how telomeres are replicated

The ends of eukaryotic chromosomes, known as telomeres, are essential protein–DNA complexes that pro-tect chromosome ends from fusion and degradation1–3. In many organisms, the DNA component of telomeres comprises tandem repeats of guanosine-rich sequences in the strand that contains the 3′ end, which protrudes to form a single-strand overhang (the G-tail). Adjacent to the telomeric repeats, the subtelomeric region is com-posed of a mosaic of repetitive elements that exhibit a great deal of polymorphism. Telomeric proteins con-stitute specialized structures that have been extensively reviewed4,5. BOX 1 summarizes the composition of yeast, ciliate and mammalian telomeres, and BOX 2 describes the enzymatic activities as well as the DNA conforma-tions and transactions that are controlled by the human telomeric proteins.

In the absence of special telomere maintenance mechanisms, linear chromosomes shorten progressively with every round of DNA replication, eventually leading to cellular senescence or apoptosis (BOX 3). Consequently, non-dividing cells are capable of maintaining their telomeric DNA over time, even during a very long period, whereas telomere erosion is inexorably linked to cell division. A cellular reverse transcriptase called telomerase counteracts telomere shortening in many organisms6. It extends the 3′ end of chromosomes by reverse-transcribing in an iterative fashion the template region of its tightly associated telomerase RNA moiety, called telomerase RNA. In cells that express telomerase, telomeric DNA trimming still occurs but can be counterbalanced either partially or completely by the elongation of the G-rich strand and by its subsequent complementary replication.

Here, we first focus on the firing of the origins that are responsible for replication of telomeric sequences and on the difficulties that the replication fork encoun-ters in replicating telomeric chromatin. We then discuss

the possible ways by which the replisome proceeds at the end of chromosomal DNA and the current views on how the telomerase complex is assembled and activated at telomere ends to compensate for replicative erosion, and how these events are coupled to replication.

The quest for the last originThe replication of eukaryotic genomes initiates bidirec-tionally at defined origins. At chromosome termini, the last origin is expected to be responsible for the replica-tion of telomeric sequences. In budding and fission yeast, as well as in the holotrichous ciliate Tetrahymena thermophila, the origin that is used to initiate telomere replication lies on the centromere-proximal side of the terminal repeats7–11. Interestingly, there are several reports that show the binding of primase to telomeres and telomerase in yeast and ciliates, which suggests the existence of some initiation events that are triggered by telomere or telomerase12–14. Alternatively, specific inter-actions between primase and telomeres might help to coordinate synthesis of the G-strand by the telomerase, with the synthesis of its complementary strand by the lagging-strand replication machinery.

In budding yeast, there is compelling evidence that replication is not initiated within the telomeric repeats8. All subtelomeric regions in budding yeast contain at least one autonomously replicating sequence (ARS), which functions as a replication origin when present on an episome, but not necessarily when located in a chromosomal context. Indeed, every core-X and Y′ subtelomeric repeat (two classes of middle-repetitive sequences that immediately flank the telomeric DNA) contains an ARS, and all subtelomeric regions include one core-X element and 0–4 copies of Y′ elements. Most of the Y′ ARSs serve as replication origins10,15,16, whereas the core-X ARSs appear to be inactive or very inefficient at certain chromosome ends but not at others16.

*Laboratoire de Biologie Moléculaire et Cellulaire, UMR5239, IFR 128, Centre National de la Recherche Scientifique, University Lyon 1, Faculty of Medicine Lyon-Sud, Hospices Civils de Lyon, Ecole Normale Supérieure de Lyon, 46 Allée d’Italie, 69364 Lyon Cedex 07, France. ‡Laboratoire d’Instabilité Génétique et Cancérogenèse (IGC), Institut de Biologie Struturale et Microbiologie, CNRS, 31 chemin Joseph Aiguier, 13402 Marseille Cedex 20, France.e-mails: [email protected]; [email protected]:10.1038/nrm2259

Cellular senescenceA permanent form of cell-cycle arrest that can be induced by different types of exogenous or endogenous stress. Replicative senescence is triggered by an excessive telomere shortening that is the consequence of multiple rounds of cell division and is considered to be an intrinsic mechanism for limiting the proliferative lifespan of normal somatic cells.

Reverse transcriptaseAn enzyme that copies single-stranded RNA into single-stranded DNA.

How telomeres are replicatedEric Gilson* and Vincent Géli‡

Abstract | The replication of the ends of linear chromosomes, or telomeres, poses unique problems, which must be solved to maintain genome integrity and to allow cell division to occur. Here, we describe and compare the timing and specific mechanisms that are required to initiate, control and coordinate synthesis of the leading and lagging strands at telomeres in yeasts, ciliates and mammals. Overall, it emerges that telomere replication relies on a strong synergy between the conventional replication machinery, telomere protection systems, DNA-damage-response pathways and chromosomal organization.

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ReplisomeA multiprotein complex at the junction of the DNA replication fork that contains all the enzymes that are required for DNA replication.

In mammals, the lack of information concerning the direction of the fork movement at or near telomeres does not rule out the possibility that telomeres also serve as replication origins. Given that the TTAGGG-repeat factor-2 (TRF2), which binds the duplex part of telomeric DNA and is essential for telomere capping (BOX 1; BOX 2), has extra-telomeric roles in the initiation of replication at the oriP origin of the epstein–barr DNA

virus17, it could contribute to the formation of origins or origin-like elements within telomeres. by analogy to the oriP situation and in light of recent findings concerning the functions of TRF2, the activation of telomeric origins could involve a direct recruitment of the origin recognition complex (oRC) by TRF2 (Ref. 17) and a TRF2-mediated opening of the double helix18. Moreover, telomeric D-loops, the formation of which is stimulated by TRF2 and many other telomeric compo-nents18,19, might be used as a template for RNA priming and subsequent DNA extension. In conclusion, it is pos-sible that telomeres might be used as replication origins in certain organisms, although this has not yet been demonstrated.

Temporal regulation: is late the rule?Late timing in yeast. In yeast, the telomeres replicate in late S phase20. In a way that is reminiscent of the ability of telomeres to repress the transcription of a subtelomeric gene (BOX 1), telomeres exert a position effect on replication initiation, inactivating or delaying most of the origins that are present in the subtelomeric X and Y′ elements10,15,21,22. Mutants that lack the telomeric transcriptional silencing factor Sir3 show a premature activation of Y′ origins15, and tethering of Sir proteins near an origin can reset replication timing from early to late23. These results are in favour of a model in which the Sir-dependent silent chromatin that emanates from telomeres blocks replica-tion initiation in the subtelomeric regions. However, the telomeric position effect on replication timing extends over a distance (~35 kb) that is beyond the 6–8 kb esti-mated for the Sir-dependent gene repression24,25. This suggests that Sir-independent chromatin-mediated mechanisms can also contribute to the late activation of telomere-associated origins.

Cells that are defective for the multifunctional pro-tein Ku, which is involved both in repair and in telomere functions, show a partial delocalization of telomeres away from the nuclear periphery and an earlier firing of subtelomeric origins. This raises the possibility that the anchoring of telomeres to the nuclear envelop delays origin firing26. However, in cells that lack Ctf18, a protein involved in cohesin loading, telomeres are delocalized in G1 phase but replicate late as in wild-type cells27. This, together with the fact that the late replication pro-gramme is committed in G1 phase28, suggests that late telomere replication does not require peripheral posi-tioning. However, these observations do not rule out the possibility that other subnuclear positions could impose late timing on telomere replication. In contrast to most Y′-containing telomeres, the left telomere of chromo-some III replicates late because of its distance (40 kb) from an early activated origin and because of a reduced rate of replication fork movement in the leftmost region of this chromosome29.

overall, the data suggest that in the subtelomeric regions of budding yeast, transcriptional silencing, the rate of fork progression and late firing are the result of a peculiar organization of the chromatin at chromosome ends, involving both Sir-dependent and Sir-independent mechanisms.

Box 1 | The basic nucleoprotein organization of telomeres

In the ciliates Stylonychia and Oxytricha, the G-rich 3′ overhang is bound by a heterodimer that is formed by the oligonucleotide/oligosaccharide-binding fold (OB fold)-containing telomere end-binding protein-α (TEBPα) and TEBPβ (labelled as α and β in the figure)148. To our knowledge, proteins that bind to duplex telomeric DNA in ciliates have not yet been identified. In yeast, the telomeric DNA is packaged in a non-nucleosomal DNA–protein complex149. In budding yeast, duplex telomeric DNA is covered by an array of Rap1 proteins that interact either with a Sir complex, which is involved in the formation of subtelomeric heterochromatin, or with a Rif complex, which negatively controls telomere elongation150. The G-tail is bound by the OB-fold protein Cdc13, which forms a heterotrimer with Stn1 and Ten1, exhibiting structural similarities to the general single-stranded DNA-binding protein RPA (replication protein A)114. In fission yeast, Rap1 does not bind directly to telomeric DNA but is recruited to telomeres by interacting with the telomeric DNA-binding protein Taz1 (Ref. 150). The G-tail is coated by a TEBPα orthologue, Pot1. As in budding yeast, telomeres serve as nucleation sites for the heterochromatization of the subtelomeric chromatin (indicated by the binding of the HP1 orthologue, Swi6, to the subtelomeric nucleosome).

In mammals, most telomeric DNA is organized in tightly packed nucleosomes with a repeat size that is ~40 bp shorter than bulk nucleosomes. Some of these telomeric nucleosomes exhibit marks of heterochromatin, including HP1 (heterochromatin protein-1) binding151. Moreover, human telomeres exert a position effect on the expression of subtelomeric genes, indicating that, similar to yeast, human telomeres can initiate the formation of repressive subtelomeric chromatin152,153. In addition to nucleosomes, mammalian telomeric chromatin contains a set of telomeric DNA-binding proteins, including the Taz1 orthologues TTAGGG-repeat factor-1 (TRF1) and TRF2 (RefS 154–156), and the αβ orthologues POT1 (protection of telomeres protein-1)–TPP1 (POT1 binding partner; formerly named TINT1/PTOP/PIP1)144,145. These proteins are involved in a network of homo- and heterotypic interactions with other proteins such as TIN2 (TRF1-interacting factor-2) and RAP1 (Ref. 4). These different proteins, as well as the telomeric nucleosomes, are arbitrarily placed along the telomeric DNA, with the exception of POT1–TPP1, which interacts with the G-tail. The thick red lines represent duplex DNA telomeric repeats, whereas the thin red lines represent G-tails. The length of the telomeric DNA is not drawn to scale and varies from tens of nucleotides for ciliate telomeric DNA in the macronuclei to hundreds for yeast and thousands for mammals.

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PrimaseThe enzyme that synthesizes an RNA primer for initiation of DNA replication. Primase is associated with DNA polymerase-α to form a four-subunit complex. The polymerase-α–primase complex functions in the initiation of DNA replication at chromosomal origins and in the discontinuous synthesis of Okazaki fragments on the lagging strand of the replication fork.

OB foldAn N-terminal oligonucleotide/oligosaccharide binding (OB) motif. The five-stranded β-sheet forms a closed β-barrel, which is capped by an α-helix located between the third and fourth strands. The OB fold is frequently used for the specific recognition of single-stranded nucleic acids.

Origin recognition complexA heteromeric six-subunit protein complex that binds to DNA at replication origin sites and functions as a scaffold for the assembly of pre-replicative complexes in the G1 phase of the cell cycle.

D-loopThe displacement loop structure that results from the displacement of a duplex DNA by a homologous single-stranded DNA.

Position effectThe influence of the chromosomal context on various DNA transactions, including transcription, replication and recombination. It often refers to the repression that is conferred by heterochromatin proximity.

Sir proteinsThe silent information regulators (Sir)-2, -3 and -4 are the structural constituents of a particular type of silent chromatin in budding yeast. At telomeres, Sir3 and Sir4 interact with the telomere-binding protein Rap1, can self-associate, and bind to deacetylated and demethylated N-terminal tails of histones H3 and H4 of subtelomeric nucleosomes. The deacetylase activity of Sir2 is required to spread the Sir complex along the chromatin toward the centromere.

Early–late timing of mammalian telomeres? The replica-tion timing of telomeres seems to differ between differ-ent species. In the Indian muntjac deer, telomeres of individual chromosomes have a characteristic timing of replication throughout S phase30. Interestingly, the tim-ing between the telomeres of homologous chromosomes is highly coordinated and no such synchrony is observed between the two telomeres of the same chromosome30.

Telomeric DNA was shown to replicate throughout S phase in a wide variety of human cell types31–33. In apparent contrast, the detection of several subtelomeric

regions of the human genome as a duplicated signal (doublet) by fluorescent in situ hybridization (FISH) occurs late during S phase34, which suggests that sub-telomeric DNA is late replicating. To reconcile these data with the early replication of bulk telomeric DNA, one possibility is that replication origins are present within the telomeric repeats, leading to an earlier replication of the telomeric DNA than of the subtelomeric regions (see above). Another, non-exclusive explanation is that the late appearance of the subtelomeric duplication results from a block during replication and/or delayed

Box 2 | Enzymatic activities and DNA conformations controlled by human telomeric proteins

The telomeric proteins TTAGGG-repeat factor-1 (TRF1), TRF2 and protection of telomeres protein-1 (POT1) modulate diverse enzymatic activities and DNA conformations that control different aspects of telomere homeostasis (see figure)5. The black arrows indicate an activation effect and the red blunted arrows indicate an inhibitory effect. The dotted lines signify that the telomeric protein has an interaction with the corresponding enzyme or DNA conformation, but that it is not known whether this results in activation or inhibition. The grey flash represents a nuclease activity. The orange strand represents the RNA component of telomerase.

In the main text, we discuss models that allow the integration of these different activities in telomere replication. For example, TRF2 interacts with different enzymes that could control G-tail formation (the nuclease XPF1–ERCC1, the MRE11–RAD50–NBS1 (MRN) complex, the RecQ helicase WRN and the 5′ exonuclease Apollo). Fine-tuning of these activities could at least partially account for events that follow either TRF2 inhibition (that is, end-to-end fusion) or TRF2 overexpression, such as rapid telomere loss. TRF1 and TRF2 appear to control the locking and opening of telomeric loops (t-loops) by means of various DNA chaperone activities and enzyme recruitments. For instance, TRF1 is able to bring portions of telomeric DNA into close proximity (parallel pairing), while TRF2 promotes double-strand invasion by the 3′ overhang (strand invasion), introduces topological stress (topology), binds to four-way Holliday junctions and activates D-loop removal by the RecQ helicases WRN and BLM18,89,90,157.

POT1 can assist TRF2 in t-loop clearance by favouring the D-loop removal by WRN and BLM158. POT1 and the helicases WRN and BLM are also able to unfold G-quadruplex (G4) DNA159. TRF1 interacts with various proteins that modulate telomerase activity, such as POT1. These interactions could account for telomere-length shortening following TRF1 overexpression. TRF1 can also act on double-stranded telomeric DNA association160, thus suggesting a role for TRF1 in t-loop formation and in telomere–telomere association in interphase nuclei. TRF2 has been shown to be physically bound to DNA polymerase β (POLβ) and to flap endonuclease-1 (FEN1). In vitro, TRF2 stimulates POLβ activity in reconstituted base-excision repair reactions169.

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Box 3 | Chromosome end-protection

Telomeres protect chromosome ends from being recognized and processed as DNA double-strand breaks and, therefore, from triggering DNA-damage-induced responses such as checkpoint activation through the ataxia telangiectasia mutated (ATM; also known as Tel1 in yeast) and ATM and Rad3-related (ATR) pathways or recombinational repair, including non-homologous end joining and homologous recombination (see figure part a). This process of telomere end-protection relies on telomere-specific DNA conformations and chromatin organization as well as double-stranded DNA telomere-associated proteins (green and blue circles) and single-stranded DNA telomere-associated proteins (purple circles) (BOX 1).

Telomeres can fold into t-loops that may result from invasion of the 3′ overhang into duplex DNA or into G-quadruplex DNA, an unusual DNA conformation that is based on a guanine quartet (‘G quartet’) (see figure part b). However, whether t-loops or G quadruplexes can provide end-protection is still unknown. The telomeric DNA is organized into a set of unusual chromatin structures (BOX 1) that provide several types of activities that could prevent DNA-repair activities (end-joining, XPF–ERCC1 nuclease; see BOX 2). These protective complexes would need to be resolved to allow DNA replication and telomere elongation by telomerase. Paradoxically, many DNA-damage-checkpoint proteins also bind to telomeres and are transiently activated during normal replication19. However, fully functional telomeres do not elicit a DNA-damage response that would be sufficient to stop cell proliferation. Senescence is triggered by either excessive telomere shortening or disruptions in the function of protective complexes. Importantly, short telomeres that trigger senescence do not appear to be completely unprotected because they are still sheltered from non-homologous end joining and homologous recombination. By contrast, the disrupted telomeres appear to be fully unprotected because they both activate the checkpoint and lead to aberrant telomere recombination events161.

t-loopA structure adopted by telomeres that may result from invasion of the 3′ overhang into duplex DNA.

G quadruplexA four-stranded structure that is held together by square planes of four guanines (‘G-quartets’), associated through Hoogsteen base pairing. Once such structures form they are extremely stable and are likely to need enzymatic activity to be unwound in vivo.

chromatid separation of the telomeric region. To resolve this question definitively, there is a vital need to measure the spatio-temporal dynamics of replication forks within telomeric repeats and immediate subtelomeric regions and to compare, for given chromosome ends, the tim-ing of replication between telomeric and subtelomeric sequences.

by combining bromodeoxyuridine (brdu) incor-poration assays with chromatin immunoprecipitations (ChIPs) of telomeric proteins in cells that were devoid of telomerase activity, verdun and Karlseder19 showed that replication of telomeres occurs in two phases, one in S phase and the other in G2–M phase. Although it cannot be excluded that the delayed incorporation results from a subset of very late telomeres, this study raises the interest-ing possibility that the completion of telomere replication is deferred to the very late stages of S phase or even in G2 phase, explaining the late appearance of the subtelomeric doublets. In this model, the late timing of the very end of human chromosomes is reminiscent of the late timing of telomeric DNA in yeast.

Possible roles of telomere late replication. The late rep-lication of the very end of telomeric regions might be important to coordinate telomere functions with cell-cycle-regulated events. For example, in budding yeast, short telomeres are preferentially elongated by telo-merase35,36 and replicate earlier than long telomeres37, suggesting that late replication impairs telomerase action. However, the links between late replication and telomere-length regulation are likely to be more complex — the kinase Rad53 (the orthologue of mam-malian CHK2 (checkpoint kinase-2)) slightly delays the firing of the subtelomeric origins38–40 and causes telomere lengthening41. In addition, formation of the G-tail, which is one of the terminal events of telomere replication, requires the activity of the cyclin-dependent kinase Cdk1 (also known as Cdc28 in budding yeast) (see below). late replication might also be required to re-establish a specific chromatin domain after DNA replication and to allow sister-chromatid cohesion.

Replication difficulties at the telomereTelomeric chromatin has the ability to form various unusual structures, which might be a source of dif-ficulties for the passage of the replication fork. Some of them have been detected in vivo, including a hetero-chromatin-like structure, G quadruplexes and t-loops. Some others, which are more hypothetical, are inferred from in vitro studies and include triple helices, four-way junctions and D-loops. In accordance with this prediction, a wealth of recent results suggests that replication forks can pause or stall naturally at telo-meres in yeast9–11 and in human primary fibroblasts19. In agreement with these replication problems, an ATM/ATR (ataxia telangiectasia mutated/ATM and Rad3-related)-dependent DNA-damage response occurs that is mediated by the MRe11–RAD50–NbS1 complex, which is involved in the initial recognition of double-strand break DNA, and the single-stranded DNA-binding protein RPA (replication protein A) appears transiently at telomeres during S phase of human cells19. The unreplicated DNA distal to a stalled fork at most genomic loci can eventually be replicated by a fork that approaches from the opposite direction. However, telomeres lack such oncoming forks and, therefore, a stalled telomeric fork cannot be rescued by a more distal one, strengthening the need for robust systems to remediate fork progression difficulties at telomeres.

How can the fork progression problems caused by the conformation of telomeric chromatin be solved? In fission yeast, the telomere-binding protein Taz1 has recently been shown to be crucial for efficient replica-tion of telomeres or internally positioned telomere sequences11. In a taz1 mutant, the replication fork pauses at telomeres and leads to an entanglement, which causes abrupt telomere loss in a telomerase-negative back-ground11. From these results, it has been extrapolated that telomere-binding proteins have an essential role in coordinating replication fork progression through telomeres by preventing stalled forks, which may result in telomere attrition or increased recombination.

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RecQ helicaseOne of a family of evolutionarily conserved helicases, mutations of which can lead to hereditary cancer-predisposition syndromes in humans. Helicases use the energy of ATP hydrolysis to unwind duplex DNA.

DNA topoisomeraseAn enzyme that changes DNA supercoiling by inserting or removing superhelical twists.

Increasing evidence indicates that RecQ helicases also have an important role during the replication of mammalian telomeres. The RecQ helicase WRN local-izes to telomeres in S phase and is required for efficient lagging-strand replication of the G-rich telomeric DNA, preventing a dramatic loss of telomeric DNA42. Moreover, expression of a WRN protein that contains a dominant-negative mutation increases the rate of telomere loss and chromosome fusion43. These results suggest that WRN helicase activity is principally involved in removing the secondary structure (for example, a G quadruplex) that can be formed in the G-rich strand42,44. blM, another RecQ helicase, appears to perform overlapping func-tions with WRN. For instance, mutations in Wrn and Blm each accentuate the pathology in later-generation mice that lack the telomerase RNA template45,46. The telomere-binding proteins TRF2 and PoT1 (protection of telomeres protein-1) are obvious candidates for the fine regulation of WRN and blM recruitment and activity (BOX 2; fIG. 1), although their recruitment to telomeres could also be mediated by components of the replication machinery such as DNA polymerase-δ or RPA44.

The presence of a t-loop is expected to represent a potent obstacle for fork progression. Moreover, the basis of a t-loop, which is formed by nucleoprotein structures organized around a D-loop and possibly a four-way junc-tion, is unlikely to be free to rotate and, therefore, can be considered as a topological barrier. Hence, when the fork approaches a t-loop, an accumulation of superhelical stress in the unreplicated DNA might hinder the actions of topoisomerases, increasing the local amount of posi-tive supercoiling ahead. Amazingly, TRF2, but not TRF1

and nucleosomes, induces positive supercoiling, which suggests that its binding to positive supercoils would be energetically favoured18. Hence, TRF2 might be highly enriched around the fork, serving as a topological stress sensor. Subsequently, the increased concentration of TRF2 might favour t-loop opening by activating the RecQ helicases and possibly other activities (BOX 2; fIG. 2). How TRF2 could promote both t-loop formation and reso-lution would possibly rely on the formation of different TRF2 subcomplexes during the cell cycle. Remarkably, in the lyme disease bacterium Borrelia burgdorferi, posi-tive supercoiling stimulates the activity of the telomere resolvase ResT by promoting its binding to DNA47. Therefore, using the free energy of positive supercoiling generated by the replication of topologically constrained domains might be a general mechanism that is exploited by telomere proteins to facilitate telomere replication.

In budding yeast, the telomeric replication fork starts to pause ~100 bp upstream of the telomeric TG1–3 repeats in a way that appears to be independent of the Sir and Rif proteins, which represent the main known compo-nents of yeast telomeric chromatin10. The slowing of the replication fork is greatly exacerbated in the absence of the Pif1-related Rrm3 helicase, which has been shown to facilitate replication through nonhistone protein–DNA complexes9,10,48. Recently, Rrm3 was shown to move with the replication fork and to be a component of the replication fork apparatus49. extra Rrm3 molecules may be recruited to particularly stable protein–DNA com-plexes such as the telomeres, or Rrm3 could be part of a chromatin remodelling complex that moves immediately ahead of the replication fork49. It is noteworthy that yeast

Figure 1 | model for fork progression through chromosome ends in mammalian cells. Heterochromatin, hairpins, G quadruplexes (G4), triple helices, four-way junctions, D-loops and t-loops may hamper the progression of the replication fork through the telomeric chromatin. This model describes the role of the various activities that are involved in removing the secondary structure that can be formed during telomere replication, particularly at the G-rich lagging strand (see main text). WRN and BLM helicases are proposed to dissociate unusual DNA structures during replication at telomeric ends and to unlock the t-loop by D-loop unwinding (see arrows 1 and 2). TTAGGG-repeat factor-2 (TRF2) and protection of telomeres protein-1 (POT1) are thought to stimulate these activities of WRN and BLM (see arrows 3 and 4)158. RPA (replication protein A) and POT1 may also bind and unfold telomeric G-quadruplex structures (see arrow 5)159,168. The effect of the 5′→3′ exonuclease, 5′ flap endonuclease FEN1 on Okazaki fragment processing and maturation appears to be stimulated by TRF2 (see question mark). From yeast findings, one can infer that the RRM3/PIF1-related helicase (see arrow 6) facilitates the progression of the replication fork through unusual nucleoprotein complexes. This helicase might travel with the replication fork (see main text). HP1, heterochromatin protein-1; PCNA, proliferating cell nuclear antigen; POLδ/ε, DNA polymerase-δ/ε; TIN2, TRF1-interacting factor-2; TPP1, POT1 binding partner.

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cells that lack Sgs1 (the orthologue of WRN) and Rrm3 accumulate gross chromosomal rearrangements that are suppressed by homologous recombination (HR)-defective mutations. This suggests that Sgs1 and Rrm3 may act in parallel to prevent illicit homologous recombination, particularly during DNA replication of telomeres50.

In summary, telomere-binding proteins recruit RecQ helicases and possibly other co-factors that are capable of removing or remodelling the telomeric structures that can impair fork progression. In agreement with this idea, these helicases can remove D-loops and unfold G-quadruplex structures in vitro51 (BOX 2).

The end point of telomere replicationHere, we examine the situation that has been most thor-oughly documented; that is, when telomere replication is initiated from a non-telomeric origin and not from a telomeric one (see above). In this case, leading-strand

synthesis produces the G-rich daughter strand and the lagging-strand synthesis produces the C-rich daughter strand. In many, if not all, organisms, a G-tail can be detected at both daughter telomeres52–56. Its formation requires the passage of the replication fork, at least in budding yeast57.

5′ resection is involved in G‑tail formation. because the length of the G-tail is not affected by the presence of telomerase53,58, it must be hypothesized that the G-tail results from the processing of the 5′ strand rather than from elongation of the 3′ strand (fIG. 3). For the leading telomere59, the end product is either blunt or 5′ pro-truding, which implies the existence of a 5′ resection activity to convert it into a 3′ overhang. Whether a 5′ resection activity also processes the lagging telomere is still unknown (see below). In budding yeast, the G-tail formation requires Cdk1, as is the case for accidental

Figure 2 | Topology and t-loop problems might be coupled during fork progression. Replication transiently generates positive supercoiling ahead of the elongating fork, which is usually rapidly relaxed by topoisomerases. However, the lock at the basis of the t-loop, which is thought to be formed by a complex nucleoprotein architecture involving a D-loop and a four-way junction, is unlikely to be free to rotate and can be considered as a topological barrier. Therefore, when the fork approaches a t-loop, one expects an accumulation of positive supercoiling in the unreplicated DNA in front of the t-loop and, ultimately, fork pause or arrest. This blockade could be rapidly relieved by t-loop opening and progression of the fork towards the very end of the chromosome (central part of the figure). It could also be efficiently released by topoisomerases and t-loop-opening activities (right part of the figure). Alternatively, if the action of topoisomerases is uncompleted in the presence of a t-loop, a residual positive supercoiling might favour fork regression and the formation of a four-way junction, called a chicken foot (left part of the figure). Chicken-foot regression or resolution, together with t-loop opening, will rescue the blocked fork. It is noteworthy that TTAGGG-repeat factor-2 (TRF2) specifically binds several of these DNA structures, perhaps as part of different shelterin subcomplexes. This might reverse the chicken foot and t-loop through TRF2-dependent activation of processes that are known to disrupt these structures (see BOX 2 and main text). Positive supercoils are also expected to represent a favoured substrate for TRF2 binding (see main text), possibly leading to a high concentration of TRF2 ahead of the fork, which might promote t-loop opening.

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double-strand breaks (DSbs)60–62. In humans, on com-pletion of telomere replication, an ATM-dependent damage signal is required for generation of the G-tail19,62.

Collectively, these findings reveal important similari-ties between the normal processing of a chromosome end and the repair of an accidental DSb60. Therefore, it is not surprising that G-tail formation involves the Mre11 complex, which consists of Mre11, Rad50 and Xrs2 in yeast. In Mre11-defective yeast strains, the length of the G-tail is reduced55. Furthermore, long telo-meric tracts inhibit the binding of Mre11 and prevent 5′ resection63. However, G-tails are still present in cells that express nuclease-dead alleles of Mre11, suggesting that the nuclease activities of Mre11 are not strictly neces-sary for telomeric 5′ resection55,64. Therefore, the role of the Mre11 complex in the 5′ resection of telomeric ends might be more structural than catalytic. In human cells, the Apollo 5′ exonuclease is associated with TRF2 and is required to protect telomeres during S phase65,66. Thus, Apollo is a potential candidate to contribute to G-tail formation through its 5′ resection activity. overall, the nature of the nuclease(s) involved in the 5′ resection activity of telomeric ends remains largely unknown.

Different processing mechanisms act at telomeres. If involvement of a 5′ resection activity is needed to explain the formation of the G-tail at the leading telo-mere (see above and fIG. 3), an incomplete okazaki fragment synthesis can, a priori, be sufficient to gen-erate the lagging G-tail. This is consistent with the telomerase-independent formation of long 3′ overhangs in cells carrying mutations that affect several genes involved in okazaki synthesis67–69. one possibility is that lagging-strand synthesis stops once the leading-strand synthesis has finished replication of the parental C-rich strand (fIG. 3). If we estimate that the replication lag between the two strands corresponds to the time necessary to synthesize an okazaki fragment (that is, ~200 nucleotides (nt) in eukaryotic species70), one would expect that the lagging-strand synthesis pauses at ~200 nt from the last nucleotide of the parental G-rich strand. In agreement with this expectation, in vitro reconstitution of linear DNA replication reveals that lagging-strand synthesis is gradually compro-mised in the terminal region, leaving a 3′ overhang of ~250 nt59. The length of this unreplicated region is com-patible with the length of the G-tail observed in human cells, although additional processing or synthesis events could also take place52,56.

In organisms that exhibit a much shorter G-tail, as in yeast, the terminal gap could be partially filled in by telomere-specific priming events. Accordingly, the yeast Cdc13–Stn1–Ten1 complex interacts with Pol1 and Pol12 (two subunits of DNA polymerase-α)71,72, and muta-tions in Pol1 (the catalytic subunit) result in longer 3′ overhangs, even in the absence of telomerase68,73. A delayed priming of the lagging strand is consistent with the transient appearance of longer G-tails at the end of S phase in yeast74 and with the late incorpora-tion of brdu at human telomeres19. Therefore, the

Figure 3 | models for g-tail formation. The diagram shows the molecular events that are expected to cause the formation of a G-tail at both daughter telomeres after each round of replication. There is mounting evidence that the leading and lagging G-tails do not have the same size and are therefore expected to be processed differently. The G-rich strand is replicated by the lagging-strand machinery, whereas the C-rich strand is replicated by the leading-strand machinery. Consequently, the parental G-rich strand is not completely duplicated, leading to a G-rich 3′ overhang. The length of this G-tail can be different if it corresponds to the length of the last primer (that is, roughly 10–12 nucleotides) or to an inhibition in the synthesis of the last Okazaki fragment, leading to a longer overhang of ~200 nucleotides (pathway on the left). An alternative mechanism is that the inappropriate synthesis of the last Okazaki fragment is compensated for by a telomere-specific recruitment of a primase complex (shown in the right pathway). Importantly, in the left pathway, there is no need to implicate a 5′ resection activity to generate a long G-tail for the lagging strand. In both pathways, there must be a specific mechanism to remove the last primer because there is no upstream Okazaki fragment to displace the primer by the 5′→3′ exonuclease, 5′ flap endonuclease FEN1/Rad27 and DNA2. This might involve an RNase H activity. For the leading strand, which must terminate as either a blunt end or as a 5′ overhang, one has to assume the existence of a 5′ resection activity to generate a G-tail. This activity involves the MRE11–RAD50–NBS1 complex, although the nature of the nuclease is still elusive. The extent of 5′ resection must be controlled by telomere factors, the mode of action of which remains unknown. The processing appears to terminate with the action of C-rich and G-rich sequence-specific nucleases, which will determine the last nucleotide.

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transient lengthening of the G-tail observed at late S phase in yeast could result from the combined action of an Mre11-dependent resection at the leading telomere and of a delayed synthesis of the last okazaki fragment at the lagging telomere (fIG. 3). This model is consistent with the fact that Mre11-compromised yeast cells still show a transient, longer G-tail in S phase55.

G‑tail length regulation. In yeast, the extent of the 5′ resection is negatively regulated by several telomeric proteins, including Cdc13 in S phase62, Ku75 and Rap1 (Ref. 63). In fission yeast and vertebrates, PoT1 controls G-tail length4,76,77. How the components of telomeric chromatin control the various activities that are required for proper processing of telomeres remains largely unknown. The fact that the XPF–eRCC1 nuclease is part of TRF2 complexes and is involved in the removal of the G-tail after TRF2 inhibition suggests that TRF2 sequesters XPF–eRCC1 in an inactive enzyme form78. However, removal of the G-tail on TRF2 loss also depends upon DNA ligase-4 (lIG4), suggesting that clipping of the overhang by XPF–eRCC1 is part of the non-homologous end joining (NHeJ) process that leads to the fusions of unprotected telomeres79. Therefore, XPF–eRCC1 is not expected to greatly contribute to G-tail-length regulation at protected telomeres.

Removal of the RNA primers. Another telomere-specific lagging-strand problem is removal of the last RNA primer. During normal replication, this reaction depends on the downstream okazaki fragment, which displaces the primer into a flap that is cleaved by the concerted action of the helicase DNA2 and flap endonuclease-1 (FeN1) in mammals (Rad27 in budding yeast) (reviewed in Ref. 80). However, a downstream okazaki fragment cannot exist at the very end of the lagging daughter strand to displace the strand and recruit FeN1, suggesting that a special telomere-specific event is required to remove the last RNA primer. For example, Pif1 helicase, a negative regulator of telomerase in bud-ding yeast, has been proposed to aid Dna2 for removal of RNA from the okazaki fragments at the telomere81. Alternatively, RNase H activity might have an important role in processing the last primer. In agreement with this possibility, in yeast, RNase H2 can aid the removal of RNA primers during lagging-strand synthesis82 and interacts with Rif2, a negative regulator of telomere elongation83. Moreover, in an in vitro assay of mam-malian telomere lagging-strand synthesis, the removal of the RNA primer was observed upon the addition of purified RNase H84.

Synthesizing the last nucleotides. Finally, at least in cili-ates and humans, sequence-specific events are involved in the precise determination of the last nucleotides of the C-rich and G-rich strands85–87. In PoT1-deficient human cells, the sequences of the 5′ extremities appear to be random, which suggests that the PoT1 protein, bound to the G-tail, modulates the activity of a nucle-ase that is responsible for determining the 5′ end88. The nature of this nuclease is still mysterious.

Higher‑order telomere assembly. once formed, the daughter telomeres fold into higher-order nucleoprotein complexes that are involved in telomere protection and telomerase regulation. The G-tail can invade the duplex part of the telomere in cis, forming a t-loop. This strand-invasion reaction can be assisted in vitro by the synergistic effect of several telomere proteins and factors that are involved in homologous recombination18,19,89. Interestingly, the binding of TRF2 to a Holliday junc-tion could stabilize the basis of the loop by locking the Holliday junction formed from the invaded G-tail90 (A. Poulet, e.G. and M.J. Giraud-Panis, unpublished observations). It is worth noting that in human cells, the leading G-tail is shorter than the lagging one56, raising the possibility that t-loops are more difficult to form at the leading than at the lagging telomeres. This would explain the fact that leading telomeres are more easily uncapped when TRF2 and DNA-PKcs (DNA-dependent protein kinase catalytic subunit) are dysfunctional, and explains why they are more prone to be elongated by telomerase91,92.

In budding yeast, telomeres are clustered into a lim-ited number of foci that are primarily associated with the nuclear envelope93. The fact that telomere movements are constrained both during interphase and S phase94 suggests the existence of telomere-specific replication factories that bring together several replicating telomeres near the nuclear periphery. This clustering might facilitate the re-assembly of a proper higher-order organization of telomeres after their replication.

Compensating for replicative erosionThe erosion of telomeric DNA that results from its replication can be compensated for by various mecha-nisms involving recombination, retrotransposition and telomerase-dependent repeat addition (BOX 4). Here, we discuss the current views on how telomerase is recruited and activated at telomeres and how these events are coupled to replication.

Telomerase recruitment in budding yeast. In budding yeast, telomere elongation by telomerase coincides with telomere replication in cells that progress synchronously through the cell cycle95. Interestingly, the replication of a telomere appears to facilitate its extension by telomerase, suggesting a coupling between in vivo telomerase activity and conventional DNA replication. In agreement with this assumption, elongation of telomeric DNA repeats by telomerase at a DSb that is flanked by a telomeric DNA tract (telomeric DSb) requires DNA polymerase-α and -δ function, as well as primase activity96.

In budding yeast, the timing of telomere elonga-tion correlates with the binding at telomeres of several proteins that are involved in telomere elongation, including the telomerase holoenzyme proteins est1, est2 and est3, Cdc13 (and its associated proteins Stn1 and Ten1), Mre11 (and its associated proteins Rad50 and Xrs2) and RPA64,97–100 (fIG. 4). It is noteworthy that Mre11 binds to telomeres just before Cdc13, est1, est2 and RPA (and probably est3), which are all recruited at the same time64. The inactivation of Mre11 appears

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to strongly affect the binding to telomeres of est1 and est2 in late S phase and to partially affect the binding of Cdc13 (RefS 64,101). Whereas the binding of est1, Cdc13, Mre11 and RPA is low or undetectable in G1 phase, the telomerase (the catalytic subunit est2 and the RNA template TlC1) is bound to capped telomeres in the G1 phase through an interaction with the Ku heterodimer (Ku70–Ku80) via the Ku-binding arm of the RNA template TlC1 (RefS 102,103). In early S phase, est2 is removed by an unknown mechanism but binds again in late S phase. This binding, in turn, is followed by a decreased association of telomerase to telomeres in G2–M phase. Therefore, the catalytic subunit of telo-merase, est2, exhibits a biphasic cell-cycle-regulated telomere-binding profile97, which is not fully elucidated (fIG. 4) (see below).

The current model is that the interaction of Cdc13 with est1 mediates the recruitment of telomerase in late S phase to chromosome ends99,104. Consistent with this view, a strain that carries an allele of cdc13 (cdc13‑2) that can no longer interact with est1 and abolishes telomerase activity in vivo105,106 exhibits a defect in the recruitment of est1 and est2 at both telomeric DSbs as well as at telomeres in late S phase97,99. Reciprocally, a non-func-tional allele of est1 (est1‑60) that is unable to interact with Cdc13, but which re-establishes the interaction with the telomerase null allele cdc13‑2, is a suppressor of cdc13‑2 (RefS 99,106).

However, the way in which telomerase is recruited to telomeres is likely to involve a more complex network of interactions. For instance, an allele of RFA2 (the gene encoding the middle subunit of RPA) exhibits a severe reduction of telomerase activity in vivo and greatly reduces the binding of est1 to telomeres, but does not

affect the binding of est2 in late S phase98. Therefore, it seems that in late S phase, telomerase activity involves Cdc13, est1, est2, est3, RPA, TlC1 and probably Ku, because telomere-elongating cdc13 alleles have been identified and shown to require Ku for their effect107.

It has recently been observed that RPA interacts with the telomerase RNA TlC1 in late S phase when RPA, Cdc13, est1 and est2 are recruited to telomeres (P. luciano, e.G. and v.G., unpublished observations). A secondary structure model of TlC1 RNA and protein-binding experiments indicate that TlC1 is organized as a central core that contains the RNA template and the est2-binding site, from which three RNA arms emanate and interact with est1, the Ku heterodimer and the Sm heteroheptamer (which is involved in ribonucleoprotein maturation), respectively108–113. These results reveal that TlC1 serves as a flexible scaffold for proteins to assemble into the telomerase holoenzyme (fIG. 4). overall, it emerges that Cdc13, which has been proposed to be a telomere-specific RPA-like complex114, est1, Ku and RPA may cooperate to activate a functional telomerase complex at chromosome ends during late S phase. We speculate that est1, Ku and RPA may act as chaperones to maintain TlC1 in a functional conformation.

Recent results in human cells show that telomerase may extend overhangs differently or affect telomere end-processing differently on the leading versus the lagging telomeres56. It is therefore possible that the mechanism by which the telomerase complex is recruited to telomeres at the end of S phase is different for the lagging telomere and the leading telomere. Therefore, in yeast, Cdc13 and RPA could have different roles in telomerase recruitment at the leading and lagging telomeres. besides its role in telomerase recruitment, the heterotrimer Cdc13–Stn1–Ten1 interacts with the polymerase-α complex to coordi-nate the synthesis of the complementary C strand, which exerts a negative effect on telomerase action71,106,115.

one of the last events of the ‘telomerase cycle’ is the removal of telomerase from the telomeres. Telomere addition is impaired by Pif1 helicase, the role of which is to limit telomerase processivity by dissociating the telomerase-RNA–telomeric-DNA hybrid that is formed during telomere replication116. Recent results indicate that the reverse transcriptase finger subdomain of telomerase cooperates with Pif1 to limit telomerase asso-ciation with chromosome ends and, consequently, limits telomere elongation117.

Checkpoint activation and telomerase action. Telomeric DNA-binding proteins regulate telomere length through a negative feedback mechanism118. The cis-repression exerted by the telomeric DNA-tract-binding proteins is thought to affect the probability that telomerase can access and elongate the telomere during one genera-tion36. As a consequence, short telomeres have a higher probability of being elongated by the telomerase than the longer ones35. Several genetic studies indicate that, in yeast, Tel1 and Mec1 (the orthologues of the DNA-damage-checkpoint kinases ATM and ATR, respec-tively) have a role in allowing telomerase access to the telomere119,120. However, whereas Tel1 acts as a major

Box 4 | Mechanisms that compensate for telomere loss

Because the length of telomeric DNA inexorably shortens during replication, there is a need for compensating mechanisms to preserve genome integrity and telomere functions. In many organisms, a specialized reverse transcriptase named telomerase catalyses the addition of short and simple repeats in a process that is tightly coupled to replication6 (see main text). However, the use of telomerase is not universal and the compensating mechanisms are far more diverse than originally thought.

In the absence of telomerase, telomeric DNA loss can also be compensated for by alternative lengthening of telomeres (ALT) mechanisms162. Based on differences in telomere structure, one can distinguish between the mechanisms that amplify subtelomeric repeats and the mechanisms that lengthen the simple telomeric repeats alone163. The ALT mechanisms are still unclear and appear to rely on homologous recombination, rolling-circle replication, extrachromosomal circle integration and break-induced replication164. The ALT pathways can be considered as backups of telomere maintenance in organisms that normally exploit the telomerase system. For instance, whereas ALT is inhibited in normal human cells, some human tumours maintain their telomeres using ALT. Whether some organisms or cell types normally rely on ALT for telomere-length maintenance is still unclear. In certain organisms, including dipteran insects, telomerase and short repeats have never been detected. Whereas the telomeres in Drosophila melanogaster are formed by long arrays of the non-long-terminal-repeat (non-LTR) retrotransposons HeT-A, TART and TAHRE, chironomids (mosquitoes) terminate their chromosomes by using tandem repeats that are in the range of hundreds of nucleotides165,166. Interestingly, dysfunctional telomeres in mammals can lead to the retrotransposition of long interspersed element-1 (LINE-1 or L1) elements167. This provides further intriguing evolutionary links between retrotransposition and mechanisms for telomere loss compensation.

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Figure 4 | Dynamics of telomerase recruitment and activation through the cell cycle in budding yeast. In this model, telomerase (comprising the catalytic subunit Est2 and the RNA template TLC1) is bound to capped telomeres in G1 phase through an interaction of TLC1 with the Ku heterodimer. In early S phase, telomerase is removed from the telomere, whereas Ku remains associated with chromosome ends. In late S phase, the replication fork passage leads to the transient disruption of telomeric chromatin. For the telomere that derives from the replication of the leading strand (leading telomere), unidentified exonuclease activities (Exo) contribute to the formation of G-rich overhangs through a telomeric 5′ resection that is controlled by Tel1 (the orthologue of the DNA-damage-checkpoint kinase ATM), the Mre11–Rad50–Xrs2 complex (MRX) and the cyclin-dependent kinase Cdk1. The generation of G-tails creates binding sites for Cdc13, the extent of the resection being controlled by the heterotrimer Cdc13–Stn1–Ten1 (CST) and Rap1. We speculate that the lagging G-tail, which results from an impaired Okazaki synthesis, is covered by replication protein A (RPA) and Cdc13. Different mechanisms may be used to recruit telomerase at the leading and the lagging telomere. Through a yet-unknown mechanism, telomerase (Est2 and TLC1) becomes engaged in an interaction involving Cdc13, Est1, Est3, RPA and probably Ku that activates telomerase. Recent results indicate that Tel1 and telomerase bind preferentially to short telomeres, which suggests that Tel1 favours their elongation by telomerase. Whereas Cdc13 binds to the newly generated G-tails, the DNA polymerase-α–Pol12–primase complex, which interacts with the CST heterotrimer, synthesizes the complementary C-rich strand and limits the activity of telomerase. As telomeres are replicated, Rap1 binds to newly synthesized telomeres, thereby inhibiting over-elongation of telomeres by telomerase. The removal of telomerase from telomeres in S–G2 phase is actively promoted by the helicase Pif1. Most of the components of the telomerase holoenzyme are removed from the telomere in G2–M phase. Rif1/2, Rap1-interacting factor-1/2.

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positive regulator of telomerase121,122, Mec1 appears to have a modest role in telomere-length regulation41,123. Tel1, but not Mec1, was shown to be required for normal levels of telomere association of est1 and est2 (but not of Cdc13) in late S phase101.

Recently, four different laboratories have uncovered evidence that Tel1 and telomerase bind preferentially to short telomeres in a Xrs2-dependent way, suggesting that Tel1 favours their elongation by telomerase124–128. Taken together, this evidence suggests that short telo-meres, in contrast to the long ones, may induce a trans-ient cell-cycle-regulated DNA-damage-checkpoint response that is required for telomerase binding. Finally, telomerase activity is sensitive to the cis-action of certain types of subtelomeric elements, providing a possible mechanism for the chromosome-specific telomere-length setting observed for yeast and human telomeres129. Interestingly, the same subtelomeric elements antagonize the anchoring of telomeres at the nuclear envelope, suggesting a functional link between the association of yeast telomeres with the nuclear periphery and telomerase regulation130.

Telomerase recruitment in ciliates. How far does the budding yeast model of telomerase recruitment hold true in other eukaryotes? Recent results obtained for the heterodimeric telomere end-binding protein (TebP)α–β complex from ciliated protozoa (α and β in BOX 1) shed light on new connections between cell-cycle progres-sion and telomerase recruitment. TebPβ does not bind telomeric DNA by itself but forms a heterodimer with TebPα in the presence of telomeric DNA. In an intriguing way, TebPα anchors telomeres to a sub-nuclear structure and recruits TebPβ131. by using anti-bodies against antiparallel G-quartet DNA, both TebP subunits were shown to be required for the maintenance of G quadruplexes in vivo. Strikingly, in the course of replication, Paeschke et al. have shown that TebPβ becomes phosphorylated and dissociates from the 3′ telomeric overhang–TebPα complex concomitantly with G-quadruplex resolution131. Therefore, during rep-lication, on phosphorylation of TebPβ and its dissocia-tion from TebPα, G-quadruplex DNA would be resolved through a process that involves telomerase. Chromosome ends would then be accessible for replication and extension of the G-rich strand by telomerase.

Telomerase recruitment in mammals. In humans, PoT1, an orthologue of TebPα, interacts with telomeres both through direct binding to the 3′ overhanging G-strand DNA and through an interaction with the TRF1–TRF2 duplex telomere DNA-binding complex (BOX 1). besides its role in telomere protection, PoT1 was also found to be a telomerase-dependent negative regulator of telomere length132,133 and a positive regulator of telomerase134,135. A similar duality was also reflected by the fact that human PoT1, depending on its location relative to the DNA 3′-end, could either inhibit telomerase action or form a preferred substrate for telomerase136,137. The question arises of how to reconcile these apparently contradictory observations.

Several groups independently identified a novel telomere protein named TPP1 (PoT1 binding partner; formerly named TINT1/PToP/PIP1), which interacts with both PoT1 and the TRF1-interacting factor TIN2 (RefS 138–142). Functional studies demonstrated that TPP1 heterodimerizes with PoT1 and regulates the recruit-ment of PoT1 to telomeres. Taken together, the results indicate that PoT1 and TPP1 appear to function together in end-protection and telomere-length regulation.

The determination of the crystal structure of the N-terminus of TPP1 revealed that TPP1 contains an ob fold143 that resembles the ob fold of the Oxytricha nova TebPβ subunit, suggesting that TPP1 is the human homologue of the TebPβ subunit144. Moreover, TPP1 was shown to associate with the telomerase in a man-ner that is dependent on the TPP1 ob fold, providing a physical link between telomerase and the components of the telomere145. TPP1 and PoT1 are thought to act syner-gistically to recruit telomerase to telomeres144,145. Indeed, TPP1 was shown to enhance the affinity of PoT1 and to induce a 3′-end preference for several telomeric oligo-nucleotides that contain the decamer core recognized by PoT1 (RefS 144,145). From all of these results, it is proposed that the heterodimer PoT1–TPP1 can switch from a state in which it inhibits telomerase access to the 3′ end of the telomere to a state that allows telomerase activity and processivity during telomere extension144,145. An important aim for future work in this area will be to integrate in this model of telomerase recruitment the TRF1-dependent cis-inhibition of telomerase146, which is mediated by PoT1 (Ref. 147).

Concluding remarksIn this Review, we have summarized some of the difficulties that the conventional replication machinery encounters when it reaches the chromosome ends, from end-replication problems at the two daughter chromatids to fork pausing and stalling. As a consequence, dividing cells are exposed to an inexorable erosion of telomeric DNA at each round of replication and, eventually, experience rapid telomere loss. Collectively, these replication defects are believed to have a crucial role in cell homeostasis by triggering senes-cence or apoptosis when one or several telomeres become too short to be fully functional.

Remarkably, telomeres have evolved a puzzling diver-sity of mechanisms to accommodate these problems and to reconstitute capped telomeres. A first category of mechanisms involves telomere-associated proteins to facilitate fork progression, to control G-tail formation and to recap the telomere after replication. For instance, a wealth of recent data revealed that TRF2 behaves as a ‘Swiss army knife’ for telomere replication by modulating the activity of several enzymes and by directly shaping the conformation of telomeric DNA (BOX 2; fIG. 1). However, it remains to be demonstrated that TRF2, similar to Taz1 (Ref. 11), indeed facilitates fork progression.

A second category of mechanisms relies on the transient activation of DNA-damage checkpoints upon telomere replication, which might overcome certain replication difficulties by activating repair pathways. A third category of mechanisms concerns the existence

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1. McClintock, B. The production of homozygous deficient tissues with mutant characteristics by means of the aberrant mitotic behavior of ring-shaped chromosomes. Genetics 23, 315–376 (1938).

2. Sandell, L. L. & Zakian, V. A. Loss of a yeast telomere: arrest, recovery, and chromosome loss. Cell 75, 729–739 (1993).

3. van Steensel, B., Smogorzewska, A. & de Lange, T. TRF2 protects human telomeres from end-to-end fusions. Cell 92, 401–413 (1998).

4. de Lange, T. Shelterin: the protein complex that shapes and safeguards human telomeres. Genes Dev. 19, 2100–2110 (2005).

5. Brunori, M., Luciano, P., Gilson, E. & Geli, V. The telomerase cycle: normal and pathological aspects. J. Mol. Med. 83, 244–257 (2005).

6. Greider, C. W. & Blackburn, E. H. Identification of a specific telomere terminal transferase activity in Tetrahymena extracts. Cell 43, 405–413 (1985).

7. Cech, T. R. & Brehm, S. L. Replication of the extrachromosomal ribosomal RNA genes of Tetrahymena thermophilia. Nucleic Acids Res. 9, 3531–3543 (1981).

8. Wellinger, R. J., Wolf, A. J. & Zakian, V. A. Structural and temporal analysis of telomere replication in yeast. Cold Spring Harb. Symp. Quant. Biol. 58, 725–732 (1993).

9. Ivessa, A. S., Zhou, J. Q., Schulz, V. P., Monson, E. K. & Zakian, V. A. Saccharomyces Rrm3p, a 5’ to 3’ DNA helicase that promotes replication fork progression through telomeric and subtelomeric DNA. Genes Dev. 16, 1383–1396 (2002).

10. Makovets, S., Herskowitz, I. & Blackburn, E. H. Anatomy and dynamics of DNA replication fork movement in yeast telomeric regions. Mol. Cell. Biol. 24, 4019–4031 (2004).

11. Miller, K. M., Rog, O. & Cooper, J. P. Semi-conservative DNA replication through telomeres requires Taz1. Nature 440, 824–828 (2006).Before this study it was assumed that telomere-binding proteins impede replication fork progression. Conversely, this study shows that Taz1 is crucial for efficient replication fork progression through the telomere.

12. Zahler, A. M. & Prescott, D. M. DNA primase and the replication of the telomeres in Oxytricha nova. Nucleic Acids Res. 17, 6299–6317 (1989).

13. Ray, S., Karamysheva, Z., Wang, L., Shippen, D. E. & Price, C. M. Interactions between telomerase and primase physically link the telomere and chromosome replication machinery. Mol. Cell. Biol. 22, 5859–5868 (2002).A physical association of telomerase and primase is shown in Euplotes crassus, supporting the proposed coordinated regulation of telomeric G- and C-strand synthesis.

14. Dahlen, M., Sunnerhagen, P. & Wang, T. S. Replication proteins influence the maintenance of telomere length and telomerase protein stability. Mol. Cell. Biol. 23, 3031–3042 (2003).

15. Stevenson, J. B. & Gottschling, D. E. Telomeric chromatin modulates replication timing near chromosome ends. Genes Dev. 13, 146–151 (1999).

16. Poloumienko, A., Dershowitz, A., De, J. & Newlon, C. S. Completion of replication map of Saccharomyces cerevisiae chromosome III. Mol. Biol. Cell 12, 3317–3327 (2001).

17. Deng, Z. et al. Telomeric proteins regulate episomal maintenance of Epstein-Barr virus origin of plasmid replication. Mol. Cell 9, 493–503 (2002).

18. Amiard, S. et al. A topological mechanism for TRF2-enhanced strand invasion. Nature Struct. Mol. Biol. 14, 147–154 (2007).

19. Verdun, R. E. & Karlseder, J. The DNA damage machinery and homologous recombination pathway act consecutively to protect human telomeres. Cell 127, 709–720 (2006).Shows that telomere ends need to be recognized as damaged DNA in order for end replication to be completed and for a telomere-specific structure to be formed at chromosome ends after replication.

20. Raghuraman, M. K. et al. Replication dynamics of the yeast genome. Science 294, 115–121 (2001).

21. Gottschling, D. E., Aparicio, O. M., Billington, B. L. & Zakian, V. A. Position effect at S. cerevisiae telomeres: reversible repression of Pol II transcription. Cell 63, 751–762 (1990).

22. Ferguson, B. M. & Fangman, W. L. A position effect on the time of replication origin activation in yeast. Cell 68, 333–339 (1992).

23. Zappulla, D. C., Sternglanz, R. & Leatherwood, J. Control of replication timing by a transcriptional silencer. Curr. Biol. 12, 869–875 (2002).

24. Pryde, F. E. & Louis, E. J. Limitations of silencing at native yeast telomeres. EMBO J. 18, 2538–2550 (1999).

25. Wyrick, J. J. et al. Chromosomal landscape of nucleosome-dependent gene expression and silencing in yeast. Nature 402, 418–421 (1999).

26. Cosgrove, A. J., Nieduszynski, C. A. & Donaldson, A. D. Ku complex controls the replication time of DNA in telomere regions. Genes Dev. 16, 2485–2490 (2002).

27. Hiraga, S., Robertson, E. D. & Donaldson, A. D. The Ctf18 RFC-like complex positions yeast telomeres but does not specify their replication time. EMBO J. 25, 1505–1514 (2006).

28. Raghuraman, M. K., Brewer, B. J. & Fangman, W. L. Cell cycle-dependent establishment of a late replication program. Science 276, 806–809 (1997).

29. Wang, Y., Vujcic, M. & Kowalski, D. DNA replication forks pause at silent origins near the HML locus in budding yeast. Mol. Cell. Biol. 21, 4938–4948 (2001).

30. Zou, Y., Gryaznov, S. M., Shay, J. W., Wright, W. E. & Cornforth, M. N. Asynchronous replication timing of telomeres at opposite arms of mammalian chromosomes. Proc. Natl Acad. Sci. USA 101, 12928–12933 (2004).

31. Tan, M., Jahn, C. L. & Price, C. M. Origin usage during Euplotes ribosomal DNA amplification. Eukaryot. Cell 2, 115–122 (2003).

32. Wright, W. E., Tesmer, V. M., Liao, M. L. & Shay, J. W. Normal human telomeres are not late replicating. Exp. Cell Res. 251, 492–499 (1999).

33. Hultdin, M. et al. Replication timing of human telomeric DNA and other repetitive sequences analyzed by fluorescence in situ hybridization and flow cytometry. Exp. Cell Res. 271, 223–229 (2001).

34. 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 96, 11434–11439 (1999).

35. Marcand, S., Brevet, V. & Gilson, E. Progressive cis-inhibition of telomerase upon telomere elongation. EMBO J. 18, 3509–3519 (1999).

36. Teixeira, M. T., Arneric, M., Sperisen, P. & Lingner, J. Telomere length homeostasis is achieved via a switch between telomerase-extendible and -nonextendible states. Cell 117, 323–335 (2004).Shows that telomerase does not act on every telomere in each cell cycle and that it exhibits a preference for short telomeres.

37. Bianchi, A. & Shore, D. Early replication of short telomeres in budding yeast. Cell 128, 1051–1062 (2007).

38. Shirahige, K. et al. Regulation of DNA-replication origins during cell-cycle progression. Nature 395, 618–621 (1998).

39. Santocanale, C. & Diffley, J. F. A Mec1- and Rad53-dependent checkpoint controls late-firing origins of DNA replication. Nature 395, 615–618 (1998).

40. Feng, W. et al. Genomic mapping of single-stranded DNA in hydroxyurea-challenged yeasts identifies origins of replication. Nature Cell Biol. 8, 148–155 (2006).

41. Longhese, M. P., Paciotti, V., Neecke, H. & Lucchini, G. Checkpoint proteins influence telomeric silencing and length maintenance in budding yeast. Genetics 155, 1577–1591 (2000).

42. Crabbe, L., Verdun, R. E., Haggblom, C. I. & Karlseder, J. Defective telomere lagging strand synthesis in cells lacking WRN helicase activity. Science 306, 1951–1953 (2004).Reports that cells that lack WRN show deletion of telomeres that were replicated by lagging-strand synthesis, suggesting that WRN is necessary for the efficient replication of G-rich telomeric DNA.

43. Bai, Y. & Murnane, J. P. Telomere instability in a human tumor cell line expressing a dominant-negative WRN protein. Hum. Genet. 113, 337–347 (2003).

44. Shen, J. & Loeb, L. A. Unwinding the molecular basis of the Werner syndrome. Mech. Ageing Dev. 122, 921–944 (2001).

45. Du, X. et al. Telomere shortening exposes functions for the mouse Werner and Bloom syndrome genes. Mol. Cell. Biol. 24, 8437–8446 (2004).

46. Chang, S. et al. Essential role of limiting telomeres in the pathogenesis of Werner syndrome. Nature Genet. 36, 877–882 (2004).

47. Bankhead, T., Kobryn, K. & Chaconas, G. Unexpected twist: harnessing the energy in positive supercoils to control telomere resolution. Mol. Microbiol. 62, 895–905 (2006).

48. Ivessa, A. S. & Zakian, V. A. To fire or not to fire: origin activation in Saccharomyces cerevisiae ribosomal DNA. Genes Dev. 16, 2459–2464 (2002).

49. Azvolinsky, A., Dunaway, S., Torres, J. Z., Bessler, J. B. & Zakian, V. A. The S. cerevisiae Rrm3p DNA helicase moves with the replication fork and affects replication of all yeast chromosomes. Genes Dev. 20, 3104–3116 (2006).

50. Schmidt, K. H. & Kolodner, R. D. Suppression of spontaneous genome rearrangements in yeast DNA helicase mutants. Proc. Natl Acad. Sci. USA 103, 18196–18201 (2006).

51. Opresko, P. L. et al. The Werner syndrome helicase and exonuclease cooperate to resolve telomeric D loops in a manner regulated by TRF1 and TRF2. Mol. Cell 14, 763–74 (2004).

52. Makarov, V. L., Hirose, Y. & Langmore, J. P. Long G tails at both ends of human chromosomes suggest a C strand degradation mechanism for telomere shortening. Cell 88, 657–666 (1997).

53. Dionne, I. & Wellinger, R. J. Cell cycle-regulated generation of single-stranded G-rich DNA in the absence of telomerase. Proc. Natl Acad. Sci. USA 93, 13902–13907 (1996).

54. Klobutcher, L. A., Swanton, M. T., Donini, P. & Prescott, D. M. All gene-sized DNA molecules in four species of hypotrichs have the same terminal sequence and an unusual 3’ terminus. Proc. Natl Acad. Sci. USA 78, 3015–3019 (1981).

55. Larrivee, M., LeBel, C. & Wellinger, R. J. The generation of proper constitutive G-tails on yeast telomeres is dependent on the MRX complex. Genes Dev. 18, 1391–1396 (2004).Demonstrates that G-tails are present outside S phase on normal yeast telomeres, and that Mre11 is essential to form this constitutive end structure.

56. Chai, W., Du, Q., Shay, J. W. & Wright, W. E. Human telomeres have different overhang sizes at leading versus lagging strands. Mol. Cell 21, 427–435 (2006).

of compensating pathways, including the addition of telomere repeats by telomerase. The fact that telomerase acts preferentially on shorter telomeres constitutes an efficient way to rapidly repair a brutal telomere loss. Therefore, telomerase can be considered as a repair factor that is specialized for eroded telomeres. This is consistent with the fact that many telomerase auxiliary factors are also involved in the DNA damage response.

In conclusion, the way in which telomeres are repli-cated determines their function, implying that telomeres must be studied in the context of their native replicon. Future studies on the function of these replicons will take biologists into new ground in terms of the relation-ships between chromosome organization, epigenetic regulation, the DNA damage response and cell-cycle regulation.

R E V I E W S



Shows that human diploid cells have longer G overhangs at telomeres generated by lagging-strand synthesis than by leading-strand synthesis, which suggests that leading and lagging daughter telomeres are generated differently.

57. Dionne, I. & Wellinger, R. J. Processing of telomeric DNA ends requires the passage of a replication fork. Nucleic Acids Res. 26, 5365–5371 (1998).

58. Hemann, M. T. & Greider, C. W. G-strand overhangs on telomeres in telomerase-deficient mouse cells. Nucleic Acids Res. 27, 3964–3969 (1999).

59. Ohki, R., Tsurimoto, T. & Ishikawa, F. In vitro reconstitution of the end replication problem. Mol. Cell. Biol. 21, 5753–5766 (2001).

60. Ira, G. et al. DNA end resection, homologous recombination and DNA damage checkpoint activation require CDK1. Nature 431, 1011–1017 (2004).

61. Frank, C. J., Hyde, M. & Greider, C. W. Regulation of telomere elongation by the cyclin-dependent kinase CDK1. Mol. Cell 24, 423–432 (2006).

62. Vodenicharov, M. D. & Wellinger, R. J. DNA degradation at unprotected telomeres in yeast is regulated by the CDK1 (Cdc28/Clb) cell-cycle kinase. Mol. Cell 24, 127–137 (2006).References 61 and 62 report evidence that cyclin-dependent kinase Cdk1/Cdc28 activity is required for the generation of 3′ single-strand overhangs at telomeres in S. cerevisiae.

63. Negrini, S., Ribaud, V., Bianchi, A. & Shore, D. DNA breaks are masked by multiple Rap1 binding in yeast: implications for telomere capping and telomerase regulation. Genes Dev. 21, 292–302 (2007).

64. Takata, H., Tanaka, Y. & Matsuura, A. Late S phase-specific recruitment of Mre11 complex triggers hierarchical assembly of telomere replication proteins in Saccharomyces cerevisiae. Mol. Cell 17, 573–583 (2005).

65. van Overbeek, M. & de Lange, T. Apollo, an Artemis-related nuclease, interacts with TRF2 and protects human telomeres in S phase. Curr. Biol. 16, 1295–1302 (2006).

66. Lenain, C. et al. The Apollo 5’ exonuclease functions together with TRF2 to protect telomeres from DNA repair. Curr. Biol. 16, 1303–1310 (2006).

67. Parenteau, J. & Wellinger, R. J. Accumulation of single-stranded DNA and destabilization of telomeric repeats in yeast mutant strains carrying a deletion of RAD27. Mol. Cell. Biol. 19, 4143–4152 (1999).

68. Adams Martin, A., Dionne, I., Wellinger, R. J. & Holm, C. The function of DNA polymerase α at telomeric G tails is important for telomere homeostasis. Mol. Cell. Biol. 20, 786–796 (2000).

69. Tomita, K. et al. Fission yeast Dna2 is required for generation of the telomeric single-strand overhang. Mol. Cell. Biol. 24, 9557–9567 (2004).

70. Hubscher, U., Maga, G. & Spadari, S. Eukaryotic DNA polymerases. Annu. Rev. Biochem. 71, 133–163 (2002).

71. Qi, H. & Zakian, V. A. The Saccharomyces telomere-binding protein Cdc13p interacts with both the catalytic subunit of DNA polymerase α and the telomerase-associated est1 protein. Genes Dev. 14, 1777–88 (2000).

72. Grossi, S., Puglisi, A., Dmitriev, P. V., Lopes, M. & Shore, D. Pol12, the B subunit of DNA polymerase α, functions in both telomere capping and length regulation. Genes Dev. 18, 992–1006 (2004).

73. Carson, M. J. & Hartwell, L. CDC17: an essential gene that prevents telomere elongation in yeast. Cell 42, 249–257 (1985).

74. Wellinger, R. J., Wolf, A. J. & Zakian, V. A. Saccharomyces telomeres acquire single-strand TG1–3 tails late in S phase. Cell 72, 51–60 (1993).

75. Bertuch, A. A. & Lundblad, V. The Ku heterodimer performs separable activities at double-strand breaks and chromosome termini. Mol. Cell. Biol. 23, 8202–8215 (2003).

76. Churikov, D., Wei, C. & Price, C. M. Vertebrate POT1 restricts G-overhang length and prevents activation of a telomeric DNA damage checkpoint but is dispensable for overhang protection. Mol. Cell. Biol. 26, 6971–6982 (2006).

77. Baumann, P. Are mouse telomeres going to pot? Cell 126, 33–36 (2006).

78. Zhu, X. D. et al. ERCC1/XPF removes the 3’ overhang from uncapped telomeres and represses formation of telomeric DNA-containing double minute chromosomes. Mol. Cell 12, 1489–1498 (2003).

79. Celli, G. B. & de Lange, T. DNA processing is not required for ATM-mediated telomere damage

response after TRF2 deletion. Nature Cell Biol. 7, 712–718 (2005).

80. Rossi, M. L., Purohit, V., Brandt, P. D. & Bambara, R. A. Lagging strand replication proteins in genome stability and DNA repair. Chem. Rev. 106, 453–473 (2006).

81. Budd, M. E., Reis, C. C., Smith, S., Myung, K. & Campbell, J. L. Evidence suggesting that Pif1 helicase functions in DNA replication with the Dna2 helicase/nuclease and DNA polymerase δ. Mol. Cell. Biol. 26, 2490–2500 (2006).

82. Qiu, J., Qian, Y., Frank, P., Wintersberger, U. & Shen, B. Saccharomyces cerevisiae RNase H(35) functions in RNA primer removal during lagging-strand DNA synthesis, most efficiently in cooperation with Rad27 nuclease. Mol. Cell. Biol. 19, 8361–8371 (1999).

83. Jeong, H. S., Backlund, P. S., Chen, H. C., Karavanov, A. A. & Crouch, R. J. RNase H2 of Saccharomyces cerevisiae is a complex of three proteins. Nucleic Acids Res. 32, 407–414 (2004).

84. Reveal, P. M., Henkels, K. M. & Turchi, J. J. Synthesis of the mammalian telomere lagging strand in vitro. J. Biol. Chem. 272, 11678–11681 (1997).

85. Fan, X. & Price, C. M. Coordinate regulation of G- and C strand length during new telomere synthesis. Mol. Biol. Cell 8, 2145–2155 (1997).

86. Jacob, N. K., Kirk, K. E. & Price, C. M. Generation of telomeric G strand overhangs involves both G and C strand cleavage. Mol. Cell 11, 1021–1032 (2003).

87. Sfeir, A. J., Chai, W., Shay, J. W. & Wright, W. E. Telomere-end processing the terminal nucleotides of human chromosomes. Mol. Cell 18, 131–138 (2005).

88. Hockemeyer, D., Sfeir, A. J., Shay, J. W., Wright, W. E. & de Lange, T. POT1 protects telomeres from a transient DNA damage response and determines how human chromosomes end. EMBO J. 24, 2667–2678 (2005).

89. 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. 20, 5532–5540 (2001).

90. Fouche, N. et al. The basic domain of TRF2 directs binding to DNA junctions irrespective of the presence of TTAGGG repeats. J. Biol. Chem. 281, 37486–37495 (2006).

91. Bailey, S. M., Cornforth, M. N., Kurimasa, A., Chen, D. J. & Goodwin, E. H. Strand-specific postreplicative processing of mammalian telomeres. Science 293, 2462–2465 (2001).

92. Wang, R. C., Smogorzewska, A. & de Lange, T. Homologous recombination generates T-loop-sized deletions at human telomeres. Cell 119, 355–368 (2004).

93. Gotta, M. et al. The clustering of telomeres and colocalization with Rap1, Sir3, and Sir4 proteins in wild-type Saccharomyces cerevisiae. J. Cell Biol. 134, 1349–1363 (1996).

94. Heun, P., Laroche, T., Shimada, K., Furrer, P. & Gasser, S. M. Chromosome dynamics in the yeast interphase nucleus. Science 294, 2181–2186 (2001).

95. Marcand, S., Brevet, V., Mann, C. & Gilson, E. Cell cycle restriction of telomere elongation. Curr. Biol. 10, 487–490 (2000).Shows that in budding yeast cells that progress synchronously through the cell cycle, telomere elongation coincides with the time of telomere replication.

96. Diede, S. J. & Gottschling, D. E. Telomerase-mediated telomere addition in vivo requires DNA primase and DNA polymerases α and δ. Cell 99, 723–733 (1999).Shows that the essential DNA polymerase-α and -δ and DNA primase are required for telomerase function, indicating that telomeric DNA synthesis by telomerase is tightly coregulated with the production of the opposite strand.

97. Taggart, A. K., Teng, S. C. & Zakian, V. A. Est1p as a cell cycle-regulated activator of telomere-bound telomerase. Science 297, 1023–1026 (2002).This study correlates the timing of telomere elongation in budding yeast with the binding at the telomeres of several proteins that are involved in telomere elongation, including the telomerase holoenzyme.

98. Schramke, V. et al. RPA regulates telomerase action by providing Est1p access to chromosome ends. Nature Genet. 36, 46–54 (2004).Shows that in budding yeast, RPA binds to telomeres at the end of S phase and is required for telomerase action.

99. Bianchi, A., Negrini, S. & Shore, D. Delivery of yeast telomerase to a DNA break depends on the recruitment functions of Cdc13 and Est1. Mol. Cell 16, 139–146 (2004).

100. Osterhage, J. L., Talley, J. M. & Friedman, K. L. Proteasome-dependent degradation of Est1p regulates the cell cycle-restricted assembly of telomerase in Saccharomyces cerevisiae. Nature Struct. Mol. Biol. 13, 720–728 (2006).

101. Goudsouzian, L. K., Tuzon, C. T. & Zakian, V. A. S. cerevisiae Tel1p and Mre11p are required for normal levels of Est1p and Est2p telomere association. Mol. Cell 24, 603–610 (2006).

102. Stellwagen, A. E., Haimberger, Z. W., Veatch, J. R. & Gottschling, D. E. Ku interacts with telomerase RNA to promote telomere addition at native and broken chromosome ends. Genes Dev. 17, 2384–2395 (2003).

103. Fisher, T. S., Taggart, A. K. & Zakian, V. A. Cell cycle-dependent regulation of yeast telomerase by Ku. Nature Struct. Mol. Biol. 11, 1198–1205 (2004).

104. Evans, S. K. & Lundblad, V. Est1 and Cdc13 as comediators of telomerase access. Science 286, 117–120 (1999).

105. Nugent, C. I., Hughes, T. R., Lue, N. F. & Lundblad, V. Cdc13p: a single-strand telomeric DNA-binding protein with a dual role in yeast telomere maintenance. Science 274, 249–252 (1996).

106. Pennock, E., Buckley, K. & Lundblad, V. Cdc13 delivers separate complexes to the telomere for end protection and replication. Cell 104, 387–396 (2001).

107. Grandin, N., Damon, C. & Charbonneau, M. Cdc13 cooperates with the yeast Ku proteins and Stn1 to regulate telomerase recruitment. Mol. Cell. Biol. 20, 8397–8408 (2000).

108. Seto, A. G., Zaug, A. J., Sobel, S. G., Wolin, S. L. & Cech, T. R. Saccharomyces cerevisiae telomerase is an Sm small nuclear ribonucleoprotein particle. Nature 401, 177–180 (1999).

109. Seto, A. G., Livengood, A. J., Tzfati, Y., Blackburn, E. H. & Cech, T. R. A bulged stem tethers Est1p to telomerase RNA in budding yeast. Genes Dev. 16, 2800–2812 (2002).

110. Peterson, S. E. et al. The function of a stem-loop in telomerase RNA is linked to the DNA repair protein Ku. Nature Genet. 27, 64–67 (2001).

111. Dandjinou, A. T. et al. A phylogenetically based secondary structure for the yeast telomerase RNA. Curr. Biol. 14, 1148–1158 (2004).

112. Zappulla, D. C. & Cech, T. R. Yeast telomerase RNA: a flexible scaffold for protein subunits. Proc. Natl Acad. Sci. USA 101, 10024–10029 (2004).Based on the interactions of yeast telomerase RNA TLC1 with Est1, Ku and Sm proteins, this study proposes that TLC1 provides a flexible tether for these proteins.

113. Zappulla, D. C., Goodrich, K. & Cech, T. R. A miniature yeast telomerase RNA functions in vivo and reconstitutes activity in vitro. Nature Struct. Mol. Biol. 12, 1072–1077 (2005).

114. Gao, H., Cervantes, R. B., Mandell, E. K., Otero, J. H. & Lundblad, V. RPA-like proteins mediate yeast telomere function. Nature Struct. Mol. Biol. 14, 208–214 (2007).

115. Grossi, S., Bianchi, A., Damay, P. & Shore, D. Telomere formation by rap1p binding site arrays reveals end-specific length regulation requirements and active telomeric recombination. Mol. Cell. Biol. 21, 8117–8128 (2001).

116. Boule, J. B., Vega, L. R. & Zakian, V. A. The yeast Pif1p helicase removes telomerase from telomeric DNA. Nature 438, 57–61 (2005).Suggests that Pif1 RNA/DNA helicase activity limits telomerase action by displacing active telomerase from DNA ends.

117. Eugster, A. et al. The finger subdomain of yeast telomerase cooperates with Pif1p to limit telomere elongation. Nature Struct. Mol. Biol. 13, 734–739 (2006).

118. Marcand, S., Gilson, E. & Shore, D. A protein-counting mechanism for telomere length regulation in yeast. Science 275, 986–990 (1997).

119. Naito, T., Matsuura, A. & Ishikawa, F. Circular chromosome formation in a fission yeast mutant defective in two ATM homologues. Nature Genet. 20, 203–206 (1998).

120. Chan, S. W., Chang, J., Prescott, J. & Blackburn, E. H. Altering telomere structure allows telomerase to act in yeast lacking ATM kinases. Curr. Biol. 11, 1240–1250 (2001).

R E V I E W S



121. Greenwell, P. W. et al. TEL1, a gene involved in controlling telomere length in S. cerevisiae, is homologous to the human ataxia telangiectasia gene. Cell 82, 823–829 (1995).

122. Viscardi, V., Baroni, E., Romano, M., Lucchini, G. & Longhese, M. P. Sudden telomere lengthening triggers a Rad53-dependent checkpoint in Saccharomyces cerevisiae. Mol. Biol. Cell 14, 3126–3143 (2003).

123. Nakamura, T. M., Moser, B. A. & Russell, P. Telomere binding of checkpoint sensor and DNA repair proteins contributes to maintenance of functional fission yeast telomeres. Genetics 161, 1437–1452 (2002).

124. Sabourin, M., Tuzon, C.T. & Zakian, V.A. Telomerase and Tel1p preferentially associate with short telomeres in S. cerevisiae. Mol. Cell 27, 550–561 (2007).

125. Bianchi, A. & Shore, D. Increased association of telomerase with short telomeres in yeast. Genes Dev. 21, 1726–1730 (2007).

126. Hector, R.E. et al. Tel1p preferentially associates with short telomeres to stimulate their elongation. Mol. Cell (in the press).

127. Arneriç, M. & Lingner, J. Tel1p kinase and subtelomere bound Tbf1p mediate preferential elongation of short telomeres by telomerase in yeast. EMBO Rep. (in the press).

128. Chang, M., Arneric, M., & Lingner, J. Telomerase repeat addition processivity is increased at critically short telomeres in a Tel1-dependent manner in Saccharomyces cerevisiae. Genes Dev. (in the press).

129. Berthiau, A. S. et al. Subtelomeric proteins negatively regulate telomere elongation in budding yeast. EMBO J. 25, 846–856 (2006).

130. Hediger, F., Berthiau, A. S., van Houwe, G., Gilson, E. & Gasser, S. M. Subtelomeric factors antagonize telomere anchoring and Tel1-independent telomere length regulation. EMBO J. 25, 857–867 (2006).

131. Paeschke, K., Simonsson, T., Postberg, J., Rhodes, D. & Lipps, H. J. Telomere end-binding proteins control the formation of G-quadruplex DNA structures in vivo. Nature Struct. Mol. Biol. 12, 847–854 (2005).

132. Loayza, D., Parsons, H., Donigian, J., Hoke, K. & de Lange, T. DNA binding features of human POT1: a nonamer 5’-TAGGGTTAG-3’ minimal binding site, sequence specificity, and internal binding to multimeric sites. J. Biol. Chem. 279, 13241–13248 (2004).

133. Ye, J. Z. et al. POT1-interacting protein PIP1: a telomere length regulator that recruits POT1 to the TIN2/TRF1 complex. Genes Dev. 18, 1649–1654 (2004).

134. Colgin, L. M., Baran, K., Baumann, P., Cech, T. R. & Reddel, R. R. Human POT1 facilitates telomere elongation by telomerase. Curr. Biol. 13, 942–946 (2003).

135. Armbruster, B. N. et al. Rescue of an hTERT mutant defective in telomere elongation by fusion with hPot1. Mol. Cell. Biol. 24, 3552–3561 (2004).

136. Kelleher, C., Kurth, I. & Lingner, J. Human protection of telomeres 1 (POT1) is a negative regulator of telomerase activity in vitro. Mol. Cell. Biol. 25, 808–818 (2005).

137. Lei, M., Zaug, A. J., Podell, E. R. & Cech, T. R. Switching human telomerase on and off with hPOT1 protein in vitro. J. Biol. Chem. 280, 20449–20456 (2005).

138. Kim, S. H. et al. TIN2 mediates functions of TRF2 at human telomeres. J. Biol. Chem. 279, 43799–43804 (2004).

139. Liu, D. et al. PTOP interacts with POT1 and regulates its localization to telomeres. Nature Cell Biol. 6, 673–680 (2004).

140. Ye, J. Z. et al. TIN2 binds TRF1 and TRF2 simultaneously and stabilizes the TRF2 complex on telomeres. J. Biol. Chem. 279, 47264–47271 (2004).

141. O’Connor, M. S., Safari, A., Xin, H., Liu, D. & Songyang, Z. A critical role for TPP1 and TIN2 interaction in high-order telomeric complex assembly. Proc. Natl Acad. Sci. USA 103, 11874–11879 (2006).

142. Houghtaling, B. R., Cuttonaro, L., Chang, W. & Smith, S. A dynamic molecular link between the telomere length regulator TRF1 and the chromosome end protector TRF2. Curr. Biol. 14, 1621–1631 (2004).

143. Murzin, A. G. OB(oligonucleotide/oligosaccharide binding)-fold: common structural and functional solution for non-homologous sequences. EMBO J. 12, 861–867 (1993).

144. Wang, F. et al. The POT1–TPP1 telomere complex is a telomerase processivity factor. Nature 445, 506–510 (2007).

145. Xin, H. et al. TPP1 is a homologue of ciliate TEBP-β and interacts with POT1 to recruit telomerase. Nature 445, 559–562 (2007).References 144 and 145 show that the human telomeric proteins TPP1 and POT1 form a complex that regulates telomerase access to the telomere and increases the processivity of the telomerase core enzyme.

146. Ancelin, K. et al. Targeting assay to study the cis functions of human telomeric proteins: evidence for inhibition of telomerase by TRF1 and for activation of telomere degradation by TRF2. Mol. Cell. Biol. 22, 3474–3487 (2002).

147. Loayza, D. & De Lange, T. POT1 as a terminal transducer of TRF1 telomere length control. Nature 423, 1013–1018 (2003).

148. Gottschling, D. E. & Cech, T. R. Chromatin structure of the molecular ends of Oxytricha macronuclear DNA: phased nucleosomes and a telomeric complex. Cell 38, 501–510 (1984).

149. Wright, J. H., Gottschling, D. E. & Zakian, V. A. Saccharomyces telomeres assume a non-nucleosomal chromatin structure. Genes Dev. 6, 197–210 (1992).

150. Teixeira, M. T. & Gilson, E. Telomere maintenance, function and evolution: the yeast paradigm. Chromosome Res. 13, 535–548 (2005).

151. Blasco, M. A. The epigenetic regulation of mammalian telomeres. Nature Rev. Genet. 8, 299–309 (2007).

152. Baur, J. A., Zou, Y., Shay, J. W. & Wright, W. E. Telomere position effect in human cells. Science 292, 2075–2077 (2001).

153. Koering, C. E. et al. Human telomeric position effect is determined by chromosomal context and telomeric chromatin integrity. EMBO Rep. 3, 1055–1061 (2002).

154. Chong, L. et al. A human telomeric protein. Science 270, 1663–1667 (1995).

155. Broccoli, D., Smogorzewska, A., Chong, L. & de Lange, T. Human telomeres contain two distinct Myb-related proteins, TRF1 and TRF2. Nature Genet. 17, 231–235 (1997).

156. Bilaud, T. et al. Telomeric localization of TRF2, a novel human telobox protein. Nature Genet. 17, 236–239 (1997).

157. Griffith, J. D. et al. Mammalian telomeres end in a large duplex loop. Cell 97, 503–514 (1999).

158. Opresko, P. L. et al. POT1 stimulates RecQ helicases WRN and BLM to unwind telomeric DNA substrates. J. Biol. Chem. 280, 32069–32080 (2005).

159. 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 102, 10864–10869 (2005).

160. Griffith, J., Bianchi, A. & de Lange, T. TRF1 promotes parallel pairing of telomeric tracts in vitro. J. Mol. Biol. 278, 79–88 (1998).

161. Bae, N. S. & Baumann, P. A RAP1/TRF2 complex inhibits nonhomologous end-joining at human telomeric DNA ends. Mol. Cell 26, 323–334 (2007).

162. Bryan, T. M., Englezou, A., Dalla-Pozza, L., Dunham, M. A. & Reddel, R. R. Evidence for an alternative mechanism for maintaining telomere length in human tumors and tumor-derived cell lines. Nature Med. 3, 1271–1274 (1997).

163. Marciniak, R. A. et al. A novel telomere structure in a human alternative lengthening of telomeres cell line. Cancer Res. 65, 2730–2737 (2005).

164. McEachern, M. J. & Haber, J. E. Break-induced replication and recombinational telomere elongation in yeast. Annu. Rev. Biochem. 75, 111–135 (2006).

165. Biessmann, H. & Mason, J. M. Telomere maintenance without telomerase. Chromosoma 106, 63–69 (1997).

166. Pardue, M. L. et al. Two retrotransposons maintain telomeres in Drosophila. Chromosome Res. 13, 443–453 (2005).

167. Morrish, T. A. et al. Endonuclease-independent LINE-1 retrotransposition at mammalian telomeres. Nature 446, 208–212 (2007).

168. Salas, T. R. et al. Human replication protein A unfolds telomeric G-quadruplexes. Nucleic Acids Res. 34, 4857–4865 (2006).

169. Muftuoglu, M. et al. Telomere repeat binding factor 2 interacts with base excision repair proteins and stimulates DNA synthesis by DNA polymerase β. Cancer Res. 66, 113–124 (2006).

Acknowledgements We would like to thank M.-J. Giraud-Panis, T. Teixeira, A. Londono-Vallejo and P. Luciano for critical reading and helpful discussions. The E.G. and V.G. laboratories are sup-ported by ‘La Ligue Nationale contre le Cancer’ (‘Equipes labellisées’). We apologize for all the important papers that could not be cited due to space limitations.

Competing interests statementThe authors declare no competing financial interests.

DATABASESEntrez Genome Project: http://www.ncbi.nlm.nih.gov/sites/entrez?db=genomeprjBorrelia burgdorferiUniProtKB: http://ca.expasy.org/sprotBLM | Cdc28 | FEN1 | Ku | Pif1 | POT1 | Rap1 | Sir3 | TRF2 | WRN

FURTHER INFORMATIONEric Gilson’s laboratory web site: http://www.ens-lyon.fr/LBMCVincent Géli’s laboratory web site: http://www.igc.cnrs-mrs.fr/spip?article=61

all links are acTive in The online pDf

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http://www.ncbi.nlm.nih.gov/sites/entrez?db=genomeprj

http://www.ncbi.nlm.nih.gov/sites/entrez?db=genomeprj

http://www.ncbi.nlm.nih.gov/sites/entrez?Db=genomeprj&cmd=ShowDetailView&TermToSearch=3

http://ca.expasy.org/sprot






http://www.expasy.org/uniprot/Q9NUX5





http://www.ens-lyon.fr/LBMC

http://www.igc.cnrs-mrs.fr/spip?article=61

how telomeres are replicated

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