor 1 and 2 (TRF1 and TRF2), repressor/activator protein 1 (RAP1), TRF1-interacting nuclear protein 2 (TIN2), protection of telomeres protein 1 (POT1), and POT1- and TIN2-interacting protein TPP1 (TINT1/PTOP/PIP1) [6]. The repeat array terminates in a single-stranded 3′ protrusion of the G-rich strand (referred to as a G-overhang). The chromosome ends are stabilized by the formation of a protective loop structure, called a T-loop (telomere loop), in which the G-overhang presumably loops back and invades the double-stranded region of telomeric DNA [7,8]. Telomeres thereby prevent chromosome ends from inappropriate recognition by DNA damage signaling and repair systems [2]. In addition, several conserved features of telomeres, such as constitutive heterochromatin, G-quadruplex (G4) DNA secondary structure, and transcription of the non-coding telomeric repeat-containing RNA (TERRA), are also involved in the regulation of telomere capping and maintenance [9,10,11,12,13].
The majority of telomeric double-stranded DNA repeats are replicated in a semi-conservative manner by conventional DNA replication machinery [14]. However, characteristic features of telomeres represent intrinsic replication fork barriers that induce stalling and/or collapse of replication machinery [3,4]. Failure of telomeric DNA replication can cause genomic instability, which in turn promotes cellular transformation or senescence [15]. Here, we summarize the recent advances in our understanding of the mechanisms that support efficient DNA replication at mammalian telomeres, with a focus on the functional interactions between shelterin components and a variety of accessory proteins that enable the replication machinery to reach the chromosomal termini.

2. Replication Origins for the Duplication of Telomeric DNA

2.1. General Regulation of Eukaryotic DNA Replication; Origin Licensing and Firing

The accurate DNA replication of eukaryotic genomes relies on strict temporal separation of chromatin loading of a replicative helicase (so-called origin licensing) from its activation followed by DNA synthesis (so-called origin firing) (Figure 1) [16,17]. In the late M to G1 phases, the MCM2–7 helicase complex is recruited onto chromatin in an inactive form in a process that is dependent on the origin-recognition complex (ORC), cell division cycle protein 6 (CDC6), and DNA replication licensing factor Cdt1 [18,19]. This step is also referred to as pre-replication complex (pre-RC) formation. In the subsequent S phase, DBF4-dependent kinase (DDK) and cyclin-dependent kinases (CDKs) trigger the recruitment of additional replication proteins to the origins, leading to the remodeling of inactive MCM2–7 complexes to active CMG (CDC45–MCM–GINS) replicative helicase complexes, and to the initiation of DNA synthesis at bidirectional replication forks [18,20,21]. According to a recent model, DNA polymerase α (Pol α) and primase complex initiate DNA synthesis, and Polδ and Polε continue lagging and leading DNA strand synthesis, respectively [22]. MCM2–7 loading is strictly inhibited after the onset of S phase through a number of redundant mechanisms, thereby preventing re-replication of the genome [23].
Positioning of sites for binding of ORC and MCM2–7 in the G1 phase is a key regulator of the chromosome-replication program, in which multiple replication-initiation sites (replication origins) are distributed along chromosomes [24,25,26,27,28,29,30,31]. Ideally, bidirectional replication forks should continue along a chromosome until they meet forks coming from adjacent origins, or they reach the end of the chromosome. However, replication forks often pause and collapse because they encounter obstacles, such as damaged DNA, interstrand DNA cross-links, or DNA-RNA hybrids that form R-loop structures, or because of exhaustion of dNTPs or of the single-stranded DNA (ssDNA)-binding protein RPA [15,32]. Because reloading of MCM2–7 in the S phase should not occur, so-called dormant origins (backup pre-RCs formed in G1 phase but not used in normal S phase) are reserved to complete genome replication in conditions of replication stress [15,33,34,35,36]. The DNA-replication-checkpoint pathway coordinates multiple mechanisms, including cell cycle arrest, protection and restart of stalled forks, and activation of dormant origins, to maintain genome integrity [37,38].

2.2. Replication Origins for Duplication of Telomeric DNA

In contrast to yeast telomeres, which are replicated in late S phase, human telomeres are duplicated throughout S phase [39,40,41,42,43]. Timing of the replication of human telomeres is specific for each chromosome arm and is dependent on subtelomeric elements, although the mechanism for this regulation is still unclear [41,42,44]. Unlike yeast cells, in which the telomeric protein Rif1 negatively regulates subtelomeric origin firing, mammalian Rif1 is not localized to telomeres and therefore may not play a role in regulation of telomeric DNA replication [45]. Single-molecule DNA-fiber analysis has enabled identification of replication origins labeled with thymidine analogs around telomeres in mouse and human cells [46,47,48]. Similar to the origin distribution in yeasts [49,50,51], origins are frequently found in mammalian subtelomeric regions. Moreover, in some cases, replication initiates within the telomeric repeats themselves. The results of nascent-strand sequencing (NS-seq) experiments also suggest that, even after normalizing for λ-exonuclease bias, human telomeric DNA is enriched in the sequences of actual firing origins [52].
Telomeres challenge the progression of replication machinery. Telomeric origins may function as a backup system that is needed to ensure completion of telomeric DNA replication. When a replication fork collapses within a telomere, additional origin activation could prevent telomere loss resulting from a large unreplicated region [15,33]. The genomic regions called common fragile sites are frequently broken upon replication stress. The chromosomal fragility is associated with the origin-poor regions of genomes [24,53,54]. It also stems from DNA secondary structures, collision with transcription of large genes, or condensed chromatin structures, all interfering with progression of replication fork. Defects in telomere replication similarly lead to chromosomal fragility [48,55,56,57,58], suggesting that origins in telomeric regions may be important for genome stability.

2.3. Mechanisms Promoting Pre-RC Formation on Telomeric DNA

Results from several studies demonstrate active ORC binding and pre-RC formation within TTAGGG repeats [59,60,61,62,63,64], and the shelterin component TRF2, which is essential for telomere capping, has been implicated in origin licensing through physical interaction with the largest ORC subunit, ORC1 [59,60,61,65,66]. TRF2 knockdown reduces ORC binding and pre-RC formation on telomeric DNA [60,61,63]. The TRFH (TRF homology) dimerization domain of TRF2, but not a mutant domain defective in dimerization, recruits ORC and pre-RC to chromatin [66]. This dimerization domain also interacts with proteins that are involved in telomere maintenance, such as 5′ exonuclease Apollo, structure-specific endonuclease subunit SLX4, regulator of telomere elongation helicase 1 (RTEL1), and RAP1 [67,68,69,70,71,72]. An interaction between ORC1 and the basic domain of TRF2 has also been proposed [59,60,65].
Several telomere-specific features may support ORC binding to telomeres. G4 DNA is a non-B-form DNA secondary structure constructed by parallel four-stranded guanine base pairing [73,74]. Systematic genome-wide studies have suggested that G4-motif sequences are associated with replication origins [24,75,76,77,78,79]. In vitro, human ORC1 physically interacts with G4-forming ssDNA and RNA [59,80]. Several lines of evidence support the presence of G4 DNAs at human telomeres [9,81,82,83]. The telomeric C-rich strand is transcribed from the subtelomeric region toward the telomere by RNA Polymerase II to generate TERRA [84,85]. TERRA then interacts with telomeres and is involved in heterochromatin organization and telomere maintenance [10,12,86,87]. TERRA–telomeric DNA hybrids form R-loop structures, which may result in the formation of G4 on the displaced G-rich ssDNA [87,88]. Further research is needed to determine whether these telomeric G4 structures promote ORC recruitment and origin firing.
Telomeric regions (and subtelomeric regions) are highly enriched with repressive epigenetic modifications [12,13]. Heterochromatin proteins that interact with ORC, such as heterochromatin protein 1 (HP1) and ORC-associated protein (ORCA, also known as LRWD1), might be involved in the regulation of telomeric replication origins [89]. ORCA localizes to heterochromatic sites including telomeres, and functions in the regulation of replication licensing through interactions with ORC, Cdt1, and geminin in a cell cycle-dependent manner [89,90,91,92,93]. Among repressive modifications of telomeres, the trimethylated lysine 20 of histone H4 (H4K20me3) is associated with ORC recruitment to replication origins [94,95]. The methyltransferase PR-Set7 (also known as Set8 and KMT5a) catalyzes H4K20 monomethylation, while other methyltransferases Suv4-20h1 and Suv4-20h2 are responsible for the transition from H4K20me1 to H4K20me2/3 [96,97,98]. Ectopic tethering of PR-Set7 promotes trimethylation and loading of ORC in a manner that is dependent on Suv4-20h1 [92,94]. Although the BAH (bromo-adjacent homology) domain of ORC1 preferentially interacts with a H4K20me2 peptide [92,99], ORC complexed with ORCA is thought to interact with H4K20me3 [92,93]. H4K20me3 is highly enriched at telomeres and other transcriptionally silenced regions [100,101,102], but the roles of this modification in telomeric replication remain to be established.

3. Shelterin and Additional Proteins that Support Telomeric DNA Replication

3.1. Telomeric Obstacles Against Passage of Replication Forks

Eukaryotic genome integrity is maintained by protecting telomeres from various problems caused by their terminal position. Incomplete lagging-strand synthesis at the chromosomal termini causes gradual loss of genetic information. The iterative telomerase action or a homologous recombination-mediated mechanism, called Alternative Lengthening of Telomeres (ALT), is therefore needed to extend and maintain the repetitive TTAGGG sequences [14,103]. Moreover, the protective shelterin complex prevents chromosomal fusions resulting from improper activation of DNA repair pathways [1,2,3]. These mechanisms are essential for genomic stability, but at the same time they cause difficulties in replication. Telomeric repeats impede the replication machinery not only in telomeres, but also in interstitial chromosomal regions that contain the repeats, or when transferred to plasmid DNAs [48,55,104,105,106], suggesting that the replication difficulties can, at least partly, be attributed to the telomeric sequences themselves. Repetitive TTAGGG sequences can form G4 structures that are more stable than the standard B-form DNA duplex, thereby presenting obstacles to the progression of replication forks [9,107] (Figure 2a). Furthermore, G4-independent fork stalling on telomeric G-rich templates has been suggested by the results of in vitro experiments [108]. In addition, protective capping structures formed by shelterin can cause replication impediments (Figure 2a). T-loop structures, as well as telomeric R-loops, DNA topological constraint, and heterochromatin may interfere with replication fork progression if they are not resolved (Figure 2a). Therefore, a number of accessory proteins are required for efficient passage of replication forks through telomeres. Whereas shelterin proteins are potential obstacles to conventional replication forks, because they bind tightly to telomeric chromatin [104,109], evidence now indicates that shelterin components facilitate replication by recruiting additional proteins that resolve other obstacles (Figure 2b). Here, we provide an update of the mechanisms that are known to underlie efficient fork progression through telomeres.

3.2. TRF1 and RecQ-Like Helicases

TRF1, a shelterin component that is not essential for telomere capping, contributes to efficient replication in mammalian telomeres [48,55,56,110]. TRF1 deletion leads to various telomeric defects, including the fragile telomere phenotype, in which FISH (fluorescence in situ hybridization) signals of telomeric probes show multiple foci at single chromosomal termini on metaphase spreads. Although detailed mechanisms of this phenotype remains to be clarified, the multi telomeric signals are thought to be a consequence of replication defects at telomeres and to reflect telomeric DNA breakage or the presence of aberrant, condensed structures. Fragile telomeres are also observed with replication stress induced by low doses of aphidicolin, an inhibitor of DNA polymerases [48,55]. Furthermore, TRF1-deleted cells exhibit activation of the DNA-replication-checkpoint kinase ATR, sister telomere association, and ultrafine anaphase bridges in mitosis, which is consistent with the presence of replication defects [48,55,111,112].
One of the suggested molecular mechanisms for the suppression of fragile telomeres by TRF1 is the recruitment of Bloom syndrome protein (BLM) [48,56], a member of the RecQ-like (RECQL) helicase family [113] that can resolve G4 DNA, D-loop (displacement loop) structures, and Holliday-junction DNA in vitro [114,115,116,117,118]. During DNA replication, G4-forming ssDNA can be produced at telomeres by unwinding of duplex DNA or unfolding of the G-overhang in the T-loop [9]. Although ssDNA-binding proteins such as RPA and POT1 can counteract G4 formation [119,120,121,122,123,124], a single-DNA-molecule-based analysis revealed that deletion of BLM decreases the rate of progression of replication forks inside telomeric tracts, and a G4-stabilizing agent enhances this slowing down of the forks, supporting the idea that BLM promotes telomeric replication by resolving G4 DNA [46]. Indeed, BLM deficiency induces fragile telomere specifically in daughter chromatids derived from G-rich templates, but not from C-rich ones [48,56,58]. In addition to the resolution of G4 during S phase, BLM localizes to telomeres in G2/M and is involved in the processing of late- or post-replicative telomeric structures resulting from both leading- and lagging-strand replication, as well as in T-loop resolution [58,125,126]. BLM acts on ultrafine anaphase bridges, a subset of which originate from telomeric DNA, to resolve these aberrant post-replicative structures that might arise from incomplete replication [58,127]. BLM can bind to basic patches in the hinge domain of TRF1, and a TRF1 variant lacking the BLM-binding patches is defective in the suppression of fragile telomeres in TRF1-deleted cells [56]. Although TRF1 has been suggested to be the major factor in the recruitment of BLM to telomeres, the helicase activity of BLM can be modulated by other shelterin components, such as TRF2 and POT1 [128,129,130].
Similar to BLM, the RECQL Werner syndrome helicase (WRN) has been implicated in resolution of telomeric G4 DNA, D-loops, and Holliday junctions, and its activity is regulated by several shelterin components [114,116,128,129,131,132,133]. The helicase activity of WRN is required for efficient replication of G-rich telomeric DNA, and its deficiency causes loss of the telomeres that use the G-rich strand as a template for synthesis [58,134,135]. Stabilization of G4 DNA perturbs telomere replication and enhances association of WRN and BLM with telomeres [136]. However, unlike BLM, deficiency of WRN does not induce the multi-telomeric signals indicative of fragile telomeres [48]. WRN is thought to be recruited by TRF2 to telomeres in S phase, and is also involved in the control of telomeric recombination events, such as T-loop assembly and disassembly, repression of sister chromatid exchange, and ALT [128,131,135,137,138,139,140]. Overall, WRN and BLM seem to have partially shared but non-redundant functions for the common goal that is complete replication of the chromosome ends.
RECQL helicase 4 (RECQL4) is altered in patients with Rothmund–Thomson syndrome, and cells derived from these patients show telomere fragility [141]. The N-terminal non-catalytic region of RECQL4 has an essential role in the initiation of DNA replication, and is a metazoan homolog of yeast Sld2 (Drc1) [142,143]. RECQL4 localizes to telomeres in S phase, and knockdown of RECQL4 causes telomere dysfunction-induced foci (TIFs) and fragile telomeres. In contrast to BLM and WRN, RECQL4 does not possess G4-unwinding activity in vitro [144], although the N-terminal region binds to G4 structures [145]. Interaction of another RECQL protein, RECQL1, with TRF2 and flap endonuclease 1 (FEN1) has also been proposed to participate in telomere replication [146,147]. In vitro, RECQL1 can resolve D-loops and Holliday junctions, but not G4 DNA, and it displaces TRF1 and TRF2 from telomeric repeats [146,148,149]. However, the detailed molecular mechanisms for how these RECQL helicases maintain telomere integrity during replication are not yet known.

3.3. RTEL1

RTEL1 is a G4-resolving helicase that is involved in telomeric DNA replication [81,150]. RTEL1-knockout mouse embryonic fibroblasts have various chromosomal abnormalities, such as fragile telomeres, telomere circles (extrachromosomal circular DNAs that contain telomeric repeat sequences), and loss of telomere signals [48,57,151,152,153]. RTEL1 contains a PIP (proliferating cell nuclear antigen (PCNA)-interacting protein) box in its C-terminal region [153]. PCNA is a fundamental component of the replication machinery that increases the processivity of DNA polymerases. The PIP box of RTEL1 is required for unwinding of G4 DNAs not only in telomeres but also genome-wide during replication [153]. A PIP box-deleted variant of mouse RTEL1, which is defective in PCNA interaction, fails to rescue the fragile telomere phenotype induced by RTEL1 deletion, but can rescue telomere circles and telomere loss [153], suggesting that RTEL1 has at least two distinct and separable functions for telomere maintenance.
The T-loop structure is essential to protect chromosome ends, but this structure must be unwound and reformed during telomere replication. RTEL1 has been proposed to be a helicase that unwinds the T-loop, in which G-overhang DNA invades the double-stranded telomere [57,151,152]. In vitro, RTEL1 preferentially unwinds a 3′-ssDNA-invaded D-loop (which resembles the structure in the T-loop) in a RPA-dependent manner [154]. Telomere-circle formation and telomere loss in RTEL1-deficient cells support the idea that RTEL1 has a role in T-loop disassembly in vivo [57]. TRF2 is a binding partner of RTEL1 [70], and they interact via the TRFH dimerization domain of TRF2. A mutation that affects the TRFH domain and disrupts the TRF2-RTEL1 interaction leads to telomere-circle formation and telomere loss [70]. In patients with Hoyeraal–Hreidarsson syndrome (a severe variant of dyskeratosis congenita), mutation affects the RTEL1 C4C4 metal-binding motif [150], so that RTEL1 no longer binds to TRF2, and this RTEL1 variant fails to rescue the telomere loss and the telomere circles induced by RTEL1 deletion [70]. Because the C4C4-defective RTEL1 variant can rescue the fragile telomere phenotype, the interaction of RTEL1 with TRF2 seems to be required for proper disassembly of the T-loop, rather than G4-unwinding, preventing loss of the telomere as a circle. Taken together, RTEL1 prevents telomere fragility via interaction with PCNA and facilitates T-loop disassembly via interaction with TRF2. However, TRF2 is also essential for the assembly of the T-loop [7,8,155]. How these contrasting activities of TRF2 are regulated during the cell cycle is not currently known.

3.4. SLX4

Telomere-circle formation and telomere loss in RTEL1-deficient cells are mediated by SLX4 (also known as FANCP or BTBD12), which serves as a scaffold protein for the structure-specific endonucleases SLX1, XPF, and MUS81 [57,156,157,158,159]. The SLX4–endonuclease complex is capable of nucleolytically resolving D-loops and Holliday junctions in vitro [71,126,156,157,158], and is involved in genome-wide resolution of Holliday junctions, and in repair of interstrand DNA cross-links [160,161,162,163]. Deletion of SLX4, SLX1, or XPF, but not MUS81, suppresses telomere-circle formation that is observed in the absence of RTEL1 [57], suggesting that SLX4–endonuclease complexes excise persistent T-loop structures. Furthermore, deletion of SLX4 leads to TIFs and fragile telomeres [72,126,164], suggesting that SLX4-mediated nucleolytic resolution of branched intermediates is required during telomere replication.
In human cells, SLX4 localizes to telomeres throughout the cell cycle via binding to the TRF2 TRFH domain [71,72]. Although SLX4 in mice is involved in telomere maintenance [57,72], the TRF2-binding motif of SLX4 (HxLxP) is conserved in primates, but not in non-primate mammals. The Holliday junction-processing activity of human SLX4 is carefully regulated by TRF1, TRF2, and BLM, preventing inappropriate telomere shortening by T-loop excision and aberrant crossover between telomeric sister chromatids [71,126,164,165]. Recently, SUMO was shown to regulate the function of human SLX4, including TRF2 binding [166,167,168], further contributing to the tight regulation of SLX4 activity for homeostasis of telomere length.

3.5. FEN1 and DNA2

FEN1, a structure-specific endonuclease, is important for proper telomere replication, independent of its general role in Okazaki fragment maturation. FEN1 has been suggested to facilitate telomeric replication by reinitiating stalled replication forks [169,170]. FEN1 depletion leads to a fragile telomere phenotype and to loss of single sister telomeres derived from lagging- or leading-strand replication [169,170,171]. Nuclease activity and interaction with WRN and TRF2 are required for FEN1 to prevent telomere fragility [169,170,171]. Although FEN1 cleaves telomeric G4-containing 5′ flaps in vitro [172], in vivo substrates during telomere replication are unknown [173]. Notably, RNase H1, an endoribonuclease that degrades the RNA strand of a DNA–RNA hybrid, can rescue the telomeric replication defect in FEN1-deleted cells [171].
DNA2, a multifunctional 5′–3′ DNA helicase with exonuclease and endonuclease activities, participates in Okazaki fragment maturation and processing of G4 DNA [173]. DNA2 heterozygous knockout in mice causes fragile telomere phenotype and telomere loss without genome-wide effects on replication [174], although the mechanisms for DNA2 function at telomeres remain to be determined.

3.6. UPF1 and Chromatin Remodelers

The up-frameshift suppressor 1 (UPF1, also known as RENT1 or SMG2) is a DNA/RNA-dependent ATPase and 5′–3′ helicase, known as a component of the RNA quality control machinery [175,176]. UPF1 ATPase activity is required to prevent telomere dysfunctions during replication [177]. UPF1 binds to telomeres through interaction with the shelterin component TPP1 [84,178]. UPF1 knockdown results in DNA damage at telomeres and frequent loss of the telomeres that are replicated by leading-strand synthesis [178]. UPF1 knockdown also results in an increased level of TERRA signal at telomeres, suggesting a role for UPF1 in displacement of TERRA [84,178]. If it is not displaced, TERRA can form a telomeric R-loop by binding to the C-rich DNA strand, and might induce replication stress and double-strand breaks during leading-strand DNA replication. Furthermore, the chromatin remodeling protein ATRX has been implicated in the displacement of TERRA to resolve recombinogenic DNA–RNA hybrid structures [179,180]. Loss of ATRX is associated with ALT in cancer cells, in which RNase H1 regulates TERRA–telomeric-DNA hybrids and telomere maintenance [179,181,182,183,184]. Deletion of mouse INO80, encoding a chromatin remodeler involved in diverse aspects of DNA metabolism [185], also results in fragile telomere phenotype [186].

3.7. Apollo

Apollo (also known as SNM1B) is a member of the metallo-β-lactamase/β-CASP family, and has 5′–3′ exonuclease activity [187]. Apollo has been implicated in several DNA damage responses including ATM activation and Fanconi anemia pathway [188,189,190,191,192]. Besides these genome-wide functions, Apollo has telomere-associated functions. Evidence indicates that Apollo localizes to telomeres through its interaction with TRF2 [67,193,194,195,196]. Studies of crystal structures showed that the C-terminal YxLxP motif of Apollo is involved in binding to TRF2, which requires the F120 residue of the TRF2 TRFH domain [67]. Knockdown of DCLRE1B, which encodes human Apollo protein, results in fragile telomere phenotype in telomeres produced by both lagging- or leading-strand replication [195]. Expressions of Apollo mutant proteins lacking the TRF2-binding or nuclease activity have dominant-negative effects on telomeric DNA replication [197,198]. Apollo has been suggested to act in the same pathway as DNA topoisomerase 2α (TOP2A), which relieves accumulating topological stress during human telomere replication [197]. However, exactly how the exonuclease activity of Apollo contributes to telomere replication is unknown. On the other hand, mouse TOP2A is recruited to telomeres in a manner that is dependent on TRF1, and which prevents the fragile telomere phenotype [112].
Mouse Apollo has a further essential role in the generation of the telomeric G-overhang after the bulk replication of telomeres [193,199,200]. Because leading-strand replication on the C-rich strand generates a blunt-ended daughter telomere, 5′-end resection of the C-rich strand by Apollo is required to form the single-stranded G-overhang. DCLRE1B-knockout mouse embryonic fibroblasts exhibit TIFs, especially in S phase, and have leading-end telomere fusion [193,199,200]. Such role of human Apollo remains to be clarified. Additional aspects of G-overhang generation, such as repression of Apollo by POT1 and following resection by exonuclease 1 (EXO1), are reviewed elsewhere [2,4,14].

3.8. POT1 and RAP1

POT1 is a shelterin component with multiple functions, and it binds directly to telomeric ssDNA. A well-documented role of POT1 is to protect the G-overhang from DNA repair activities by excluding RPA and ATR activation from the 3’ overhang ssDNA [201,202,203,204,205,206]. Other functions of POT1 in the regulation of G-overhang generation and telomere length (by controlling telomerase activity) are reviewed in detail elsewhere [103,207,208]. Moreover, POT1 has been proposed to overcome RPA accumulation on ssDNA during DNA replication and to repress sister telomere association [56,134]. TRF1 acts as a platform to recruit POT1 through the interaction mediated by TIN2–TPP1 in shelterin [56,205]. Mutations encoding POT1 variants defective in ssDNA binding have been found in patients with cancer, and expression of these variants elicits fragile telomere and ATR-dependent TIFs, which are signs of telomeric replication defects [209,210,211,212]. The function of POT1 in efficient replication seems to be mediated by the interaction with the CST (CTC1–STN1–TEN1) ternary complex [212], which stimulates replication fork restart. Knockdown of CTC1 or STN1 induces fragile telomere and TIFs and is epistatic to POT1 mutations [212]. Another function of POT1 is the unwinding of G4 DNA on the G-rich template strands [121,122,123,124]. In parallel, mouse Rap1, a shelterin component that interacts with TRF2, is required to prevent telomere fragility, telomere recombination, and telomere shortening, whereas human RAP1 inhibits chromosome fusions at telomeres [213,214,215].

3.9. The CST Complex

The CST complex is a ssDNA-binding complex, which is structurally related to RPA, and which is involved in the regulation of telomeric G-overhangs [216,217,218]. Besides high-affinity binding to telomeric ssDNA, interaction with TPP1–POT1 heterodimer regulates the telomeric localization of the CST complex [200,219,220]. CST stimulates RNA priming and DNA synthesis by the primase-Polα complex to fill in the C-strand after G-strand extension by telomerase and/or EXO1-mediated resection [200,220,221,222,223,224,225,226]. Because replication forks stall naturally at mammalian telomeres, an ATR-dependent fork restart mechanism is needed to complete DNA replication [227,228,229]. Knockdown of expression of CST components decreases bromodeoxyuridine incorporation at telomeres after release from hydroxyurea-induced replication fork arrest, and elicits telomere fragility [212,226,230,231,232,233,234,235,236]. Several lines of evidence suggest that CST contributes to fork restart not only in telomeres but also genome-wide under conditions of replication stress [232,233,235,237]. Stimulation of the primase-Polα complex has been implicated in the restart of stalled fork [226,231,236], but DNA-fiber analysis has also suggested that CST promotes replication recovery partly by activating dormant origins [232,235,237]. Results of deep-sequencing analysis revealed that CST recruits RAD51, a recombination protein, to GC-rich repetitive regions including telomeres in response to hydroxyurea [238]. The DNA repair protein BRCA2 also contributes to telomere replication as a RAD51 loader [239]. Recruited RAD51 would facilitate fork restart by strand exchange of the collapsed fork.

4. Restart of Replication to Complete Telomere Duplication

Prolonged replication fork arrest ultimately leads to irreversible fork collapse (Figure 3) [240]. In general, broken forks are rescued by incoming replication forks or repaired by recombination-mediated fork-restart mechanisms [241,242]. If no dormant origin exists in the telomeric region distal to the broken fork, the region may remain unreplicated. Therefore, loading of backup MCM replicative helicase in the telomere may be particularly important for the completion of telomere duplication. Homologous recombination-mediated processes, such as break-induced replication, provide alternative pathways to rescue the collapsed replication fork [243,244]. Recently, break-induced replication by PCNA–Polδ was shown to occur at mammalian telomeres [245,246]. MCM helicase may be dispensable for the break-induced telomere synthesis, and the DNA-unwinding mechanism in this process is unknown [246]. It has been suggested that break-induced replication promotes ALT to maintain the telomere length in telomerase-negative cancer cells [245,246]. The rescue of fork collapse by firing of dormant origins may contribute to prevent such aberrant telomere extension.

5. Concluding Remarks

Efficient replication of telomeric DNA requires a number of interactions between telomere-specific proteins and non-telomere-specific proteins to support fork progression (Figure 2). In the absence of these factors, replication forks frequently stall, collapse, and give rise to aberrant recombination, leading to telomere fragility. Unreplicated regions of telomeres or improper recombination such as sister telomere association may cause aberrant chromosome segregation in mitosis (Figure 3). Because the factors that overcome the impediments to telomeric replication are also involved in general DNA replication, repair, and recombination, and are sometimes essential for viability, separation-of-function mutants have been valuable tools to elucidate the mechanisms for the preservation of telomere integrity. Telomeres regulate cellular lifespan and their dysfunction is a driver of genomic instability. Besides the simple telomere protection, efficient replication of telomeres has emerged as another factor that influences aging and carcinogenesis [55,150,212].
It is now clear that DNA replication at telomeres is supported by multiple mechanisms, which are discussed in this review and another recent review [247]. However, much remains unknown about how these mechanisms are controlled during the cell cycle, differentiation, aging, and cancer development. In particular, several factors appear to have opposing effects on telomeric DNA replication. For example, G4 DNA may contribute to specification of replication origins, but impairs replication fork progression. Activation of the ATR-dependent checkpoint pathway is repressed by POT1 at telomeres, but ATR is required to prevent telomere fragility. In addition, although R-loops formed by TERRA transcripts are an obstacle to the replication machinery, TERRA is suggested to promote the switch from RPA to POT1 at the G-overhang after replication [10,11]. How are the apparently conflicting roles of these factors coordinated? One major future challenge is to understand how telomeres manage to complete their duplication while avoiding the potential harmful effects of this process, including replication stress, telomere shortening, and genomic instability. Whether the replication machinery itself modulates a complex network of telomeric protein–protein interactions in response to fork stalling is an important question. Indeed, Timeless, a component of the fork protection complex that travels with the replication fork, is required for efficient telomere replication, and interacts with TRF1 and TRF2 [248]. Posttranslational modifications of telomeric factors and nuclear localization of telomeres might determine the appropriate use of multiple factors at telomeres.
Further elucidation of the molecular mechanisms that ensure efficient telomere replication is an important issue in telomere biology. The molecular mechanisms that coordinate dormant origin firing, homology-directed repair, and break-induced replication in response to fork collapse at telomeres are largely unknown. Another question is whether telomere length has an impact on telomere fragility. It is well known that short telomeres cause telomere deprotection and cell death. On the other hand, longer telomeres might increase the probability of fork stalling and collapse, leading to telomere loss. Comparing the frequency of telomeric fork stalling, collapse, or restart in broad biological contexts (e.g., normal vs. cancer cells, young vs. old cells) could provide insights into the endogenous sources of telomere fragility. Furthermore, it is important to uncover the molecular mechanisms underlying abnormal telomere shortening and cancer predisposition in short telomere diseases (so-called telomeropathy), such as Werner syndrome and Hoyeraal–Hreidarsson syndrome [150,198], which are caused by mutations in genes encoding factors involved in telomere replication. It will be critical to understand how semi-conservative replication influences telomere elongation by telomerase and vice versa. A comprehensive and integrated understanding of these processes could yield novel targets and strategies for disease diagnosis and therapy.

Acknowledgments

We apologize to those whose work we were unable to cite due to space limitations. We thank members of the Fujita lab for helpful discussion and comments on the manuscript. This work was supported in part by Grants to Masatoshi Fujita and Kazumasa Yoshida from the Ministry of Education, Science, Sports, Technology and Culture of Japan.

Author Contributions

Mitsunori Higa, Masatoshi Fujita, and Kazumasa Yoshida wrote the paper and produced the figures.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Palm, W.; de Lange, T. How Shelterin Protects Mammalian Telomeres. Annu. Rev. Genet. 2008, 42, 301–334. [Google Scholar] [CrossRef] [PubMed]
  2. Doksani, Y.; de Lange, T. The Role of Double-Strand Break Repair Pathways at Functional and Dysfunctional Telomeres. Cold Spring Harb. Perspect. Biol. 2014, 6, a016576. [Google Scholar] [CrossRef] [PubMed]
  3. Arnoult, N.; Karlseder, J. Complex interactions between the DNA-damage response and mammalian telomeres. Nat. Struct. Mol. Biol. 2015, 22, 859–866. [Google Scholar] [CrossRef] [PubMed]
  4. Martínez, P.; Blasco, M.A. Replicating through telomeres: A means to an end. Trends Biochem. Sci. 2015, 40, 504–515. [Google Scholar] [CrossRef] [PubMed]
  5. Davis, T.; Kipling, D. Telomeres and telomerase biology in vertebrates: Progress towards a non-human model for replicative senescence and ageing. Biogerontology 2005, 6, 371–385. [Google Scholar] [CrossRef] [PubMed]
  6. Lewis, K.A.; Wuttke, D.S. Telomerase and telomere-associated proteins: Structural insights into mechanism and evolution. Structure 2012, 20, 28–39. [Google Scholar] [CrossRef] [PubMed]
  7. Doksani, Y.; Wu, J.Y.; de Lange, T.; Zhuang, X. Super-resolution fluorescence imaging of telomeres reveals TRF2-dependent T-loop formation. Cell 2013, 155, 345–356. [Google Scholar] [CrossRef] [PubMed]
  8. Griffith, J.D.; Comeau, L.; Rosenfield, S.; Stansel, R.M.; Bianchi, A.; Moss, H.; de Lange, T. Mammalian telomeres end in a large duplex loop. Cell 1999, 97, 503–514. [Google Scholar] [CrossRef]
  9. Rhodes, D.; Lipps, H.J. G-quadruplexes and their regulatory roles in biology. Nucleic Acids Res. 2015, 43, 8627–8637. [Google Scholar] [CrossRef] [PubMed]
  10. Azzalin, C.M.; Lingner, J. Telomere functions grounding on TERRA firma. Trends Cell Biol. 2015, 25, 29–36. [Google Scholar] [CrossRef] [PubMed]
  11. Cusanelli, E.; Chartrand, P. Telomeric repeat-containing RNA TERRA: A noncoding RNA connecting telomere biology to genome integrity. Front. Genet. 2015. [Google Scholar] [CrossRef] [PubMed]
  12. Schoeftner, S.; Blasco, M.A. Chromatin regulation and non-coding RNAs at mammalian telomeres. Semin. Cell Dev. Biol. 2010, 21, 186–193. [Google Scholar] [CrossRef] [PubMed]
  13. Blasco, M.A. The epigenetic regulation of mammalian telomeres. Nat. Rev. Genet. 2007, 8, 299–309. [Google Scholar] [CrossRef] [PubMed]
  14. Pfeiffer, V.; Lingner, J. Replication of telomeres and the regulation of telomerase. Cold Spring Harb. Perspect. Biol. 2013, 5, a010405. [Google Scholar] [CrossRef] [PubMed]
  15. Magdalou, I.; Lopez, B.S.; Pasero, P.; Lambert, S.A.E. The causes of replication stress and their consequences on genome stability and cell fate. Semin. Cell Dev. Biol. 2014, 30, 154–164. [Google Scholar] [CrossRef] [PubMed]
  16. Fragkos, M.; Ganier, O.; Coulombe, P.; Méchali, M. DNA replication origin activation in space and time. Nat. Rev. Mol. Cell Biol. 2015, 16, 360–374. [Google Scholar] [CrossRef] [PubMed]
  17. Siddiqui, K.; On, K.F.; Diffley, J.F.X. Regulating DNA replication in Eukarya. Cold Spring Harb. Perspect. Biol. 2013, 5, a012930. [Google Scholar] [CrossRef] [PubMed]
  18. Deegan, T.D.; Diffley, J.F.X. MCM: One ring to rule them all. Curr. Opin. Struct. Biol. 2016, 37, 145–151. [Google Scholar] [CrossRef] [PubMed]
  19. Yardimci, H.; Walter, J.C. Prereplication-complex formation: A molecular double take? Nat. Struct. Mol. Biol. 2014, 21, 20–25. [Google Scholar] [CrossRef] [PubMed]
  20. Tanaka, S.; Araki, H. Helicase activation and establishment of replication forks at chromosomal origins of replication. Cold Spring Harb. Perspect. Biol. 2013, 5, a010371. [Google Scholar] [CrossRef] [PubMed]
  21. Zegerman, P. Evolutionary conservation of the CDK targets in eukaryotic DNA replication initiation. Chromosoma 2015, 124, 309–321. [Google Scholar] [CrossRef] [PubMed]
  22. Lujan, S.A.; Williams, J.S.; Kunkel, T.A. DNA Polymerases Divide the Labor of Genome Replication. Trends Cell Biol. 2016, 26, 640–654. [Google Scholar] [CrossRef] [PubMed]
  23. Hills, S.A.; Diffley, J.F.X. DNA replication and oncogene-induced replicative stress. Curr. Biol. 2014, 24, R435–R444. [Google Scholar] [CrossRef] [PubMed]
  24. Miotto, B.; Ji, Z.; Struhl, K. Selectivity of ORC binding sites and the relation to replication timing, fragile sites, and deletions in cancers. Proc. Natl. Acad. Sci. USA 2016, 113, E4810–E4819. [Google Scholar] [CrossRef] [PubMed]
  25. Dellino, G.I.; Cittaro, D.; Piccioni, R.; Luzi, L.; Banfi, S.; Segalla, S.; Cesaroni, M.; Mendoza-Maldonado, R.; Giacca, M.; Pelicci, P.G. Genome-wide mapping of human DNA-replication origins: Levels of transcription at ORC1 sites regulate origin selection and replication timing. Genome Res. 2013, 23, 1–11. [Google Scholar] [CrossRef] [PubMed]
  26. Petryk, N.; Kahli, M.; D’Aubenton-Carafa, Y.; Jaszczyszyn, Y.; Shen, Y.; Maud, S.; Thermes, C.; Chen, C.L.; Hyrien, O. Replication landscape of the human genome. Nat. Commun. 2016. [Google Scholar] [CrossRef] [PubMed]
  27. Powell, S.K.; MacAlpine, H.K.; Prinz, J.A.; Li, Y.; Belsky, J.A.; MacAlpine, D.M. Dynamic loading and redistribution of the Mcm2-7 helicase complex through the cell cycle. EMBO J. 2015, 34, 531–543. [Google Scholar] [CrossRef] [PubMed]
  28. Hyrien, O. How MCM loading and spreading specify eukaryotic DNA replication initiation sites. F1000Research 2016. [Google Scholar] [CrossRef] [PubMed]
  29. Sugimoto, N.; Maehara, K.; Yoshida, K.; Yasukouchi, S.; Osano, S.; Watanabe, S.; Aizawa, M.; Yugawa, T.; Kiyono, T.; Kurumizaka, H.; et al. Cdt1-binding protein GRWD1 is a novel histone-binding protein that facilitates MCM loading through its influence on chromatin architecture. Nucleic Acids Res. 2015, 43, 5898–5911. [Google Scholar] [CrossRef] [PubMed]
  30. Renard-Guillet, C.; Kanoh, Y.; Shirahige, K.; Masai, H. Temporal and spatial regulation of eukaryotic DNA replication: From regulated initiation to genome-scale timing program. Semin. Cell Dev. Biol. 2014, 30, 110–120. [Google Scholar] [CrossRef] [PubMed]
  31. Hyrien, O. Peaks cloaked in the mist: The landscape of mammalian replication origins. J. Cell Biol. 2015, 208, 147–160. [Google Scholar] [CrossRef] [PubMed]
  32. Zeman, M.K.; Cimprich, K.A. Causes and consequences of replication stress. Nat. Cell Biol. 2014, 16, 2–9. [Google Scholar] [CrossRef] [PubMed]
  33. Alver, R.C.; Chadha, G.S.; Blow, J.J. The contribution of dormant origins to genome stability: From cell biology to human genetics. DNA Repair 2014, 19, 182–189. [Google Scholar] [CrossRef] [PubMed]
  34. Ge, X.Q.; Jackson, D.A.; Blow, J.J. Dormant origins licensed by excess Mcm2-7 are required for human cells to survive replicative stress. Genes Dev. 2007, 21, 3331–3341. [Google Scholar] [CrossRef] [PubMed]
  35. Ibarra, A.; Schwob, E.; Méndez, J. Excess MCM proteins protect human cells from replicative stress by licensing backup origins of replication. Proc. Natl. Acad. Sci. USA 2008, 105, 8956–8961. [Google Scholar] [CrossRef] [PubMed]
  36. Kawabata, T.; Luebben, S.W.; Yamaguchi, S.; Ilves, I.; Matise, I.; Buske, T.; Botchan, M.R.; Shima, N. Stalled Fork Rescue via Dormant Replication Origins in Unchallenged S Phase Promotes Proper Chromosome Segregation and Tumor Suppression. Mol. Cell 2011, 41, 543–553. [Google Scholar] [CrossRef] [PubMed]
  37. Yekezare, M.; Gomez-Gonzalez, B.; Diffley, J.F.X. Controlling DNA replication origins in response to DNA damage—Inhibit globally, activate locally. J. Cell Sci. 2013, 126, 1297–1306. [Google Scholar] [CrossRef] [PubMed]
  38. Jones, R.M.; Petermann, E. Replication fork dynamics and the DNA damage response. Biochem. J. 2012, 443, 13–26. [Google Scholar] [CrossRef] [PubMed]
  39. Ten Hagen, K.G.; Gilbert, D.M.; Willard, H.F.; Cohen, S.N. Replication Timing of DNA Sequences Associated with Human Centromeres and Telomeres. Mol. Cell. Biol. 1990, 10, 6348–6355. [Google Scholar] [CrossRef] [PubMed]
  40. Wright, W.E.; Tesmer, V.M.; Liao, M.L.; Shay, J.W. Normal human telomeres are not late replicating. Exp. Cell Res. 1999, 251, 492–499. [Google Scholar] [CrossRef] [PubMed]
  41. Arnoult, N.; Schluth-Bolard, C.; Letessier, A.; Drascovic, I.; Bouarich-Bourimi, R.; Campisi, J.; Kim, S.H.; Boussouar, A.; Ottaviani, A.; Magdinier, F.; et al. Replication timing of human telomeres is chromosome arm-specific, influenced by subtelomeric structures and connected to nuclear localization. PLoS Genet. 2010, 6, e1000920. [Google Scholar] [CrossRef] [PubMed]
  42. Piqueret-stephan, L.; Ricoul, M.; Hempel, W.M.; Sabatier, L. Replication Timing of Human Telomeres is Conserved during Immortalization and Influenced by Respective Subtelomeres. Sci. Rep. 2016. [Google Scholar] [CrossRef] [PubMed]
  43. Hultdin, M.; Gronlund, E.; Norrback, K.F.; Just, T.; Taneja, K.; Roos, G. Replication Timing of Human Telomeric DNA and Other Repetitive Sequences Analyzed by Fluorescence in Situ Hybridization and Flow Cytometry. Exp. Cell Res. 2001, 271, 223–229. [Google Scholar] [CrossRef] [PubMed]
  44. Ofir, R.; Wong, A.C.; McDermid, H.E.; Skorecki, K.L.; Selig, S. Position effect of human telomeric repeats on replication timing. Proc. Natl. Acad. Sci. USA 1999, 96, 11434–11439. [Google Scholar] [CrossRef] [PubMed]
  45. Yamazaki, S.; Hayano, M.; Masai, H. Replication timing regulation of eukaryotic replicons: Rif1 as a global regulator of replication timing. Trends Genet. 2013, 29, 449–460. [Google Scholar] [CrossRef] [PubMed]
  46. Drosopoulos, W.C.; Kosiyatrakul, S.T.; Schildkraut, C.L. BLM helicase facilitates telomere replication during leading strand synthesis of telomeres. J. Cell Biol. 2015, 210, 191–208. [Google Scholar] [CrossRef] [PubMed]
  47. Drosopoulos, W.C.; Kosiyatrakul, S.T.; Yan, Z.; Calderano, S.G.; Schildkraut, C.L. Human telomeres replicate using chromosomespecific, rather than universal, replication programs. J. Cell Biol. 2012, 197, 253–266. [Google Scholar] [CrossRef] [PubMed]
  48. Sfeir, A.; Kosiyatrakul, S.T.; Hockemeyer, D.; MacRae, S.L.; Karlseder, J.; Schildkraut, C.L.; De Lange, T. Mammalian Telomeres Resemble Fragile Sites and Require TRF1 for Efficient Replication. Cell 2009, 138, 90–103. [Google Scholar] [CrossRef] [PubMed]
  49. Newman, T.J.; Mamun, M.A.; Nieduszynski, C.A.; Blow, J.J. Replisome stall events have shaped the distribution of replication origins in the genomes of yeasts. Nucleic Acids Res. 2013, 41, 9705–9718. [Google Scholar] [CrossRef] [PubMed]
  50. Hayashi, M.; Katou, Y.; Itoh, T.; Tazumi, M.; Yamada, Y.; Takahashi, T.; Nakagawa, T.; Shirahige, K.; Masukata, H. Genome-wide localization of pre-RC sites and identification of replication origins in fission yeast. EMBO J. 2007, 26, 1327–1339. [Google Scholar] [CrossRef] [PubMed]
  51. Heichinger, C.; Penkett, C.J.; Bähler, J.; Nurse, P. Genome-wide characterization of fission yeast DNA replication origins. EMBO J. 2006, 25, 5171–5179. [Google Scholar] [CrossRef] [PubMed]
  52. Foulk, M.S.; Urban, J.M.; Casella, C.; Gerbi, S.A. Characterizing and controlling intrinsic biases of lambda exonuclease in nascent strand sequencing reveals phasing between nucleosomes and G-quadruplex motifs around a subset of human replication origins. Genome Res. 2015, 25, 725–735. [Google Scholar] [CrossRef] [PubMed]
  53. Ozeri-Galai, E.; Lebofsky, R.; Rahat, A.; Bester, A.C.; Bensimon, A.; Kerem, B. Failure of Origin Activation in Response to Fork Stalling Leads to Chromosomal Instability at Fragile Sites. Mol. Cell 2011, 43, 122–131. [Google Scholar] [CrossRef] [PubMed]
  54. Debatisse, M.; Le Tallec, B.; Letessier, A.; Dutrillaux, B.; Brison, O. Common fragile sites: Mechanisms of instability revisited. Trends Genet. 2012, 28, 22–32. [Google Scholar] [CrossRef] [PubMed]
  55. Martínez, P.; Thanasoula, M.; Muñoz, P.; Liao, C.; Tejera, A.; McNees, C.; Flores, J.M.; Fernández-Capetillo, O.; Tarsounas, M.; Blasco, M.A. Increased telomere fragility and fusions resulting from TRF1 deficiency lead to degenerative pathologies and increased cancer in mice. Genes Dev. 2009, 23, 2060–2075. [Google Scholar] [CrossRef] [PubMed]
  56. Zimmermann, M.; Kibe, T.; Kabir, S.; de Lange, T. TRF1 negotiates TTAGGG repeat associated-replication problems by recruiting the BLM helicase and the TPP1/POT1 repressor of ATR signaling. Genes Dev. 2014, 28, 2477–2491. [Google Scholar] [CrossRef] [PubMed]
  57. Vannier, J.B.; Pavicic-Kaltenbrunner, V.; Petalcorin, M.I.R.; Ding, H.; Boulton, S.J. RTEL1 dismantles T loops and counteracts telomeric G4-DNA to maintain telomere integrity. Cell 2012, 149, 795–806. [Google Scholar] [CrossRef] [PubMed]
  58. Barefield, C.; Karlseder, J. The BLM helicase contributes to telomere maintenance through processing of late-replicating intermediate structures. Nucleic Acids Res. 2012, 40, 7358–7367. [Google Scholar] [CrossRef] [PubMed]
  59. Deng, Z.; Norseen, J.; Wiedmer, A.; Riethman, H.; Lieberman, P.M. TERRA RNA Binding to TRF2 Facilitates Heterochromatin Formation and ORC Recruitment at Telomeres. Mol. Cell 2009, 35, 403–413. [Google Scholar] [CrossRef] [PubMed]
  60. Deng, Z.; Dheekollu, J.; Broccoli, D.; Dutta, A.; Lieberman, P.M. The Origin Recognition Complex Localizes to Telomere Repeats and Prevents Telomere-Circle Formation. Curr. Biol. 2007, 17, 1989–1995. [Google Scholar] [CrossRef] [PubMed]
  61. Tatsumi, Y.; Ezura, K.; Yoshida, K.; Yugawa, T.; Narisawa-Saito, M.; Kiyono, T.; Ohta, S.; Obuse, C.; Fujita, M. Involvement of human ORC and TRF2 in pre-replication complex assembly at telomeres. Genes Cells 2008, 13, 1045–1059. [Google Scholar] [CrossRef] [PubMed]
  62. Déjardin, J.; Kingston, R.E. Purification of Proteins Associated with Specific Genomic Loci. Cell 2009, 136, 175–186. [Google Scholar] [CrossRef] [PubMed]
  63. Bartocci, C.; Diedrich, J.K.; Ouzounov, I.; Li, J.; Piunti, A.; Pasini, D.; Yates, J.R.; Lazzerini Denchi, E. Isolation of chromatin from dysfunctional telomeres reveals an important role for Ring1b in NHEJ-mediated chromosome fusions. Cell Rep. 2014, 7, 1320–1332. [Google Scholar] [CrossRef] [PubMed]
  64. Grolimund, L.; Aeby, E.; Hamelin, R.; Armand, F.; Chiappe, D.; Moniatte, M.; Lingner, J. A quantitative telomeric chromatin isolation protocol identifies different telomeric states. Nat. Commun. 2013. [Google Scholar] [CrossRef] [PubMed]
  65. Atanasiu, C.; Deng, Z.; Wiedmer, A.; Norseen, J.; Lieberman, P.M. ORC binding to TRF2 stimulates OriP replication. EMBO Rep. 2006, 7, 716–721. [Google Scholar] [CrossRef] [PubMed]
  66. Higa, M.; Kushiyama, T.; Kurashige, S.; Kohmon, D.; Enokitani, K.; Iwahori, S.; Sugimoto, N.; Yoshida, K.; Fujita, M. TRF2 recruits ORC through TRFH domain dimerization. Biochim. Biophys. Acta—Mol. Cell Res. 2017, 1864, 191–201. [Google Scholar] [CrossRef] [PubMed]
  67. Chen, Y.; Yang, Y.; van Overbeek, M.; Donigian, J.R.; Baciu, P.; de Lange, T.; Lei, M. A shared docking motif in TRF1 and TRF2 used for differential recruitment of telomeric proteins. Science 2008, 319, 1092–1096. [Google Scholar] [CrossRef] [PubMed]
  68. Gaullier, G.; Miron, S.; Pisano, S.; Buisson, R.; Le Bihan, Y.V.; Tellier-Lebegue, C.; Messaoud, W.; Roblin, P.; Guimaras, B.G.; Thai, R.; et al. A higher-order entity formed by the flexible assembly of RAP1 with TRF2. Nucleic Acids Res. 2016, 44, 1962–1976. [Google Scholar] [CrossRef] [PubMed]
  69. Kim, H.; Lee, O.; Xin, H.; Chen, L.; Qin, J.; Chae, H.K.; Lin, S.; Safari, A.; Liu, D.; Songyang, Z. TRF2 functions as a protein hub and regulates telomere maintenance by recognizing specific peptide motifs. Nat. Struct. Mol. Biol. 2009, 16, 372–379. [Google Scholar] [CrossRef] [PubMed]
  70. Sarek, G.; Vannier, J.; Panier, S.; Petrini, J.H.J.; Boulton, S.J. TRF2 Recruits RTEL1 to Telomeres in S Phase to Promote T-Loop Unwinding. Mol. Cell 2015, 57, 622–635. [Google Scholar] [CrossRef] [PubMed]
  71. Wan, B.; Yin, J.; Horvath, K.; Sarkar, J.; Chen, Y.; Wu, J.; Wan, K.; Lu, J.; Gu, P.; Yu, E.Y.; et al. SLX4 Assembles a Telomere Maintenance Toolkit by Bridging Multiple Endonucleases with Telomeres. Cell Rep. 2013, 4, 861–869. [Google Scholar] [CrossRef] [PubMed]
  72. Wilson, J.S.J.; Tejera, A.M.; Castor, D.; Toth, R.; Blasco, M.A.; Rouse, J. Localization-Dependent and -Independent Roles of SLX4 in Regulating Telomeres. Cell Rep. 2013, 4, 853–860. [Google Scholar] [CrossRef] [PubMed]
  73. Patel, D.J.; Phan, A.T.; Kuryavyi, V. Human telomere, oncogenic promoter and 5’-UTR G-quadruplexes: Diverse higher order DNA and RNA targets for cancer therapeutics. Nucleic Acids Res. 2007, 35, 7429–7455. [Google Scholar] [CrossRef] [PubMed]
  74. Huppert, J.L. Structure, location and interactions of G-quadruplexes. FEBS J. 2010, 277, 3452–3458. [Google Scholar] [CrossRef] [PubMed]
  75. Valton, A.L.; Hassan-Zadeh, V.; Lema, I.; Boggetto, N.; Alberti, P.; Saintome, C.; Riou, J.F.; Prioleau, M.N. G4 motifs affect origin positioning and efficiency in two vertebrate replicators. EMBO J. 2014, 33, 732–746. [Google Scholar] [CrossRef] [PubMed]
  76. Valton, A.L.; Prioleau, M.N. G-Quadruplexes in DNA Replication: A Problem or a Necessity? Trends Genet. 2016, 32, 697–706. [Google Scholar] [CrossRef] [PubMed]
  77. Cayrou, C.; Coulombe, P.; Puy, A.; Rialle, S.; Kaplan, N.; Segal, E.; Méchali, M. New insights into replication origin characteristics in metazoans. Cell Cycle 2012, 11, 658–667. [Google Scholar] [CrossRef] [PubMed]
  78. Besnard, E.; Babled, A.; Lapasset, L.; Milhavet, O.; Parrinello, H.; Dantec, C.; Marin, J.M.; Lemaitre, J.M. Unraveling cell type–specific and reprogrammable human replication origin signatures associated with G-quadruplex consensus motifs. Nat. Struct. Mol. Biol. 2012, 19, 837–844. [Google Scholar] [CrossRef] [PubMed]
  79. Prioleau, M.; Macalpine, D.M. DNA replication origins—Where do we begin? Genes Dev. 2016, 30, 1683–1697. [Google Scholar] [CrossRef] [PubMed]
  80. Hoshina, S.; Yura, K.; Teranishi, H.; Kiyasu, N.; Tominaga, A.; Kadoma, H.; Nakatsuka, A.; Kunichika, T.; Obuse, C.; Waga, S. Human origin recognition complex binds preferentially to G-quadruplex-preferable RNA and single-stranded DNA. J. Biol. Chem. 2013, 288, 30161–30171. [Google Scholar] [CrossRef] [PubMed]
  81. Mendoza, O.; Bourdoncle, A.; Boule, J.B.; Brosh, R.M., Jr.; Mergny, J.L. G-quadruplexes and helicases. Nucleic Acids Res. 2016, 44, 1989–2006. [Google Scholar] [CrossRef] [PubMed]
  82. Neidle, S.; Parkinson, G.N. The structure of telomeric DNA. Curr. Opin. Struct. Biol. 2003, 13, 275–283. [Google Scholar] [CrossRef]
  83. Biffi, G.; Tannahill, D.; McCafferty, J.; Balasubramanian, S. Quantitative visualization of DNA G-quadruplex structures in human cells. Nat. Chem. 2013, 5, 182–186. [Google Scholar] [CrossRef] [PubMed]
  84. Azzalin, C.M.; Reichenbach, P.; Khoriauli, L.; Giulotto, E.; Lingner, J. Telomeric repeat containing RNA and RNA surveillance factors at mammalian chromosome ends. Science 2007, 318, 798–801. [Google Scholar] [CrossRef] [PubMed]
  85. Schoeftner, S.; Blasco, M. Developmentally regulated transcription of mammalian telomeres by DNA-dependent RNA polymerase II. Nat. Cell Biol. 2008, 10, 228–236. [Google Scholar] [CrossRef] [PubMed]
  86. Montero, J.J.; López de Silanes, I.; Graña, O.; Blasco, M.A. Telomeric RNAs are essential to maintain telomeres. Nat. Commun. 2016. [Google Scholar] [CrossRef] [PubMed]
  87. Rippe, K.; Luke, B. TERRA and the state of the telomere. Nat. Struct. Mol. Biol. 2015, 22, 853–858. [Google Scholar] [CrossRef] [PubMed]
  88. Balk, B.; Dees, M.; Bender, K.; Luke, B. The differential processing of telomeres in response to increased telomeric transcription and RNA-DNA hybrid accumulation. RNA Biol. 2014, 11, 95–100. [Google Scholar] [CrossRef] [PubMed]
  89. Chakraborty, A.; Shen, Z.; Prasanth, S.G. “ORCanization” on heterochromatin linking DNA replication initiation to chromatin organization. Epigenetics 2011, 6, 665–670. [Google Scholar] [CrossRef] [PubMed]
  90. Shen, Z.; Sathyan, K.M.; Geng, Y.; Zheng, R.; Chakraborty, A.; Freeman, B.; Wang, F.; Prasanth, K.V.; Prasanth, S.G. A WD-repeat protein stabilizes ORC binding to chromatin. Mol. Cell 2010, 40, 99–111. [Google Scholar] [CrossRef] [PubMed]
  91. Shen, Z.; Chakraborty, A.; Jain, A.; Giri, S.; Ha, T.; Prasanth, K.V.; Prasanth, S.G. Dynamic Association of ORCA with Prereplicative Complex Components Regulates DNA Replication Initiation. Mol. Cell. Biol. 2012, 32, 3107–3120. [Google Scholar] [CrossRef] [PubMed]
  92. Beck, D.B.; Burton, A.; Oda, H.; Ziegler-Birling, C.; Torres-Padilla, M.E.; Reinberg, D. The role of PR-Set7 in replication licensing depends on Suv4-20h. Genes Dev. 2012, 26, 2580–2589. [Google Scholar] [CrossRef] [PubMed]
  93. Vermeulen, M.; Eberl, H.C.; Matarese, F.; Marks, H.; Denissov, S.; Butter, F.; Lee, K.K.; Olsen, J.V.; Hyman, A.A.; Stunnenberg, H.G.; Mann, M. Quantitative Interaction Proteomics and Genome-wide Profiling of Epigenetic Histone Marks and Their Readers. Cell 2010, 142, 967–980. [Google Scholar] [CrossRef] [PubMed]
  94. Tardat, M.; Brustel, J.; Kirsh, O.; Lefevbre, C.; Callanan, M.; Sardet, C.; Julien, E. The histone H4 Lys 20 methyltransferase PR-Set7 regulates replication origins in mammalian cells. Nat. Cell Biol. 2010, 12, 1086–1093. [Google Scholar] [CrossRef] [PubMed]
  95. Tardat, M.; Murr, R.; Herceg, Z.; Sardet, C.; Julien, E. PR-Set7-dependent lysine methylation ensures genome replication and stability through S phase. J. Cell Biol. 2007, 179, 1413–1426. [Google Scholar] [CrossRef] [PubMed]
  96. Oda, H.; Okamoto, I.; Murphy, N.; Chu, J.; Price, S.M.; Shen, M.M.; Torres-Padilla, M.E.; Heard, E.; Reinberg, D. Monomethylation of histone H4-lysine 20 is involved in chromosome structure and stability and is essential for mouse development. Mol. Cell. Biol. 2009, 29, 2278–2295. [Google Scholar] [CrossRef] [PubMed]
  97. Schotta, G.; Sengupta, R.; Kubicek, S.; Malin, S.; Kauer, M.; Callén, E.; Celeste, A.; Pagani, M.; Opravil, S.; De La Rosa-Velazquez, I.A.; et al. A chromatin-wide transition to H4K20 monomethylation impairs genome integrity and programmed DNA rearrangements in the mouse. Genes Dev. 2008, 22, 2048–2061. [Google Scholar] [CrossRef] [PubMed]
  98. Jørgensen, S.; Schotta, G.; Sørensen, C.S. Histone H4 Lysine 20 methylation: Key player in epigenetic regulation of genomic integrity. Nucleic Acids Res. 2013, 41, 2797–2806. [Google Scholar] [CrossRef] [PubMed]
  99. Kuo, A.J.; Song, J.; Cheung, P.; Ishibe-Murakami, S.; Yamazoe, S.; Chen, J.K.; Patel, D.J.; Gozani, O. The BAH domain of ORC1 links H4K20me2 to DNA replication licensing and Meier-Gorlin syndrome. Nature 2012, 484, 115–119. [Google Scholar] [CrossRef] [PubMed]
  100. Schotta, G.; Lachner, M.; Sarma, K.; Ebert, A.; Sengupta, R.; Reuter, G.; Reinberg, D.; Jenuwein, T. A silencing pathway to induce H3-K9 and H4-K20 trimethylation at constitutive heterochromatin. Genes Dev. 2004, 18, 1251–1262. [Google Scholar] [CrossRef] [PubMed]
  101. Gonzalo, S.; García-Cao, M.; Fraga, M.F.; Schotta, G.; Peters, A.H.F.M.; Cotter, S.E.; Eguía, R.; Dean, D.C.; Esteller, M.; Jenuwein, T.; et al. Role of the RB1 family in stabilizing histone methylation at constitutive heterochromatin. Nat. Cell Biol. 2005, 7, 420–428. [Google Scholar] [CrossRef] [PubMed]
  102. Regha, K.; Sloane, M.A.; Huang, R.; Pauler, F.M.; Warczok, K.E.; Melikant, B.; Radolf, M.; Martens, J.H.A.; Schotta, G.; Jenuwein, T.; et al. Active and Repressive Chromatin Are Interspersed without Spreading in an Imprinted Gene Cluster in the Mammalian Genome. Mol. Cell 2007, 27, 353–366. [Google Scholar] [CrossRef] [PubMed]
  103. Nandakumar, J.; Cech, T.R. Finding the end: Recruitment of telomerase to telomeres. Nat. Rev. Mol. Cell Biol. 2013, 14, 69–82. [Google Scholar] [CrossRef] [PubMed]
  104. Ohki, R.; Ishikawa, F. Telomere-bound TRF1 and TRF2 stall the replication fork at telomeric repeats. Nucleic Acids Res. 2004, 32, 1627–1637. [Google Scholar] [CrossRef] [PubMed]
  105. Bosco, N.; De Lange, T. A TRF1-controlled common fragile site containing interstitial telomeric sequences. Chromosoma 2012, 121, 465–474. [Google Scholar] [CrossRef] [PubMed]
  106. Edwards, D.N.; Machwe, A.; Wang, Z.; Orren, D.K. Intramolecular telomeric G-quadruplexes dramatically inhibit DNA synthesis by replicative and translesion polymerases, revealing their potential to lead to genetic change. PLoS ONE 2014, 9, e80664. [Google Scholar] [CrossRef] [PubMed]
  107. Phan, A.T. Human telomeric G-quadruplex: Structures of DNA and RNA sequences. FEBS J. 2010, 277, 1107–1117. [Google Scholar] [CrossRef] [PubMed]
  108. Lormand, J.D.; Buncher, N.; Murphy, C.T.; Kaur, P.; Lee, M.Y.; Burgers, P.; Wang, H.; Kunkel, T.A.; Opresko, P.L. DNA polymerase δ stalls on telomeric lagging strand templates independently from G-quadruplex formation. Nucleic Acids Res. 2013, 41, 10323–10333. [Google Scholar] [CrossRef] [PubMed]
  109. Nera, B.; Huang, H.S.; Lai, T.; Xu, L. Elevated levels of TRF2 induce telomeric ultrafine anaphase bridges and rapid telomere deletions. Nat. Commun. 2015. [Google Scholar] [CrossRef] [PubMed]
  110. Okamoto, K.; Iwano, T.; Tachibana, M.; Shinkai, Y. Distinct roles of TRF1 in the regulation of telomere structure and lengthening. J. Biol. Chem. 2008, 283, 23981–23988. [Google Scholar] [CrossRef] [PubMed]
  111. Martínez, P.; Flores, J.M.; Blasco, M.A. 53BP1 deficiency combined with telomere dysfunction activates ATR-dependent DNA damage response. J. Cell Biol. 2012, 197, 283–300. [Google Scholar] [CrossRef] [PubMed]
  112. d’Alcontres, M.S.; Palacios, J.A.; Mejias, D.; Blasco, M.A. TopoIIα prevents telomere fragility and formation of ultra thin DNA bridges during mitosis through TRF1-dependent binding to telomeres. Cell Cycle 2014, 13, 1463–1481. [Google Scholar] [CrossRef] [PubMed]
  113. Croteau, D.L.; Popuri, V.; Opresko, P.L.; Bohr, V.A. Human RecQ Helicases in DNA Repair, Recombination, and Replication. Annu. Rev. Biochem. 2014, 83, 519–552. [Google Scholar] [CrossRef] [PubMed]
  114. Mohaghegh, P.; Karow, J.K.; Brosh, R.M.; Bohr, V.A.; Hickson, I.D. The Bloom’s and Werner’s syndrome proteins are DNA structure-specific helicases. Nucleic Acids Res. 2001, 29, 2843–2849. [Google Scholar] [CrossRef] [PubMed]
  115. Sun, H.; Karow, J.K.; Hickson, I.D.; Maizels, N. The Bloom’s syndrome helicase unwinds G4 DNA. J. Biol. Chem. 1998, 273, 27587–27592. [Google Scholar] [CrossRef] [PubMed]
  116. Machwe, A.; Karale, R.; Xu, X.; Liu, Y.; Orren, D.K. The Werner and Bloom syndrome proteins help resolve replication blockage by converting (regressed) Holliday junctions to functional replication forks. Biochemistry 2011, 50, 6774–6788. [Google Scholar] [CrossRef] [PubMed]
  117. Chatterjee, S.; Zagelbaum, J.; Savitsky, P.; Sturzenegger, A.; Huttner, D.; Janscak, P.; Hickson, I.D.; Gileadi, O.; Rothenberg, E. Mechanistic insight into the interaction of BLM helicase with intra-strand G-quadruplex structures. Nat. Commun. 2014. [Google Scholar] [CrossRef] [PubMed]
  118. Huber, M.D.; Lee, D.C.; Maizels, N. G4 DNA unwinding by BLM and Sgs1p: Substrate specificity and substrate-specific inhibition. Nucleic Acids Res. 2002, 30, 3954–3961. [Google Scholar] [CrossRef] [PubMed]
  119. Safa, L.; Gueddouda, N.M.; Thiebaut, F.; Delagoutte, E.; Petruseva, I.; Lavrik, O.; Mendoza, O.; Bourdoncle, A.; Alberti, P.; Riou, J.F.; et al. 5’ to 3’ unfolding directionality of DNA secondary structures by replication protein A: G-quadruplexes and duplexes. J. Biol. Chem. 2016, 291, 21246–21256. [Google Scholar] [CrossRef] [PubMed]
  120. Salas, T.R.; Petruseva, I.; Lavrik, O.; Bourdoncle, A.; Mergny, J.L.; Favre, A.; Saintomé, C. Human replication protein A unfolds telomeric G-quadruplexes. Nucleic Acids Res. 2006, 34, 4857–4865. [Google Scholar] [CrossRef] [PubMed]
  121. Zaug, A.J.; Podell, E.R.; Cech, T.R. Human POT1 disrupts telomeric G-quadruplexes allowing telomerase extension in vitro. Proc. Natl. Acad. Sci. USA 2005, 102, 10864–10869. [Google Scholar] [CrossRef] [PubMed]
  122. Gomez, D.; O’Donohue, M.F.; Wenner, T.; Douarre, C.; Macadré, J.; Koebel, P.; Giraud-Panis, M.J.; Kaplan, H.; Kolkes, A.; Shin-Ya, K.; et al. The G-quadruplex ligand telomestatin inhibits POT1 binding to telomeric sequences in vitro and induces GFP-POT1 dissociation from telomeres in human cells. Cancer Res. 2006, 66, 6908–6912. [Google Scholar] [CrossRef] [PubMed]
  123. Wang, H.; Nora, G.J.; Ghodke, H.; Opresko, P.L. Single molecule studies of physiologically relevant telomeric tails reveal POT1 mechanism for promoting G-quadruplex unfolding. J. Biol. Chem. 2011, 286, 7479–7489. [Google Scholar] [CrossRef] [PubMed]
  124. Hwang, H.; Buncher, N.; Opresko, P.L.; Myong, S. POT1-TPP1 regulates telomeric overhang structural dynamics. Structure 2012, 20, 1872–1880. [Google Scholar] [CrossRef] [PubMed]
  125. Chan, K.L.; North, P.S.; Hickson, I.D. BLM is required for faithful chromosome segregation and its localization defines a class of ultrafine anaphase bridges. EMBO J. 2007, 26, 3397–3409. [Google Scholar] [CrossRef] [PubMed]
  126. Sarkar, J.; Wan, B.; Yin, J.; Vallabhaneni, H.; Horvath, K.; Kulikowicz, T.; Bohr, V.A.; Zhang, Y.; Lei, M.; Liu, Y. SLX4 contributes to telomere preservation and regulated processing of telomeric joint molecule intermediates. Nucleic Acids Res. 2015, 43, 5912–5923. [Google Scholar] [CrossRef] [PubMed]
  127. Mankouri, H.W.; Huttner, D.; Hickson, I.D. How unfinished business from S-phase affects mitosis and beyond. EMBO J. 2013, 32, 2661–2671. [Google Scholar] [CrossRef] [PubMed]
  128. Opresko, P.L.; Von Kobbe, C.; Laine, J.P.; Harrigan, J.; Hickson, I.D.; Bohr, V.A. Telomere-binding protein TRF2 binds to and stimulates the Werner and Bloom syndrome helicases. J. Biol. Chem. 2002, 277, 41110–41119. [Google Scholar] [CrossRef] [PubMed]
  129. Opresko, P.L.; Mason, P.A.; Podell, E.R.; Lei, M.; Hickson, I.D.; Cech, T.R.; Bohr, V.A. POT1 stimulates RecQ helicases WRN and BLM to unwind telomeric DNA substrates. J. Biol. Chem. 2005, 280, 32069–32080. [Google Scholar] [CrossRef] [PubMed]
  130. Stavropoulos, D.J.; Bradshaw, P.S.; Li, X.; Pasic, I.; Truong, K.; Ikura, M.; Ungrin, M.; Meyn, M.S. The Bloom syndrome helicase BLM interacts with TRF2 in ALT cells and promotes telomeric DNA synthesis. Hum. Mol. Genet. 2002, 11, 3135–3144. [Google Scholar] [CrossRef] [PubMed]
  131. Opresko, P.L.; Otterlei, M.; Graakjær, J.; Bruheim, P.; Dawut, L.; Kølvraa, S.; May, A.; Seidman, M.M.; Bohr, V.A. The werner syndrome helicase and exonuclease cooperate to resolve telomeric D loops in a manner regulated by TRF1 and TRF2. Mol. Cell 2004, 14, 763–774. [Google Scholar] [CrossRef] [PubMed]
  132. Machwe, A.; Lozada, E.; Wold, M.S.; Li, G.M.; Orren, D.K. Molecular cooperation between the Werner syndrome protein and replication protein A in relation to replication fork blockage. J. Biol. Chem. 2011, 286, 3497–3508. [Google Scholar] [CrossRef] [PubMed]
  133. Edwards, D.N.; Machwe, A.; Chen, L.; Bohr, V.A.; Orren, D.K. The DNA structure and sequence preferences of WRN underlie its function in telomeric recombination events. Nat. Commun. 2015. [Google Scholar] [CrossRef] [PubMed]
  134. Arnoult, N.; Saintome, C.; Ourliac-Garnier, I.; Riou, J.F.; Londoño-Vallejo, A. Human POT1 is required for efficient telomere C-rich strand replication in the absence of WRN. Genes Dev. 2009, 23, 2915–2924. [Google Scholar] [CrossRef] [PubMed]
  135. Crabbe, L.; Verdun, R.E.; Haggblom, C.I.; Karlseder, J. Defective telomere lagging strand synthesis in cells lacking WRN helicase activity. Science 2004, 306, 1951–1953. [Google Scholar] [CrossRef] [PubMed]
  136. Rizzo, A.; Salvati, E.; Porru, M.; D’Angelo, C.; Stevens, M.F.; D’Incalci, M.; Leonetti, C.; Gilson, E.; Zupi, G.; Biroccio, A. Stabilization of quadruplex DNA perturbs telomere replication leading to the activation of an ATR-dependent ATM signaling pathway. Nucleic Acids Res. 2009, 37, 5353–5364. [Google Scholar] [CrossRef] [PubMed]
  137. Gocha, A.R.S.; Acharya, S.; Groden, J. WRN loss induces switching of telomerase-independent mechanisms of telomere elongation. PLoS ONE 2014, 9, e93991. [Google Scholar] [CrossRef] [PubMed]
  138. Laud, P.R.; Multani, A.S.; Bailey, S.M.; Wu, L.; Ma, J.; Kingsley, C.; Lebel, M.; Pathak, S.; DePinho, R.A.; Chang, S. Elevated telomere-telomere recombination in WRN-deficient, telomere dysfunctional cells promotes escape from senescence and engagement of the ALT pathway. Genes Dev. 2005, 19, 2560–2570. [Google Scholar] [CrossRef] [PubMed]
  139. Mendez-Bermudez, A.; Hidalgo-Bravo, A.; Cotton, V.E.; Gravani, A.; Jeyapalan, J.N.; Royle, N.J. The roles of WRN and BLM RecQ helicases in the Alternative Lengthening of Telomeres. Nucleic Acids Res. 2012, 40, 10809–10820. [Google Scholar] [CrossRef] [PubMed]
  140. Edwards, D.N.; Orren, D.K.; Machwe, A. Strand exchange of telomeric DNA catalyzed by the Werner syndrome protein (WRN) is specifically stimulated by TRF2. Nucleic Acids Res. 2014, 42, 7748–7761. [Google Scholar] [CrossRef] [PubMed]
  141. Ghosh, A.K.; Rossi, M.L.; Singh, D.K.; Dunn, C.; Ramamoorthy, M.; Croteau, D.L.; Liu, Y.; Bohr, V.A. RECQL4, the protein mutated in Rothmund-Thomson syndrome, functions in telomere maintenance. J. Biol. Chem. 2012, 287, 196–209. [Google Scholar] [CrossRef] [PubMed]
  142. Matsuno, K.; Kumano, M.; Kubota, Y.; Hashimoto, Y.; Takisawa, H. The N-terminal noncatalytic region of Xenopus RecQ4 is required for chromatin binding of DNA polymerase alpha in the initiation of DNA replication. Mol. Cell. Biol. 2006, 26, 4843–4852. [Google Scholar] [CrossRef] [PubMed]
  143. Sangrithi, M.N.; Bernal, J.A.; Madine, M.; Philpott, A.; Lee, J.; Dunphy, W.G.; Venkitaraman, A.R. Initiation of DNA replication requires the RECQL4 protein mutated in Rothmund-Thomson syndrome. Cell 2005, 121, 887–898. [Google Scholar] [CrossRef] [PubMed]
  144. Rossi, M.L.; Ghosh, A.K.; Kulikowicz, T.; Croteau, D.L.; Bohr, V.A. Conserved helicase domain of human RecQ4 is required for strand annealing-independent DNA unwinding. DNA Repair 2010, 9, 796–804. [Google Scholar] [CrossRef] [PubMed]
  145. Keller, H.; Kiosze, K.; Sachsenweger, J.; Haumann, S.; Ohlenschläger, O.; Nuutinen, T.; Syväoja, J.E.; Görlach, M.; Grosse, F.; Pospiech, H. The intrinsically disordered amino-terminal region of human RecQL4: Multiple DNA-binding domains confer annealing, strand exchange and G4 DNA binding. Nucleic Acids Res. 2014, 42, 12614–12627. [Google Scholar] [CrossRef] [PubMed]
  146. Popuri, V.; Hsu, J.; Khadka, P.; Horvath, K.; Liu, Y.; Croteau, D.L.; Bohr, V.A. Human RECQL1 participates in telomere maintenance. Nucleic Acids Res. 2014, 42, 5671–5688. [Google Scholar] [CrossRef] [PubMed]
  147. Sami, F.; Lu, X.; Parvathaneni, S.; Roy, R.; Gary, R.K.; Sharma, S. RECQ1 interacts with FEN-1 and promotes binding of FEN-1 to telomeric chromatin. Biochem. J. 2015, 468, 227–244. [Google Scholar] [CrossRef] [PubMed]
  148. Popuri, V.; Bachrati, C.Z.; Muzzolini, L.; Mosedale, G.; Costantini, S.; Giacomini, E.; Hickson, I.D.; Vindigni, A. The human RecQ helicases, BLM and RECQ1, display distinct DNA substrate specificities. J. Biol. Chem. 2008, 283, 17766–17776. [Google Scholar] [CrossRef] [PubMed]
  149. Sommers, J.A.; Banerjee, T.; Hinds, T.; Wan, B.; Wold, M.S.; Lei, M.; Brosh, R.M., Jr. Novel function of the fanconi anemia group J or RECQ1 helicase to disrupt protein-DNA complexes in a replication protein A-stimulated manner. J. Biol. Chem. 2014, 289, 19928–19941. [Google Scholar] [CrossRef] [PubMed]
  150. Vannier, J.B.; Sarek, G.; Boulton, S.J. RTEL1: Functions of a disease-associated helicase. Trends Cell Biol. 2014, 24, 416–425. [Google Scholar] [CrossRef] [PubMed]
  151. Ding, H.; Schertzer, M.; Wu, X.; Gertsenstein, M.; Selig, S.; Kammori, M.; Pourvali, R.; Poon, S.; Vulto, I.; Chavez, E.; et al. Regulation of murine telomere length by Rtel: An essential gene encoding a helicase-like protein. Cell 2004, 117, 873–886. [Google Scholar] [CrossRef] [PubMed]
  152. Uringa, E.J.; Lisaingo, K.; Pickett, H.A.; Brind’Amour, J.; Rohde, J.H.; Zelensky, A.; Essers, J.; Lansdorp, P.M. RTEL1 contributes to DNA replication and repair and telomere maintenance. Mol. Biol. Cell 2012, 23, 2782–2792. [Google Scholar] [CrossRef] [PubMed]
  153. Vannier, J.B.; Sandhu, S.; Petalcorin, M.I.; Wu, X.; Nabi, Z.; Ding, H.; Boulton, S.J. RTEL1 Is a Replisome-Associated Helicase That Promotes Telomere and Genome-Wide Replication. Science 2013, 342, 239–242. [Google Scholar] [CrossRef] [PubMed]
  154. Youds, J.L.; Mets, D.G.; Mcllwraith, M.J.; Martin, J.S.; Ward, J.D.; ONeil, N.J.; Rose, A.M.; West, S.C.; Meyer, B.J.; Boulton, S.J. RTEL-1 Enforces Meiotic Crossover Interference and Homeostasis. Science 2010, 327, 1254–1258. [Google Scholar] [CrossRef] [PubMed]
  155. Stansel, R.M.; De Lange, T.; Griffith, J.D. T-loop assembly in vitro involves binding of TRF2 near the 3′ telomeric overhang. EMBO J. 2001, 20, 5532–5540. [Google Scholar] [CrossRef] [PubMed]
  156. Svendsen, J.M.; Smogorzewska, A.; Sowa, M.E.; O’Connell, B.C.; Gygi, S.P.; Elledge, S.J.; Harper, J.W. Mammalian BTBD12/SLX4 Assembles A Holliday Junction Resolvase and Is Required for DNA Repair. Cell 2009, 138, 63–77. [Google Scholar] [CrossRef] [PubMed]
  157. Fekairi, S.; Scaglione, S.; Chahwan, C.; Taylor, E.R.; Tissier, A.; Coulon, S.; Dong, M.Q.; Ruse, C.; Yates, J.R.; Russell, P.; et al. Human SLX4 Is a Holliday Junction Resolvase Subunit that Binds Multiple DNA Repair/Recombination Endonucleases. Cell 2009, 138, 78–89. [Google Scholar] [CrossRef] [PubMed]
  158. Muñoz, I.M.; Hain, K.; Déclais, A.C.; Gardiner, M.; Toh, G.W.; Sanchez-Pulido, L.; Heuckmann, J.M.; Toth, R.; Macartney, T.; Eppink, B.; et al. Coordination of Structure-Specific Nucleases by Human SLX4/BTBD12 Is Required for DNA Repair. Mol. Cell 2009, 35, 116–127. [Google Scholar] [CrossRef] [PubMed]
  159. Yin, J.; Wan, B.; Sarkar, J.; Horvath, K.; Wu, J.; Chen, Y.; Cheng, G.; Wan, K.; Chin, P.; Lei, M.; et al. Dimerization of SLX4 contributes to functioning of the SLX4-nuclease complex. Nucleic Acids Res. 2016, 44, 4871–4880. [Google Scholar] [CrossRef] [PubMed]
  160. Wyatt, H.D.M.; Sarbajna, S.; Matos, J.; West, S.C. Coordinated actions of SLX1-SLX4 and MUS81-EME1 for holliday junction resolution in human cells. Mol. Cell 2013, 52, 234–247. [Google Scholar] [CrossRef] [PubMed]
  161. Garner, E.; Kim, Y.; Lach, F.P.; Kottemann, M.C.; Smogorzewska, A. Human GEN1 and the SLX4-Associated Nucleases MUS81 and SLX1 Are Essential for the Resolution of Replication-Induced Holliday Junctions. Cell Rep. 2013, 5, 207–215. [Google Scholar] [CrossRef] [PubMed]
  162. Castor, D.; Nair, N.; Déclais, A.C.; Lachaud, C.; Toth, R.; Macartney, T.J.; Lilley, D.M.J.; Arthur, J.S.C.; Rouse, J. Cooperative control of holliday junction resolution and DNA Repair by the SLX1 and MUS81-EME1 nucleases. Mol. Cell 2013, 52, 221–233. [Google Scholar] [CrossRef] [PubMed]
  163. Kim, Y.; Spitz, G.S.; Veturi, U.; Lach, F.P.; Auerbach, A.D.; Smogorzewska, A. Regulation of multiple DNA repair pathways by the Fanconi anemia protein SLX4. Blood 2013, 121, 54–63. [Google Scholar] [CrossRef] [PubMed]
  164. Saint-Léger, A.; Koelblen, M.; Civitelli, L.; Bah, A.; Djerbi, N.; Giraud-Panis, M.J.; Londonõ-Vallejo, A.; Ascenzioni, F.; Gilson, E. The basic N-terminal domain of TRF2 limits recombination endonuclease action at human telomeres. Cell Cycle 2014, 13, 2469–2479. [Google Scholar] [CrossRef] [PubMed]
  165. Pickett, H.A.; Cesare, A.J.; Johnston, R.L.; Neumann, A.A.; Reddel, R.R. Control of telomere length by a trimming mechanism that involves generation of t-circles. EMBO J. 2009, 28, 799–809. [Google Scholar] [CrossRef] [PubMed]
  166. Guervilly, J.H.; Takedachi, A.; Naim, V.; Scaglione, S.; Chawhan, C.; Lovera, Y.; Despras, E.; Kuraoka, I.; Kannouche, P.; Rosselli, F.; et al. The SLX4 complex is a SUMO E3 ligase that impacts on replication stress outcome and genome stability. Mol. Cell 2015, 57, 123–137. [Google Scholar] [CrossRef] [PubMed]
  167. Ouyang, J.; Garner, E.; Hallet, A.; Nguyen, H.D.; Rickman, K.A.; Gill, G.; Smogorzewska, A.; Zou, L. Noncovalent Interactions with SUMO and Ubiquitin Orchestrate Distinct Functions of the SLX4 Complex in Genome Maintenance. Mol. Cell 2015, 57, 108–122. [Google Scholar] [CrossRef] [PubMed]
  168. González-Prieto, R.; Cuijpers, S.A.G.; Luijsterburg, M.S.; van Attikum, H.; Vertegaal, A.C.O. SUMOylation and PARylation cooperate to recruit and stabilize SLX4 at DNA damage sites. EMBO Rep. 2015, 16, 512–519. [Google Scholar] [CrossRef] [PubMed]
  169. Saharia, A.; Guittat, L.; Crocker, S.; Lim, A.; Steffen, M.; Kulkarni, S.; Stewart, S.A. Flap Endonuclease 1 Contributes to Telomere Stability. Curr. Biol. 2008, 18, 496–500. [Google Scholar] [CrossRef] [PubMed]
  170. Saharia, A.; Teasley, D.C.; Duxin, J.P.; Dao, B.; Chiappinelli, K.B.; Stewart, S.A. FEN1 ensures telomere stability by facilitating replication fork re-initiation. J. Biol. Chem. 2010, 285, 27057–27066. [Google Scholar] [CrossRef] [PubMed]
  171. Teasley, D.C.; Parajuli, S.; Nguyen, M.; Moore, H.R.; Alspach, E.; Lock, Y.J.; Honaker, Y.; Saharia, A.; Piwnica-Worms, H.; Stewart, S.A. Flap endonuclease 1 limits telomere fragility on the leading strand. J. Biol. Chem. 2015, 290, 15133–15145. [Google Scholar] [CrossRef] [PubMed]
  172. Vallur, A.C.; Maizels, N. Distinct activities of exonuclease 1 and flap endonuclease 1 at telomeric G4 DNA. PLoS ONE 2010, 5, e8908. [Google Scholar] [CrossRef] [PubMed]
  173. León-Ortiz, A.M.; Svendsen, J.; Boulton, S.J. Metabolism of DNA secondary structures at the eukaryotic replication fork. DNA Repair 2014, 19, 152–162. [Google Scholar] [CrossRef] [PubMed]
  174. Lin, W.; Sampathi, S.; Dai, H.; Liu, C.; Zhou, M.; Hu, J.; Huang, Q.; Campbell, J.; Shin-Ya, K.; Zheng, L.; et al. Mammalian DNA2 helicase/nuclease cleaves G-quadruplex DNA and is required for telomere integrity. EMBO J. 2013, 32, 1425–1439. [Google Scholar] [CrossRef] [PubMed]
  175. Maquat, L.E.; Gong, C. Gene expression networks: Competing mRNA decay pathways in mammalian cells. Biochem. Soc. Trans. 2009, 37, 1287–1292. [Google Scholar] [CrossRef] [PubMed]
  176. Nicholson, P.; Yepiskoposyan, H.; Metze, S.; Zamudio Orozco, R.; Kleinschmidt, N.; Mühlemann, O. Nonsense-mediated mRNA decay in human cells: Mechanistic insights, functions beyond quality control and the double-life of NMD factors. Cell. Mol. Life Sci. 2010, 67, 677–700. [Google Scholar] [CrossRef] [PubMed]
  177. Azzalin, C.M. Upf1. Nucleus 2012, 3, 16–21. [Google Scholar] [CrossRef] [PubMed]
  178. Chawla, R.; Redon, S.; Raftopoulou, C.; Wischnewski, H.; Gagos, S.; Azzalin, C.M. Human UPF1 interacts with TPP1 and telomerase and sustains telomere leading-strand replication. EMBO J. 2011, 30, 4047–4058. [Google Scholar] [CrossRef] [PubMed]
  179. 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. [Google Scholar] [CrossRef] [PubMed]
  180. Arora, R.; Azzalin, C.M. Telomere elongation chooses TERRA ALTernatives. RNA Biol. 2015, 12, 938–941. [Google Scholar] [CrossRef] [PubMed]
  181. Arora, R.; Lee, Y.; Wischnewski, H.; Brun, C.M.; Schwarz, T.; Azzalin, C.M. RNaseH1 regulates TERRA-telomeric DNA hybrids and telomere maintenance in ALT tumour cells. Nat. Commun. 2014, 5, 5220. [Google Scholar] [CrossRef] [PubMed]
  182. Heaphy, C.M.; De Wilde, R.F.; Jiao, Y.; Klein, A.P.; Edil, B.H.; Shi, C.; Bettegowda, C.; Rodriguez, F.J.; Eberhart, C.G.; Hebbar, S.; et al. Altered telomeres in tumors with ATRX and DAXX mutations. Science 2011, 333, 425. [Google Scholar] [CrossRef] [PubMed]
  183. Lovejoy, C.A.; Li, W.; Reisenweber, S.; Thongthip, S.; Bruno, J.; De Lange, T.; De, S.; Petrini, J.H.J.; Sung, P.A.; Jasin, M.; et al. Loss of ATRX, genome instability, and an altered DNA damage response are hallmarks of the alternative lengthening of Telomeres pathway. PLoS Genet. 2012, 8, e1002772. [Google Scholar] [CrossRef] [PubMed]
  184. Schwartzentruber, J.; Korshunov, A.; Liu, X.Y.; Jones, D.T.; Pfaff, E.; Jacob, K.; Sturm, D.; Fontebasso, A.M.; Quang, D.A.; Tönjes, M.; et al. Driver mutations in histone H3. 3 and chromatin remodelling genes in paediatric glioblastoma. Nature 2012, 482, 226–231. [Google Scholar] [CrossRef] [PubMed]
  185. Morrison, A.J.; Shen, X. Chromatin remodelling beyond transcription: The INO80 and SWR1 complexes. Nat. Rev. Mol. Cell Biol. 2009, 10, 373–384. [Google Scholar] [CrossRef] [PubMed]
  186. Min, J.N.; Tian, Y.; Xiao, Y.; Wu, L.; Li, L.; Chang, S. The mINO80 chromatin remodeling complex is required for efficient telomere replication and maintenance of genome stability. Cell Res. 2013, 23, 1396–1413. [Google Scholar] [CrossRef] [PubMed]
  187. Yan, Y.; Akhter, S.; Zhang, X.; Legerski, R. The multifunctional SNM1 gene family: Not just nucleases. Futur. Oncol. 2010, 6, 1015–1029. [Google Scholar] [CrossRef] [PubMed]
  188. Demuth, I.; Bradshaw, P.S.; Lindner, A.; Anders, M.; Heinrich, S.; Kallenbach, J.; Schmelz, K.; Digweed, M.; Meyn, M.S.; Concannon, P. Endogenous hSNM1B/Apollo interacts with TRF2 and stimulates ATM in response to ionizing radiation. DNA Repair 2008, 7, 1192–1201. [Google Scholar] [CrossRef] [PubMed]
  189. Demuth, I.; Digweed, M.; Concannon, P. Human SNM1B is required for normal cellular response to both DNA interstrand crosslink-inducing agents and ionizing radiation. Oncogene 2004, 23, 8611–8618. [Google Scholar] [CrossRef] [PubMed]
  190. Bae, J.B.; Mukhopadhyay, S.S.; Liu, L.; Zhang, N.; Tan, J.; Akhter, S.; Liu, X.; Shen, X.; Li, L.; Legerski, R.J. Snm1B/Apollo mediates replication fork collapse and S Phase checkpoint activation in response to DNA interstrand cross-links. Oncogene 2008, 27, 5045–5056. [Google Scholar] [CrossRef] [PubMed]
  191. Mason, J.M.; Das, I.; Arlt, M.; Patel, N.; Kraftson, S.; Glover, T.W.; Sekiguchi, J.M. The SNM1B/Apollo DNA nuclease functions in resolution of replication stress and maintenance of common fragile site stability. Hum. Mol. Genet. 2013, 22, 4901–4913. [Google Scholar] [CrossRef] [PubMed]
  192. Mason, J.M.; Sekiguchi, J.M. Snm1B/Apollo functions in the Fanconi anemia pathway in response to DNA interstrand crosslinks. Hum. Mol. Genet. 2011, 20, 2549–2559. [Google Scholar] [CrossRef] [PubMed]
  193. Wu, P.; van Overbeek, M.; Rooney, S.; de Lange, T. Apollo Contributes to G Overhang Maintenance and Protects Leading-End Telomeres. Mol. Cell 2010, 39, 606–617. [Google Scholar] [CrossRef] [PubMed]
  194. Freibaum, B.D.; Counter, C.M. hSnm1B is a novel telomere-associated protein. J. Biol. Chem. 2006, 281, 15033–15036. [Google Scholar] [CrossRef] [PubMed]
  195. Van Overbeek, M.; de Lange, T. Apollo, an Artemis-Related Nuclease, Interacts with TRF2 and Protects Human Telomeres in S Phase. Curr. Biol. 2006, 16, 1295–1302. [Google Scholar] [CrossRef] [PubMed]
  196. Lenain, C.; Bauwens, S.; Amiard, S.; Brunori, M.; Giraud-Panis, M.J.; Gilson, E. The Apollo 5’ Exonuclease Functions Together with TRF2 to Protect Telomeres from DNA Repair. Curr. Biol. 2006, 16, 1303–1310. [Google Scholar] [CrossRef] [PubMed]
  197. Ye, J.; Lenain, C.; Bauwens, S.; Rizzo, A.; Saint-Léger, A.; Poulet, A.; Benarroch, D.; Magdinier, F.; Morere, J.; Amiard, S.; et al. TRF2 and Apollo Cooperate with Topoisomerase 2α to Protect Human Telomeres from Replicative Damage. Cell 2010, 142, 230–242. [Google Scholar] [CrossRef] [PubMed]
  198. Touzot, F.; Callebaut, I.; Soulier, J.; Gaillard, L.; Azerrad, C.; Durandy, A.; Fischer, A.; de Villartay, J.P.; Revy, P. Function of Apollo (SNM1B) at telomere highlighted by a splice variant identified in a patient with Hoyeraal-Hreidarsson syndrome. Proc. Natl. Acad. Sci. USA 2010, 107, 10097–10102. [Google Scholar] [CrossRef] [PubMed]
  199. Lam, Y.C.; Akhter, S.; Gu, P.; Ye, J.; Poulet, A.; Giraud-Panis, M.J.; Bailey, S.M.; Gilson, E.; Legerski, R.J.; Chang, S. SNMIB/Apollo protects leading-strand telomeres against NHEJ-mediated repair. EMBO J. 2010, 29, 2230–2241. [Google Scholar] [CrossRef] [PubMed]
  200. Wu, P.; Takai, H.; de Lange, T. Telomeric 3’ overhangs derive from resection by Exo1 and apollo and fill-in by POT1b-associated CST. Cell 2012, 150, 39–52. [Google Scholar] [CrossRef] [PubMed]
  201. Denchi, E.L.; de Lange, T. Protection of telomeres through independent control of ATM and ATR by TRF2 and POT1. Nature 2007, 448, 1068–1071. [Google Scholar] [CrossRef] [PubMed]
  202. Guo, X.; Deng, Y.; Lin, Y.; Cosme-Blanco, W.; Chan, S.; He, H.; Yuan, G.; Brown, E.J.; Chang, S. Dysfunctional telomeres activate an ATM-ATR-dependent DNA damage response to suppress tumorigenesis. EMBO J. 2007, 26, 4709–4719. [Google Scholar] [CrossRef] [PubMed]
  203. Gong, Y.; de Lange, T. A Shld1-Controlled POT1a Provides Support for Repression of ATR Signaling at Telomeres through RPA Exclusion. Mol. Cell 2010, 40, 377–387. [Google Scholar] [CrossRef] [PubMed]
  204. Flynn, R.L.; Centore, R.C.; O’Sullivan, R.J.; Rai, R.; Tse, A.; Songyang, Z.; Chang, S.; Karlseder, J.; Zou, L. TERRA and hnRNPA1 orchestrate an RPA-to-POT1 switch on telomeric single-stranded DNA. Nature 2011, 471, 532–536. [Google Scholar] [CrossRef] [PubMed]
  205. Takai, K.K.; Kibe, T.; Donigian, J.R.; Frescas, D.; de Lange, T. Telomere Protection by TPP1/POT1 Requires Tethering to TIN2. Mol. Cell 2011, 44, 647–659. [Google Scholar] [CrossRef] [PubMed]
  206. Kibe, T.; Zimmermann, M.; de Lange, T. TPP1 Blocks an ATR-Mediated Resection Mechanism at Telomeres. Mol. Cell 2016, 61, 236–246. [Google Scholar] [CrossRef] [PubMed]
  207. Hockemeyer, D.; Collins, K. Control of telomerase action at human telomeres. Nat. Struct. Mol. Biol. 2015, 22, 848–852. [Google Scholar] [CrossRef] [PubMed]
  208. Lloyd, N.R.; Dickey, T.H.; Hom, R.A.; Wuttke, D.S. Tying up the ends: Plasticity in the recognition of single-stranded DNA at telomeres. Biochemistry 2016, 55, 5326–5340. [Google Scholar] [CrossRef] [PubMed]
  209. Robles-Espinoza, C.D.; Harland, M.; Ramsay, A.J.; Aoude, L.G.; Quesada, V.; Ding, Z.; Pooley, K.A.; Pritchard, A.L.; Tiffen, J.C.; Petljak, M.; et al. POT1 loss-of-function variants predispose to familial melanoma. Nat. Genet. 2014, 46, 478–481. [Google Scholar] [CrossRef] [PubMed]
  210. Shi, J.; Yang, X.R.; Ballew, B.; Rotunno, M.; Calista, D.; Fargnoli, M.C.; Ghiorzo, P.; Bressac-de Paillerets, B.; Nagore, E.; Avril, M.F.; et al. Rare missense variants in POT1 predispose to familial cutaneous malignant melanoma. Nat. Genet. 2014, 46, 482–486. [Google Scholar] [CrossRef] [PubMed]
  211. Ramsay, A.J.; Quesada, V.; Foronda, M.; Conde, L.; Martínez-Trillos, A.; Villamor, N.; Rodríguez, D.; Kwarciak, A.; Garabaya, C.; Gallardo, M.; et al. POT1 mutations cause telomere dysfunction in chronic lymphocytic leukemia. Nat. Genet. 2013, 45, 526–530. [Google Scholar] [CrossRef] [PubMed]
  212. Pinzaru, A.M.; Hom, R.A.; Beal, A.; Phillips, A.F.; Ni, E.; Cardozo, T.; Nair, N.; Choi, J.; Wuttke, D.S.; Sfeir, A.; et al. Telomere Replication Stress Induced by POT1 Inactivation Accelerates Tumorigenesis. Cell Rep. 2016, 15, 2170–2184. [Google Scholar] [CrossRef] [PubMed]
  213. Sarthy, J.; Bae, N.S.; Scrafford, J.; Baumann, P. Human RAP1 inhibits non-homologous end joining at telomeres. EMBO J. 2009, 28, 3390–3399. [Google Scholar] [CrossRef] [PubMed]
  214. Martinez, P.; Thanasoula, M.; Carlos, A.R.; Gómez-López, G.; Tejera, A.M.; Schoeftner, S.; Dominguez, O.; Pisano, D.G.; Tarsounas, M.; Blasco, M.A. Mammalian Rap1 controls telomere function and gene expression through binding to telomeric and extratelomeric sites. Nat. Cell Biol. 2010, 12, 768–780. [Google Scholar] [CrossRef] [PubMed]
  215. Sfeir, A.; Kabir, S.; van Overbeek, M.; Celli, G.B.; de Lange, T. Loss of Rap1 Induces Telomere Recombination in the Absence of NHEJ or a DNA damage Signal. Science 2010, 327, 1657–1661. [Google Scholar] [CrossRef] [PubMed]
  216. Miyake, Y.; Nakamura, M.; Nabetani, A.; Shimamura, S.; Tamura, M.; Yonehara, S.; Saito, M.; Ishikawa, F. RPA-like Mammalian Ctc1-Stn1-Ten1 Complex Binds to Single-Stranded DNA and Protects Telomeres Independently of the Pot1 Pathway. Mol. Cell 2009, 36, 193–206. [Google Scholar] [CrossRef] [PubMed]
  217. Surovtseva, Y.V.; Churikov, D.; Boltz, K.A.; Song, X.; Lamb, J.C.; Warrington, R.; Leehy, K.; Heacock, M.; Price, C.M.; Shippen, D.E. Conserved Telomere Maintenance Component 1 Interacts with STN1 and Maintains Chromosome Ends in Higher Eukaryotes. Mol. Cell 2009, 36, 207–218. [Google Scholar] [CrossRef] [PubMed]
  218. Rice, C.; Skordalakes, E. Structure and function of the telomeric CST complex. Comput. Struct. Biotechnol. J. 2016, 14, 161–167. [Google Scholar] [CrossRef] [PubMed]
  219. Wan, M.; Qin, J.; Songyang, Z.; Liu, D. OB fold-containing protein 1 (OBFC1), a human homolog of yeast Stn1, associates with TPP1 and is implicated in telomere length regulation. J. Biol. Chem. 2009, 284, 26725–26731. [Google Scholar] [CrossRef] [PubMed]
  220. Chen, L.Y.; Redon, S.; Lingner, J. The human CST complex is a terminator of telomerase activity. Nature 2012, 488, 540–544. [Google Scholar] [CrossRef] [PubMed]
  221. Casteel, D.E.; Zhuang, S.; Zeng, Y.; Perrino, F.W.; Boss, G.R.; Goulian, M.; Pilz, R.B. A DNA polymerase-α·primase cofactor with homology to replication protein A-32 regulates DNA replication in mammalian cells. J. Biol. Chem. 2009, 284, 5807–5818. [Google Scholar] [CrossRef] [PubMed]
  222. Nakaoka, H.; Nishiyama, A.; Saito, M.; Ishikawa, F. Xenopus laevis Ctc1-Stn1-Ten1 (xCST) protein complex is involved in priming DNA synthesis on single-stranded DNA template in Xenopus egg extract. J. Biol. Chem. 2012, 287, 619–627. [Google Scholar] [CrossRef] [PubMed]
  223. Lue, N.F.; Chan, J.; Wright, W.E.; Hurwitz, J. The CDC13-STN1-TEN1 complex stimulates Pol α activity by promoting RNA priming and primase-to-polymerase switch. Nat. Commun. 2014, 5, 5762. [Google Scholar] [CrossRef] [PubMed]
  224. Sun, J.; Yu, E.Y.; Yang, Y.; Confer, L.A.; Sun, S.H.; Wan, K.; Lue, N.F.; Lei, M. Stn1-Ten1 is an Rpa2-Rpa3-like complex at telomeres. Genes Dev. 2009, 23, 2900–2914. [Google Scholar] [CrossRef] [PubMed]
  225. Gelinas, A.D.; Paschini, M.; Reyes, F.E.; Héroux, A.; Batey, R.T.; Lundblad, V.; Wuttke, D.S. Telomere capping proteins are structurally related to RPA with an additional telomere-specific domain. Proc. Natl. Acad. Sci. USA 2009, 106, 19298–19303. [Google Scholar] [CrossRef] [PubMed]
  226. Wang, F.; Stewart, J.A.; Kasbek, C.; Zhao, Y.; Wright, W.E.; Price, C.M. Human CST Has Independent Functions during Telomere Duplex Replication and C-Strand Fill-In. Cell Rep. 2012, 2, 1096–1103. [Google Scholar] [CrossRef] [PubMed]
  227. Verdun, R.E.; Karlseder, J. The DNA Damage Machinery and Homologous Recombination Pathway Act Consecutively to Protect Human Telomeres. Cell 2006, 127, 709–720. [Google Scholar] [CrossRef] [PubMed]
  228. McNees, C.J.; Tejera, A.M.; Martínez, P.; Murga, M.; Mulero, F.; Fernandez-Capetillo, O.; Blasco, M.A. ATR suppresses telomere fragility and recombination but is dispensable for elongation of short telomeres by telomerase. J. Cell Biol. 2010, 188, 639–652. [Google Scholar] [CrossRef] [PubMed]
  229. Pennarun, G.; Hoffschir, F.; Revaud, D.; Granotier, C.; Gauthier, L.R.; Mailliet, P.; Biard, D.S.; Boussin, F.D. ATR contributes to telomere maintenance in human cells. Nucleic Acids Res. 2010, 38, 2955–2963. [Google Scholar] [CrossRef] [PubMed]
  230. Boccardi, V.; Razdan, N.; Kaplunov, J.; Mundra, J.J.; Kimura, M.; Aviv, A.; Herbig, U. Stn1 is critical for telomere maintenance and long-term viability of somatic human cells. Aging Cell 2015, 14, 372–381. [Google Scholar] [CrossRef] [PubMed]
  231. Gu, P.; Min, J.N.; Wang, Y.; Huang, C.; Peng, T.; Chai, W.; Chang, S. CTC1 deletion results in defective telomere replication, leading to catastrophic telomere loss and stem cell exhaustion. EMBO J. 2012, 31, 2309–2321. [Google Scholar] [CrossRef] [PubMed]
  232. Stewart, J.A.; Wang, F.; Chaiken, M.F.; Kasbek, C.; Chastain, P.D.; Wright, W.E.; Price, C.M. Human CST promotes telomere duplex replication and general replication restart after fork stalling. EMBO J. 2012, 31, 3537–3549. [Google Scholar] [CrossRef] [PubMed]
  233. Kasbek, C.; Wang, F.; Price, C.M. Human TEN1 maintains telomere integrity and functions in genome-wide replication restart. J. Biol. Chem. 2013, 288, 30139–30150. [Google Scholar] [CrossRef] [PubMed]
  234. Bryan, C.; Rice, C.; Harkisheimer, M.; Schultz, D.C.; Skordalakes, E. Structure of the Human Telomeric Stn1-Ten1 Capping Complex. PLoS ONE 2013, 8, e66756. [Google Scholar] [CrossRef] [PubMed]
  235. Bhattacharjee, A.; Stewart, J.; Chaiken, M.; Price, C.M. STN1 OB Fold Mutation Alters DNA Binding and Affects Selective Aspects of CST Function. PLoS Genet. 2016, 12, e1006342. [Google Scholar] [CrossRef] [PubMed]
  236. Huang, C.; Dai, X.; Chai, W. Human Stn1 protects telomere integrity by promoting efficient lagging strand synthesis at telomeres and mediating C-strand fill-in. Cell Res. 2012, 22, 1681–1695. [Google Scholar] [CrossRef] [PubMed]
  237. Wang, F.; Stewart, J.; Price, C.M. Human CST abundance determines recovery from diverse forms of DNA damage and replication stress. Cell Cycle 2014, 13, 3488–3498. [Google Scholar] [CrossRef] [PubMed]
  238. Chastain, M.; Zhou, Q.; Shiva, O.; Fadri-Moskwik, M.; Whitmore, L.; Jia, P.; Dai, X.; Huang, C.; Ye, P.; Chai, W. Human CST Facilitates Genome-wide RAD51 Recruitment to GC-Rich Repetitive Sequences in Response to Replication Stress. Cell Rep. 2016, 16, 1300–1314. [Google Scholar] [CrossRef] [PubMed]
  239. Badie, S.; Escandell, J.M.; Bouwman, P.; Carlos, A.R.; Thanasoula, M.; Gallardo, M.M.; Suram, A.; Jaco, I.; Benitez, J.; Herbig, U.; et al. BRCA2 acts as a RAD51 loader to facilitate telomere replication and capping. Nat. Struct. Mol. Biol. 2010, 17, 1461–1469. [Google Scholar] [CrossRef] [PubMed]
  240. Petermann, E.; Orta, M.L.; Issaeva, N.; Schultz, N.; Helleday, T. Hydroxyurea-Stalled Replication Forks Become Progressively Inactivated and Require Two Different RAD51-Mediated Pathways for Restart and Repair. Mol. Cell 2010, 37, 492–502. [Google Scholar] [CrossRef] [PubMed]
  241. Muñoz, S.; Méndez, J. DNA replication stress: From molecular mechanisms to human disease. Chromosoma 2016. [Google Scholar]
  242. Costes, A.; Lambert, S. Homologous Recombination as a Replication Fork Escort: Fork-Protection and Recovery. Biomolecules 2012, 3, 39–71. [Google Scholar] [CrossRef] [PubMed]
  243. Sotiriou, S.K.; Kamileri, I.; Lugli, N.; Evangelou, K.; Da-Re, C.; Huber, F.; Padayachy, L.; Tardy, S.; Nicati, N.L.; Barriot, S.; et al. Mammalian RAD52 Functions in Break-Induced Replication Repair of Collapsed DNA Replication Forks. Mol. Cell 2016, 64, 1127–1134. [Google Scholar] [CrossRef] [PubMed]
  244. Costantino, L.; Sotiriou, S.K.; Rantala, J.K.; Magin, S.; Mladenov, E.; Helleday, T.; Haber, J.E.; Iliakis, G.; Kallioniemi, O.P.; Halazonetis, T.D. Break-induced replication repair of damaged forks induces genomic duplications in human cells. Science 2014, 343, 88–91. [Google Scholar] [CrossRef] [PubMed]
  245. Roumelioti, F.M.; Sotiriou, S.K.; Katsini, V.; Chiourea, M.; Halazonetis, T.D.; Gagos, S. Alternative lengthening of human telomeres is a conservative DNA replication process with features of break-induced replication. EMBO Rep. 2016, 17, 1731–1737. [Google Scholar] [CrossRef] [PubMed]
  246. Dilley, R.L.; Verma, P.; Cho, N.W.; Winters, H.D.; Wondisford, A.R.; Greenberg, R.A. Break-induced telomere synthesis underlies alternative telomere maintenance. Nature 2016, 539, 54–58. [Google Scholar] [CrossRef] [PubMed]
  247. Maestroni, L.; Matmati, S.; Coulon, S. Solving the Telomere Replication Problem. Genes 2017, 8, E55. [Google Scholar] [CrossRef] [PubMed]
  248. Leman, A.R.; Dheekollu, J.; Deng, Z.; Lee, S.W.; Das, M.M.; Lieberman, P.M.; Noguchi, E. Timeless preserves telomere length by promoting efficient DNA replication through human telomeres. Cell Cycle 2012, 11, 2337–2347. [Google Scholar] [CrossRef] [PubMed]
Figure 1. Initiation of eukaryotic DNA replication. Eukaryotic DNA replication is strictly regulated through two non-overlapping steps, origin licensing and firing. During the licensing step, which occurs from late M to G1 phases, the origin-recognition complex (ORC), and subsequently cell division cycle protein 6 (CDC6), DNA replication licensing factor Cdt1, and the MCM2–7 complex, bind to chromatin to form the pre-replication complex (pre-RC). The firing step requires S phase-specific kinases DBF4-dependent kinase (DDK) and cyclin-dependent kinase (CDK) that facilitate the loading of cell division cycle protein 45 (CDC45), the GINS complex (Sld5–Psf1–Psf2–Psf3), and several other proteins, to form the CMG (CDC45–MCM–GINS) helicase complex, which unwinds the DNA duplex, enabling DNA polymerases to initiate DNA synthesis at the replication fork. Multiple MCM2–7 double hexamers are loaded onto chromatin (not depicted in the figure). Licensed origins are sequentially activated during S phase. Some origins (called dormant origins) do not fire, are passively replicated in normal S phase, and act as backup origins upon replication stress.
Figure 1. Initiation of eukaryotic DNA replication. Eukaryotic DNA replication is strictly regulated through two non-overlapping steps, origin licensing and firing. During the licensing step, which occurs from late M to G1 phases, the origin-recognition complex (ORC), and subsequently cell division cycle protein 6 (CDC6), DNA replication licensing factor Cdt1, and the MCM2–7 complex, bind to chromatin to form the pre-replication complex (pre-RC). The firing step requires S phase-specific kinases DBF4-dependent kinase (DDK) and cyclin-dependent kinase (CDK) that facilitate the loading of cell division cycle protein 45 (CDC45), the GINS complex (Sld5–Psf1–Psf2–Psf3), and several other proteins, to form the CMG (CDC45–MCM–GINS) helicase complex, which unwinds the DNA duplex, enabling DNA polymerases to initiate DNA synthesis at the replication fork. Multiple MCM2–7 double hexamers are loaded onto chromatin (not depicted in the figure). Licensed origins are sequentially activated during S phase. Some origins (called dormant origins) do not fire, are passively replicated in normal S phase, and act as backup origins upon replication stress.
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Figure 2. The causes of replication fork stalling and the mechanisms that overcome telomeric obstacles. (a) The bulk of telomeric DNA is duplicated by conventional semi-conservative DNA replication. When a telomeric replication fork progresses unidirectionally toward the chromosomal end, G-rich and C-rich strands are replicated by lagging-strand and leading-strand synthesis, respectively. The replication machinery encounters various obstacles that compromise passage of the fork through the telomere. (b) Telomere-specific and non-telomere-specific proteins overcome the obstacles and prevent telomere fragility during DNA replication. Components of shelterin complex have prominent roles in the recruitment of accessory factors to telomeres, while recruitment of several factors, such as RecQ-like helicase 4 (RECQL4), DNA replication helicase/nuclease 2 (DNA2), chromatin remodeling proteins INO80 and ATRX, and the DNA repair protein breast cancer 2 (BRCA2), may be independent of shelterin. The defects in many of these factors result in fragile telomere phenotype, suggesting that these obstacles naturally exist in cells and are potential causes of genomic instability. Although these factors are also involved in other telomere-maintenance mechanisms or in general DNA metabolism, the focus here is on their functions in relation to telomeric DNA replication. When replication machinery unwinds duplex of telomeric DNA, G-quadruplex (G4) DNA structure can be formed on the G-rich strand of telomeres, which is basically used as a template of lagging strand synthesis. Werner syndrome RecQ-like helicase (WRN), Bloom syndrome RecQ-like helicase (BLM), and regulator of telomere elongation helicase 1 (RTEL1) resolve G4 DNAs in concert with single-stranded DNA binding proteins. These helicases also participate in resolution of D-loop (displacement loop) at the base of T-loop (telomere loop) structure. Disassembly of T-loop is required for replication fork to arrive at the end of chromosome. In the absence of RTEL1, persistent T-loop will be one of substrates of structure-specific SLX4-associated endonucleases. Structural barriers to replication fork are also generated by R-loop derived from telomeric repeat-containing RNA (TERRA) binding to telomeric DNA, and by topological stress along chromosome. Furthermore, homologous recombination of the telomeric DNA should be tightly regulated, because inappropriate recombination causes telomere defects such as multi-telomeric signals, sister telomere association, and end-to-end fusion of chromosomes. Proper recombination at the stalled replication fork is also essential for stability and restart of the fork. See the main text for details.
Figure 2. The causes of replication fork stalling and the mechanisms that overcome telomeric obstacles. (a) The bulk of telomeric DNA is duplicated by conventional semi-conservative DNA replication. When a telomeric replication fork progresses unidirectionally toward the chromosomal end, G-rich and C-rich strands are replicated by lagging-strand and leading-strand synthesis, respectively. The replication machinery encounters various obstacles that compromise passage of the fork through the telomere. (b) Telomere-specific and non-telomere-specific proteins overcome the obstacles and prevent telomere fragility during DNA replication. Components of shelterin complex have prominent roles in the recruitment of accessory factors to telomeres, while recruitment of several factors, such as RecQ-like helicase 4 (RECQL4), DNA replication helicase/nuclease 2 (DNA2), chromatin remodeling proteins INO80 and ATRX, and the DNA repair protein breast cancer 2 (BRCA2), may be independent of shelterin. The defects in many of these factors result in fragile telomere phenotype, suggesting that these obstacles naturally exist in cells and are potential causes of genomic instability. Although these factors are also involved in other telomere-maintenance mechanisms or in general DNA metabolism, the focus here is on their functions in relation to telomeric DNA replication. When replication machinery unwinds duplex of telomeric DNA, G-quadruplex (G4) DNA structure can be formed on the G-rich strand of telomeres, which is basically used as a template of lagging strand synthesis. Werner syndrome RecQ-like helicase (WRN), Bloom syndrome RecQ-like helicase (BLM), and regulator of telomere elongation helicase 1 (RTEL1) resolve G4 DNAs in concert with single-stranded DNA binding proteins. These helicases also participate in resolution of D-loop (displacement loop) at the base of T-loop (telomere loop) structure. Disassembly of T-loop is required for replication fork to arrive at the end of chromosome. In the absence of RTEL1, persistent T-loop will be one of substrates of structure-specific SLX4-associated endonucleases. Structural barriers to replication fork are also generated by R-loop derived from telomeric repeat-containing RNA (TERRA) binding to telomeric DNA, and by topological stress along chromosome. Furthermore, homologous recombination of the telomeric DNA should be tightly regulated, because inappropriate recombination causes telomere defects such as multi-telomeric signals, sister telomere association, and end-to-end fusion of chromosomes. Proper recombination at the stalled replication fork is also essential for stability and restart of the fork. See the main text for details.
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Figure 3. A model for the consequences of telomeric replication fork arrest and the different recovery mechanisms. (a) Telomeres present intrinsic obstacles, impeding the passage of replication machinery. Shelterin and accessory proteins prevent the fork stalling and repress improper recombination activity. Inappropriate recombination of the telomeric fork may result in telomere defects. (b) Persistent fork arrest might lead to fork collapse at telomeres. Restart of replication is necessary to avoid leaving an unreplicated region. Dormant origins could fire to complete telomere replication. Homologous recombination is also a general recovery mechanism from replication fork collapse. However, recombination at telomeres might increase the risk of cellular immortalization by ALT (alternative lengthening of telomeres). It is currently unknown how shelterin exactly contributes to restart of replication. BIR: break-induced replication; HDR: homology-directed repair; MTS: multi-telomeric signals; UFB: ultrafine anaphase bridge.
Figure 3. A model for the consequences of telomeric replication fork arrest and the different recovery mechanisms. (a) Telomeres present intrinsic obstacles, impeding the passage of replication machinery. Shelterin and accessory proteins prevent the fork stalling and repress improper recombination activity. Inappropriate recombination of the telomeric fork may result in telomere defects. (b) Persistent fork arrest might lead to fork collapse at telomeres. Restart of replication is necessary to avoid leaving an unreplicated region. Dormant origins could fire to complete telomere replication. Homologous recombination is also a general recovery mechanism from replication fork collapse. However, recombination at telomeres might increase the risk of cellular immortalization by ALT (alternative lengthening of telomeres). It is currently unknown how shelterin exactly contributes to restart of replication. BIR: break-induced replication; HDR: homology-directed repair; MTS: multi-telomeric signals; UFB: ultrafine anaphase bridge.
Genes 08 00112 g003

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Genes EISSN 2073-4425 Published by MDPI AG, Basel, Switzerland RSS E-Mail Table of Contents Alert
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