The EMBO Journal (2009) 28, 810–820
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2009 European Molecular Biology Organization | All Rights Reserved 0261-4189/09
THE
EMBO
JOURNAL
Differential arrival of leading and lagging strand
DNA polymerases at fission yeast telomeres
Bettina A Moser1, Lakxmi Subramanian1,
Ya-Ting Chang1, Chiaki Noguchi2,
Eishi Noguchi2 and Toru M Nakamura1,*
1
Department of Biochemistry and Molecular Genetics, University of
Illinois at Chicago, Chicago, IL, USA and 2Department of Biochemistry
and Molecular Biology, Drexel University College of Medicine,
Philadelphia, PA, USA
To maintain genomic integrity, telomeres must undergo
switches from a protected state to an accessible state that
allows telomerase recruitment. To better understand how
telomere accessibility is regulated in fission yeast, we
analysed cell cycle-dependent recruitment of telomerespecific proteins (telomerase Trt1, Taz1, Rap1, Pot1 and
Stn1), DNA replication proteins (DNA polymerases, MCM,
RPA), checkpoint protein Rad26 and DNA repair protein
Nbs1 to telomeres. Quantitative chromatin immunoprecipitation studies revealed that MCM, Nbs1 and Stn1 could
be recruited to telomeres in the absence of telomere
replication in S-phase. In contrast, Trt1, Pot1, RPA and
Rad26 failed to efficiently associate with telomeres unless
telomeres are actively replicated. Unexpectedly, the leading strand DNA polymerase e (Pole) arrived at telomeres
earlier than the lagging strand DNA polymerases a (Pola)
and d (Pold). Recruitment of RPA and Rad26 to telomeres
matched arrival of DNA Pole, whereas S-phase specific
recruitment of Trt1, Pot1 and Stn1 matched arrival of DNA
Pola. Thus, the conversion of telomere states involves an
unanticipated intermediate step where lagging strand
synthesis is delayed until telomerase is recruited.
The EMBO Journal (2009) 28, 810–820. doi:10.1038/
emboj.2009.31; Published online 12 February 2009
Subject Categories: cell cycle; genome stability & dynamics
Keywords: cell cycle; DNA polymerase; DNA replication;
pot1; telomerase
Introduction
To maintain genomic integrity, telomeres must fulfill two
major functions (Blackburn, 2001). First, telomeres must be
able to prevent DNA repair and DNA damage checkpoint
proteins from causing uncontrolled fusion and degradation of
telomeric DNA and eliciting permanent cell cycle arrest.
Second, telomeres must provide access to telomerase, a
reverse transcriptase that can extend the GT-rich telomeric
repeat sequence by using its RNA subunit as a template.
*Corresponding author. Department of Biochemistry and Molecular
Genetics, University of Illinois at Chicago, 900 S. Ashland Ave. MC669,
Chicago, IL 60607, USA. Tel.: þ 1 312 996 1988; Fax: þ 1 312 996 1988;
E-mail: [email protected]
Received: 24 October 2008; accepted: 20 January 2009; published
online: 12 February 2009
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Recruitment of telomerase is essential to prevent loss of
telomeric DNA after each round of DNA replication because
replicative DNA polymerases cannot fully replicate ends of
linear DNA molecules. Thus, telomeres must undergo dynamic switches from a highly protected state that prevents
full access to DNA repair and DNA damage checkpoint
proteins to a more accessible state that allows recruitment
of telomerase, as well as various DNA repair and DNA
damage checkpoint proteins (Verdun and Karlseder, 2007).
Previous studies have found that such changes in telomere
status are likely to occur in a cell cycle-dependent manner
(Gilson and Geli, 2007). Specifically, replication of telomeres
has been proposed to have an important function in controlling accessibility of telomeres.
Evolutionarily conserved unique structural features of
telomeric DNA include species-specific GT-rich repetitive
double-stranded DNA (dsDNA) as well as a 30 single-stranded
GT-rich overhang (G-tail). These features are important for
both protection and replication associated functions of telomeres as they provide binding sites for telomere-specific
proteins and telomerase (Verdun and Karlseder, 2007). The
G-tail is required for the extension of telomeric DNA by
telomerase, as the telomerase RNA subunit cannot anneal
to blunt ends (Lingner and Cech, 1996). Loss of telomerespecific factors, such as the budding yeast G-tail-binding
capping protein Cdc13, can lead to degradation of the telomeric CA-rich strand, massive accumulation of DNA repair
and checkpoint factors at telomeres, and cell cycle arrest
(Melo et al, 2001). The length of the G-tail is cell cycle
regulated, and increases during S-phase in both budding
yeast Saccharomyces cerevisiae and fission yeast
Schizosaccharomyces pombe (Wellinger et al, 1996; Tomita
et al, 2004).
Semi-conservative DNA replication of linear chromosomes
generates two structurally distinct termini at telomeres (Ohki
et al, 2001). The strand replicated by lagging strand synthesis
will leave a 30 single-stranded overhang (G-tail), whereas the
strand replicated by leading strand synthesis will likely
produce a blunt terminus. Therefore, leading-strand telomeres must be processed to re-generate the G-tail. Evidence
for this processing comes from studies showing that both
ends of chromosomes terminate in a 30 overhang in yeasts,
ciliates and humans (Wellinger et al, 1993; Makarov et al,
1997; Jacob et al, 2001; Munoz-Jordan et al, 2001). This end
processing does not require telomerase, as the 30 overhang
was still present at both ends of chromosomes in the absence
of telomerase (Wellinger et al, 1996; Makarov et al, 1997).
The existence of two distinct types of end processing mechanisms at telomeres has been supported by studies that
observed (1) chromosomal fusions only among leadingstrand telomeres in mammalian cells carrying mutant versions of TRF2 or DNA-PKcs (Bailey et al, 2001), (2) preferential loss of lagging-strand telomeres in human cells
defective in RecQ DNA helicase WRN (Crabbe et al, 2004)
and (3) much longer G-tails on lagging-strand telomeres than
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leading-strand telomeres in human cells lacking active telomerase (Chai et al, 2006a).
Chromatin immunoprecipitation (ChIP) analyses in
S. cerevisiae have shown increased association of the G-tailbinding protein Cdc13 and the telomerase subunit Est1 in late
S-phase (Taggart et al, 2002; Schramke et al, 2004). Est2, the
catalytic subunit of telomerase, is loaded to telomeres in G1phase through specific interaction between the Ku70–Ku80
complex and telomerase RNA, but it also shows increased
association with telomeres during late S-phase (Fisher et al,
2004). Timing of maximum recruitment for the singlestranded DNA (ssDNA)-binding protein complex RPA (replication protein A) and DNA polymerase e to telomeres coincide with the timing of the maximum recruitment for Est1
and Cdc13, suggesting that Est1 and Cdc13 are recruited to
telomeres when the replication fork arrives at telomeres in
budding yeast (Schramke et al, 2004; Bianchi and Shore,
2007). However, it is also possible that MRX (Mre11–Rad50–
Xrs2)-dependent resection of the CA-strand, rather than arrival of the DNA replication fork at telomeres itself, is responsible for the observed RPA loading to telomeres, as increased
binding of the G-tail-binding protein Cdc13 to telomeres
during late S-phase requires MRX-dependent generation of
long G-tails (Larrivee et al, 2004; Takata et al, 2005;
Goudsouzian et al, 2006). Association of budding yeast
MRX with telomeres also increases during late S-phase
(Takata et al, 2005). Likewise, S-phase specific association
of human NBS1 (budding yeast Xrs2 ortholog) with telomeres
has been observed (Wu et al, 2000; Zhu et al, 2000). On the
other hand, unlike yeast MRX, human MRE11 and RAD50
appear to constitutively associate with telomeres, perhaps
through specific interaction with TRF2 (Zhu et al, 2000). This
difference might be due to the fact that S. cerevisiae lacks
TRF1/TRF2-like proteins.
In fact, S. cerevisiae has diverged significantly in telomere
protein components from humans, whereas telomere components in fission yeast S. pombe are more closely related to
human telomere proteins. For example, S. cerevisiae lacks
orthologs for human telomeric dsDNA-binding proteins TRF1
and TRF2, whereas S. pombe Taz1 shows sequence and
functional similarity to both TRF1 and TRF2. In S. cerevisiae,
Rap1 directly binds to telomeric dsDNA. In contrast, human
and S. pombe Rap1 proteins do not directly bind to telomeric
DNA, but they are brought to telomeres through protein–
protein interaction with TRF2 and Taz1, respectively
(Smogorzewska and de Lange, 2004). Budding yeast Cdc13
also lacks any obvious ortholog in S. pombe or human,
although Pot1 proteins from both human and S. pombe are
likely to play similar functional roles as Cdc13. It was recently
shown that S. pombe Pot1 forms a multi-protein complex that
closely resembles the mammalian shelterin complex
(Miyoshi et al, 2008; Palm and de Lange, 2008). Therefore,
although studies in S. cerevisiae have provided the most
detailed molecular description of telomere components to
date, S. pombe should provide new insights as a model
system more closely resembling the mechanisms of human
telomere maintenance.
Fission yeast telomeres are replicated very late in S-phase
(Kim and Huberman, 2001). Presumably, late replication of
telomeres is the consequence of a local chromatin environment, which is composed of GT-rich repetitive telomeric
repeat DNA (both double-stranded and single-stranded with
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30 overhang), telomere-specific dsDNA-binding proteins
(Taz1 and Rap1), telomere-specific ssDNA-binding proteins
(Pot1–Tpz1–Poz1–Ccq1 and Stn1–Ten1 complexes), various
dsDNA break repair proteins (Ku70–Ku80 and Mre11–Rad50–
Nbs1 complexes), and DNA damage checkpoint proteins
(Rad3–Rad26 and Rad9–Rad1–Hus1 complexes) (Kanoh and
Ishikawa, 2001; Nakamura et al, 2002; Martin et al, 2007;
Miyoshi et al, 2008). Furthermore, proteins involved in
heterochromatin formation might also have a function in
establishing late S-phase replication of telomeres (Mickle
et al, 2007). All these proteins are expected to contribute in
regulating telomere accessibility changes during the cell
cycle; however, currently no data are available with regard
to how binding of telomere-associated proteins at telomeres
is regulated during the fission yeast cell cycle. Therefore, we
decided to characterise cell cycle specific changes in telomere
protein composition and directly co-relate them with the
timing of telomere replication.
Results
Telomere replication occurs in late S-phase and
is inhibited by hydroxyurea
To investigate how late S-phase replication of telomeres is
established and coordinated with the extension of telomeric
repeats by telomerase in fission yeast, we used the temperature sensitive cdc25-22 allele to synchronise cells (Alfa et al,
1993). After incubating cdc25-22 cells at the non-permissive
temperature (361C) for 3 h, late G2-phase arrested cells were
shifted to the permissive temperature (251C) for synchronous
cell cycle re-entry, and samples were collected every 20 min
and processed for ChIP analysis. On the basis of 5-bromo-20 deoxyuridine (BrdU) incorporation analysis (Hodson et al,
2003), the early replication origins ars2004 and ars3002
replicated 60–100 min after the temperature shift, whereas
telomeres replicated at 100–140 min (Figure 1A and data not
shown), confirming the previously established timing of
replication for these regions (Kim and Huberman, 2001). In
agreement with previous studies, when cells were released
from the cdc25-22 arrest in the presence of 15 mM hydroxyurea (HU), BrdU incorporation at telomeres was abolished,
whereas BrdU was still efficiently incorporated at ars2004
(Hayashi et al, 2007; Mickle et al, 2007; Supplementary
Figure S3). Previous studies have shown that firing of late
origins, including telomeres, are extremely delayed and/or
inhibited by the addition of HU, and that prevention of origin
firing at telomeres in the presence of HU requires S-phase
checkpoint proteins, such as Rad3 and Cds1 kinases (Hayashi
et al, 2007; Mickle et al, 2007).
We next monitored the cell cycle-regulated recruitment of
the hexameric MCM complex to telomeres and ars2004. MCM
is loaded to DNA through the function of the origin recognition complex (ORC), and it is believed to be the replicative
helicase required for both initiation and elongation stages of
DNA replication (Bochman and Schwacha, 2008). Recent
genome-wide ChIP analyses have found that sub-telomeric
regions are highly enriched with ORC and MCM (Hayashi
et al, 2007). However, kinetics of MCM loading to telomeres
has not been compared with other replicating origins. We
found that recruitment of Mcm2 (a subunit of the MCM
complex) to telomeres was delayed (binding peak at
100 min) compared with ars2004 (binding peak at 80 min)
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Figure 1 Late S-phase replication of telomeres is reflected in delayed recruitment of MCM and RPA to telomeres compared with early
replication origin ars2004. Data are from multiple (n) independent experiments, and error bars represent average deviation (n ¼ 2) or standard
deviation (n ¼ 3). For the definition of relative precipitation (Rel. precipitation), see ‘ChIP analysis’ sub-section in the Materials and methods.
These values represent background corrected percentage precipitation, normalised to the peak values in minus HU experiments. (A) BrdU
incorporation at telomeres and ars2004 for cdc25-22 synchronised cells (n ¼ 3). (B) Recruitment of MCM (Mcm2) to telomeres and ars2004 for
cdc25-22 synchronised cells (n ¼ 2). (C) Recruitment of RPA (Rad11) to telomeres and ars2004 for cdc25-22 synchronised cells (n ¼ 2).
(D) Comparison of peak percentage precipitation values (see Materials and methods) at telomeres and ars2004 in the absence or presence of
15 mM HU for MCM and RPA. The peak percentage precipitation value ratios (telomere/ars2004) in the absence or presence of HU were
calculated for the indicated proteins.
(Figure 1B). At both telomeres and ars2004, recruitment of
Mcm2 preceded BrdU incorporation; however, we observed a
greater time lag between MCM loading and BrdU incorporation at telomeres than at ars2004 (Supplementary Figure S4A
and B). Therefore, we conclude that delayed loading of MCM
to telomeres is only partially responsible for establishing the
late S-phase replication of telomeric DNA.
In the presence of HU, increased binding of Mcm2 was
observed at both telomeres and ars2004 (Supplementary
Figure S4C and D). Although MCM dissociated from
ars2004 as DNA replication was completed, MCM remained
bound to telomeres for the duration of our experiment,
correlating well with the observation that DNA replication
is inhibited at telomeres in the presence of HU
(Supplementary Figures S3, S4C and D). HU had little effect
on the initial recruitment timing of MCM to telomeres.
Moreover, the ratio of the peak percentage precipitation
values at telomeres against ars2004 for MCM was altered
only slightly by the addition of HU (Figure 1D). Therefore, it
appears that inhibition of DNA replication at telomeres in the
presence of HU is caused by inhibition of a step after loading
of MCM to telomeres.
Another hallmark of DNA replication is recruitment of the
ssDNA-binding protein RPA after the replicative helicase unwinds DNA. Therefore, we next monitored recruitment of
RPA to telomeres and ars2004. As shown in Figure 1C, we
found that Rad11 (the largest subunit of the RPA complex) is
recruited maximally to telomeres at 120 min after release
from the cdc25-22 arrest, compared with 80 min at ars2004.
RPA binding to telomeres matched closely to the timing of
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BrdU incorporation at telomeres, whereas RPA binding to
ars2004 preceded BrdU incorporation and more closely resembled the timing of Mcm2 recruitment (Supplementary
Figure S5).
Interestingly, we also found that RPA was able to precipitate approximately four-fold more telomeric DNA than
ars2004 DNA when the peak percentage precipitation values
were compared (Figure 1D). Thus, telomeres appear to
accumulate significantly more ssDNA than ars2004 when
the DNA replication fork moves in. In the presence of HU,
the DNA replication-induced peak of RPA binding at telomeres was essentially eliminated, consistent with the observation that telomeres are not replicated in the presence of HU
(Supplementary Figure S5C). In contrast, RPA binding to
ars2004 was extended and increased approximately fivefold in the presence of HU, and RPA precipitated approximately 2.5-fold more ars2004 DNA compared with telomeric
DNA (Figure 1D; Supplementary Figure S5D). Such results
are consistent with the notion that the HU-induced stalling/
collapse of replication forks results in accumulation of ssDNA
at actively replicating regions of the genome.
Telomeres are recognised by checkpoint protein Rad26
and DNA repair protein Nbs1 as damaged DNA
Given that much more RPA (approximately four-fold) was
recruited to telomeres than to ars2004 during replication
(Figure 1D), and the peak of RPA recruitment to telomeres
in the absence of HU (2.47±0.70% precipitation) was comparable to the peak of recruitment of RPA to ars2004 in the
presence of HU (3.05±0.50% precipitation), we suspected
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Figure 2 Rad26 and Nbs1 recognise telomeres as damaged DNA during S-phase. For explanation of error bars, see Figure 1. (A, B) Recruitment
of Rad26 to telomeres (A) or ars2004 (B) in the absence or presence of 15 mM HU for cdc25-22 synchronised cells (n ¼ 3). (C, D) Recruitment of
Nbs1 to telomeres (C) or ars2004 (D) in the absence or presence of 15 mM HU for cdc25-22 synchronised cells (n ¼ 3). (E) Comparison of peak
percentage precipitation values (see Materials and methods) at telomeres and ars2004 in the absence or presence of 15 mM HU for Rad26 and
Nbs1. The peak percentage precipitation value ratios (telomere/ars2004) in the absence or presence of HU were calculated for the indicated
proteins. (F) Recruitment kinetics of Rad26 and RPA to telomeres are very similar in both absence and presence of HU. Data from panel A
(Rad26) and Supplementary Figure S3C (Rad11/RPA) are combined.
that replicating telomeres might be recognised as stalled/
damaged forks by DNA damage/replication checkpoint proteins. Therefore, we next monitored the cell cycle-regulated
recruitment of the checkpoint protein Rad26 (mammalian
ATRIP ortholog) to telomeres and ars2004.
Indeed, Rad26 binding to telomeres increased as telomeres
were replicated, with the peak of recruitment at 120 min after
the release from cdc25-22 arrest, whereas DNA replicationspecific Rad26 recruitment to telomeres was largely abolished
in the presence of HU (Figure 2A). Very little Rad26 was
detected at ars2004 during unperturbed cell cycle progression
(0.073±0.038% precipitation), but Rad26 was efficiently
recruited and retained at ars2004 in the presence of HU
(0.30±0.10% precipitation), consistent with the notion that
actively replicating regions are recognised and protected by Sphase checkpoint proteins after addition of HU (Figure 2B).
We also found that timing and level of RPA and Rad26
recruitment to telomeres matched well either in the presence
or absence of HU (Figures 1D, 2E and F). This is expected, as
previous studies have established that Rad26/ATRIP is recruited to ssDNA bound by RPA (Zou and Elledge, 2003).
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Taken together, these results establish that Rad26 transiently
recognises telomeres as stalled forks and/or damaged DNA
during unperturbed replication of telomeres in fission yeast.
We next monitored recruitment of the DNA repair protein
Nbs1 to telomeres. Nbs1, a component of the Mre11–Rad50–
Nbs1 (MRN) complex, is needed to recruit Tel1 (ATM) kinase
to DNA damage sites (You et al, 2005). Previously, MRN–Tel1
and the Rad3–Rad26 complexes have been shown to represent two redundant pathways that are essential for fission
yeast cells to maintain stable telomeres and to prevent
chromosome circularisation (Naito et al, 1998; Nakamura
et al, 2002; Chahwan et al, 2003). Similarly, budding yeast
MRX–Tel1 and Mec1–Ddc2 (Rad3–Rad26 orthologs) are redundantly required to maintain stable telomeres (Craven
et al, 2002), and MRX–Tel1 are recruited to telomeres in
late S-phase to promote recruitment of telomerase preferentially to shorter telomeres (Goudsouzian et al, 2006; Hector
et al, 2007; Sabourin et al, 2007).
In contrast to budding yeast, we found that Nbs1 in fission
yeast shows constitutive binding to telomeres throughout the
cell cycle with a broad approximately two-fold increase in
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binding during S-phase (Figure 2C). Much like RPA and
Rad26, the peak percentage precipitation value was approximately five-fold higher at telomeres compared with ars2004
during unperturbed S-phase (Figure 2E). However, in contrast
to RPA and Rad26, Nbs1 binding to telomeres still increased
as cells enter S-phase in the presence of HU. Therefore, it
appears that increased association of Nbs1 to telomeres does
not require actual replication of telomeric DNA. Given the
role of MRN as a sensor of DNA breaks, the presence of DNA
ends at telomeres may be the main cause for Nbs1 recruitment to telomeres. During S-phase, even before actual replication, telomeric DNA ends might become more accessible to
Nbs1 due to changes in local chromatin structure. At ars2004,
we observed that Nbs1 recruitment is markedly increased
when cells enter S-phase in the presence of HU, consistent
with the notion that MRN is involved in detection/repair of
HU-induced collapsed replication forks (Figure 2D and E).
Arrival of lagging strand DNA polymerases a and d is
delayed compared with arrival of leading strand DNA
polymerase e at telomeres
DNA replication of genomic DNA in eukaryotic cells is
performed by DNA polymerases a, d and e. DNA polymerase
a (Pola/Pol1) is in a complex with DNA primase, and thus, it
is continually required for synthesis of Okazaki fragments on
the lagging strand. Recent studies in budding yeast have
established that DNA polymerase e (Pole/Pol2) is primarily
involved in leading strand synthesis, whereas DNA polymerase d (Pold/Pol3) is mostly involved in lagging strand synthesis (Pursell et al, 2007; Nick McElhinny et al, 2008).
Interestingly, studies in both budding and fission yeasts
have found that mutations in Pole lead to shorter telomeres,
whereas mutations in DNA Pola and Pold lead to longer
telomeres (Adams-Martin et al, 2000; Ohya et al, 2002;
Dahlén et al, 2003). Thus, defects in leading strand synthesis
and lagging strand synthesis show opposite effects on telomere maintenance in both yeasts.
When cell cycle-regulated recruitment kinetics of the DNA
polymerases were compared (Figure 3A–C; Supplementary
Figures S6–S8), we were surprised to find that Pole arrived at
telomeres significantly earlier (peak at 120 min) than Pola
and Pold (peaks at 140 min) (Figure 3D). As expected,
recruitment of all three polymerases to telomeres was essentially abolished when HU was added (Supplementary Figures
S6C, S7C and S8C). In contrast, recruitment kinetics for all
three polymerases were similar at ars2004 or non-ARS
Figure 3 Differential arrival of leading strand polymerase (Pole) and lagging strand polymerases (Pola and Pold) at telomeres. For explanation
of error bars, see Figure 1. (A–C) Recruitment of Pola (A), Pole (B) or Pold (C) to telomeres and ars2004 for cdc25-22 synchronised cells (n ¼ 2).
(D) The peak of Pole recruitment to telomeres is earlier than those of Pola and Pold. Data from panels A–C are combined. (E) All three
polymerase (Pola, Pold and Pole) are recruited to ars2004 with similar kinetics. Data from panels A–C are combined. (F) Recruitment kinetics of
Pole, RPA and Rad26 to telomeres are very similar. To better compare S-phase specific recruitment of these factors, Rad26 (right y-axis) was
offset from Pole and RPA (left y-axis) to match baselines. Data from Figures 1C, 2A and 3B are combined.
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(a region approximately 30 kb from ars2004) (Figure 3E;
Supplementary Figure S9A). All polymerases showed increased and extended association with ars2004 in the presence of HU (Supplementary Figures S6D, S7D and S8D).
We also compared recruitment kinetics of Pola and Pole at
centromeres to test whether delayed arrival of the lagging
strand polymerases at telomeres might be related to the
presence of heterochromatin. However, Pola and Pole arrived
at centromeres with similar timing (Supplementary Figure
S9B). Thus, delayed arrival of lagging strand polymerases
(Pola and Pold) compared with leading strand polymerase
(Pole) is not a general feature of heterochromatin regions, but
unique to telomeres.
When recruitment patterns for Pole, RPA and Rad26 were
aligned, telomere recruitment kinetics matched quite well for
these three proteins (Figure 3F). Thus, our observations are
consistent with the notion that delayed arrival of Pola/Pold
compared with Pole could lead to accumulation of RPAcoated ssDNA on the lagging strand telomeres, which are in
turn recognised by the checkpoint protein Rad26. Once Pola/
Pold is recruited and lagging strand synthesis is completed,
RPA and Rad26 association would then be reduced, as
telomeres no longer carry long ssDNA G-tails.
S-phase specific recruitment of telomerase and Pot1,
but not Stn1, to telomeres during S-phase is dependent
on telomere replication
Telomeres are bound by various telomere-specific factors. To
understand the dynamics of those proteins that are crucial for
telomere function and found only at telomeres, we next
monitored the recruitment of telomerase, Taz1, Rap1, Pot1
and Stn1 to telomeres during cell cycle progression. The
catalytic subunit of telomerase Trt1 (TERT, ortholog of budding yeast Est2) was maximally recruited to telomeres at
140 min after release from the cdc25-22 arrest (Figure 4A).
Thus, timing of Trt1 recruitment matches more closely to the
recruitment timing of Pola (Figure 4E) than Pole. Unlike in
budding yeast (Taggart et al, 2002; Fisher et al, 2004), we did
not see recruitment of Trt1 to telomeres in G1. This is not
surprising, as previous studies in fission yeast have found
that the Ku70–Ku80 complex does not associate with Trt1,
Figure 4 Cell cycle-regulated association of telomere specific factors to telomeres. For explanation of error bars, see Figure 1. (A–D)
Recruitment of Trt1 (A; n ¼ 3), Taz1 and Rap1 (B; n ¼ 2), Pot1 (C; n ¼ 3) or Stn1 (D; n ¼ 3) to telomeres in the absence or presence of 15 mM of
HU for cdc25-22 synchronised cells. (E) S-phase specific recruitment kinetics of Pola, Trt1 and Pot1 to telomeres are very similar. To better
compare S-phase specific recruitment of these factors, Pot1 (right y-axis) was offset from Pola and Trt1 (left y-axis) to match baselines. Data
from Figures 3A, 4A and C are combined. (F) S-phase specific recruitment kinetics of Pot1 and Stn1 to telomeres are very similar. To better
compare S-phase specific recruitment of these factors, Stn1 (right y-axis) was offset from Pot1 (left y-axis) to match lowest precipitated DNA
values. Data from panels C and D are combined.
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and Ku70 is dispensable for recruitment of Trt1 to telomeres
(Subramanian et al, 2008; Webb and Zakian, 2008).
In the presence of HU, recruitment of Trt1 to telomeres was
greatly reduced (Figure 4A). Therefore, we conclude that
maximum recruitment of Trt1 to telomeres requires arrival
of the replication fork at telomeres. However, we noticed that
a significant amount of Trt1 was still recruited to telomeres in
the presence of HU (Figure 4A), even though recruitment of
replicative polymerases to telomeres was essentially eliminated by addition of HU (Supplementary Figures S6C, S7C
and S8C). Therefore, initial recruitment of telomerase to
telomeres might not be strictly dependent on arrival of the
DNA replication fork. Rather, other earlier events in G1/
S-phase, such as loading of MCM, increased binding of
Nbs1, or reduced binding of Taz1, Pot1 and Stn1 (see
below) might allow changes in the chromatin environment
at telomeres to allow some recruitment of Trt1 to telomeres
before replication of telomeric DNA. Conversely, replicationindependent extension of telomeres by telomerase might also
contribute to loading of Stn1 and Nbs1. Further studies are
needed to sort out these possibilities.
We found that the telomere dsDNA-binding protein Taz1
was constitutively bound to telomeres but showed broad
approximately two-fold reduction in telomere association
throughout S-phase. On the other hand, Rap1, a protein
thought to be recruited to telomeric dsDNA by Taz1, showed
very little changes in binding during the cell cycle
(Figure 4B). Therefore, Rap1 might be able to stay bound to
telomeres in the absence of Taz1. In fact, there are evidences
that Rap1 has a Taz1-independent role in protection of
telomeres despite the fact that Rap1 cannot be detected at
telomeres in taz1D cells by ChIP assays (Miller et al, 2005;
Subramanian et al, 2008). Alternatively, Taz1 might become
less accessible to the antibody used for ChIP, due to recruitment of other factors during S-phase. Nbs1, which shows a
broad approximately two-fold increase in S-phase association
to telomeres, could be one of those factors. In this regard, it is
interesting to note that mammalian TRF2 (Taz1 ortholog) has
been reported to associate with the MRN complex (Zhu et al,
2000). Previous studies have also reported that budding yeast
Rap1 and Rif1 (Smith et al, 2003) and mammalian TRF1
(Verdun et al, 2005) show reduced binding to telomeres
during S-phase.
Association of the ssDNA telomere-binding proteins Pot1
and Stn1 to telomeres was reduced in M/G1-phase to 20–30%
of maximum binding, followed by increases in association as
telomeres are replicated (Figure 4C and D). In fact, the Sphase specific increase in Pot1 and Stn1 association occurred
with very similar timing as increase in binding for Pola and
Trt1 (Figure 4E and F). Previous studies have found that Pola
and Trt1 can be co-immunoprecipitated during S-phase in a
DNA-independent manner, and that the Pot1-associated protein Tpz1 can pull down catalytically active telomerase from
cell extract (Dahlén et al, 2003; Miyoshi et al, 2008).
Furthermore, we found that addition of HU resulted in
reduction of telomere binding for Trt1, Pot1 and Pola
(Figure 4A and C; Supplementary Figure S6C). Thus, it is
possible that recruitment of Trt1, Pola and Pot1 might be
influenced by protein–protein interactions among these complexes.
Although Pot1 and Stn1 showed similar cell cycle-regulated recruitment patterns to telomeres in the absence of HU,
816 The EMBO Journal VOL 28 | NO 7 | 2009
Pot1 binding was much more severely reduced in the presence of HU than Stn1 binding (Figure 4C and D). As most
Stn1 (approximately 80% compared with non-HU culture)
could still be recruited to telomeres and stayed bound at an
elevated level in the presence of HU, we conclude that
recruitment of Stn1 to telomeres is not strictly dependent
on actual replication of telomeres. In addition, it appears that
there is no strong interaction between the Pot1 complex and
the Stn1–Ten1 complex (Martin et al, 2007; Supplementary
Figure S10). Thus, although late S-phase recruitment of Stn1
and Pot1 occurs with very similar timing, it is unlikely that
Pot1, Stn1 and Ten1 are recruited to telomeres as a stable
RPA-like complex, as it has been proposed for the Cdc13–
Stn1–Ten1 complex in budding yeast (Gao et al, 2007).
Conversely, as both Pot1 and Stn1 have been shown to be
essential for protection of telomeres from fusions and checkpoint activation (Baumann and Cech, 2001; Martin et al,
2007), late S-phase recruitment of Pot1 and Stn1 might
prevent checkpoint activation by inhibiting the association
of Rad3–Rad26 at telomeres either by contributing to the
timely arrival of Pola/Pold to reduce ssDNA or by competing
for G-tail binding with RPA.
Discussion
In this study, we analysed the cell cycle-regulated recruitment
of various DNA replication proteins (MCM, RPA, Pola, Pold
and Pole), the DNA damage checkpoint protein Rad26, the
DNA repair protein Nbs1 and various telomere proteins
(Trt1/TERT, Taz1, Rap1, Pot1 and Stn1) to telomeres to better
understand how semi-conservative DNA replication and telomerase-dependent telomere extension are coordinated in
fission yeast. Prior to our study, no information regarding
cell cycle-regulated behaviour of these proteins at telomeres
was available in fission yeast.
Not all changes in protein association at telomeres
during S-phase are coupled strictly to actual replication
of telomeric DNA
As summarised in Figure 5A, recruitment of MCM, Nbs1 and
Stn1 to telomeres was not inhibited by the addition of HU, a
chemical that inhibits DNA replication at telomeres. Thus, it
suggests that these factors undergo changes in telomere
association during S-phase in a manner independent of the
actual arrival of the DNA replication fork. A previous study
has shown that Stn1 recruitment to telomeres is increased in
cells carrying longer G-tails (Martin et al, 2007). In addition,
the MRN complex is known to be involved in the G-tail
generation at telomeres (Tomita et al, 2003). Therefore, one
interesting possibility is that a gradual increase in telomere
association of MRN during S-phase could lead to increased Gtail without DNA replication at telomeres, and allow recruitment of Stn1. Alternatively, Stn1 might be recruited to
telomeres by interacting with other factors (such as Nbs1
and MCM) that show increased and sustained recruitment to
telomeres without an actual increase in G-tail length at
telomeres, as increased binding in RPA and Rad26 was not
observed at telomeres during HU arrest. In fact, S-phase
specific telomere recruitment of RPA, Rad26 and Pot1 was
strongly inhibited by the addition of HU (Figures 2F and 4C),
indicating that recruitment of these factors is more strictly
coupled to replication of telomeric DNA.
& 2009 European Molecular Biology Organization
Telomere replication in fission yeast
BA Moser et al
Figure 5 Summary and model for cell cycle-regulated dynamics of telomere-associated factors. (A) Summary of ChIP data. The timeline for the
peak of protein binding at telomeres after release from cdc25-22 arrest are indicated as boxes with protein names. Degrees of reduction in
S-phase specific telomere binding by HU are also indicated by different shades on the boxes. (B) A model of telomere replication incorporating
the differential arrival of leading and lagging strand DNA polymerases at telomeres. Delayed synthesis of Okazaki fragments on the lagging
strand would lead to exposure of extended ssDNA at lagging strand telomeres, which is then bound by RPA and Rad3–Rad26. Thus, RPA and
Rad3–Rad26 are proposed to be especially important for controlling the accessibility of lagging strand telomeres to telomerase. On the other
hand, MRN–Tel1 and Dna2, likely to be involved in re-generation of the G-tail on leading strand telomeres, are expected to be more important
in controlling accessibility of leading strand telomeres to telomerase.
Telomerase catalytic subunit Trt1 was maximally recruited
to telomeres significantly after DNA Pole arrived at telomeres
(Figures 3B and 4A). Therefore, it appears that arrival of the
DNA replication fork precedes the recruitment of Trt1 to
telomeres. However, we note that residual Trt1 recruitment
in S-phase was observed even in the presence of HU
(Figure 4A). Thus, there may be a mechanism of telomerase
recruitment that is coupled to S-phase but independent of
telomere replication in fission yeast.
Delayed arrival of lagging strand DNA polymerases can
lead to transient formation of long G-tails on the
lagging strand telomeres
We were surprised to find that arrival of lagging strand DNA
polymerases (Pola and Pold) at telomeres is significantly
delayed (B20 min) compared with the arrival of the leading
strand DNA polymerase (Pole) (Figure 3D). To our knowledge, such a large delay has not been observed before, and
our data suggest that replication forks may carry an unusually large ssDNA loop on the lagging strand as they
approach telomeres, due to a long delay in synthesis of new
Okazaki fragments. The transient presence of extended
& 2009 European Molecular Biology Organization
ssDNA on the lagging strand telomere is also supported by
our observation that the ssDNA-binding protein complex RPA
and the DNA damage/replication checkpoint protein Rad26/
ATRIP accumulate at telomeres when Pole arrives at telomeres, and dissociate once the lagging strand polymerases
and telomerase arrive at telomeres (Figures 3D, F and 5B).
On the basis of a previous in vitro study, leading strand
synthesis can replicate linear DNA fully to the end, whereas
lagging strand synthesis gradually halts near the end to leave
an approximately 500 bp region as ssDNA (Ohki et al, 2001).
Therefore, our observation might simply reflect the fact that
DNA Pola-primase is less efficient in initiating lagging strand
synthesis near the end of linear DNA. In addition, repetitive
sequences found at telomeres and telomere-proximal regions
(Sugawara, 1988), heterochromatin structure at telomeric
and sub-telomeric regions (Kanoh et al, 2005; Mickle et al,
2007), and/or binding of telomere-specific factors and various DNA repair/checkpoint proteins (Nakamura et al, 2002)
could also contribute to delaying the arrival of the lagging
strand polymerases at telomeres.
Fission yeast cells lacking the telomerase catalytic subunit
Trt1 are estimated to lose only approximately three bases of
The EMBO Journal
VOL 28 | NO 7 | 2009 817
Telomere replication in fission yeast
BA Moser et al
telomeric DNA per cell division (Nakamura et al, 1997).
Thus, native telomeres appear to possess telomerase-independent mechanism(s) that promote initiation of lagging
strand synthesis very close to the ends of parental telomeres,
rather than leaving large gaps. Efficient and timely synthesis
of Okazaki fragments very close to the ends of lagging strand
telomeres may also be critical in reducing ssDNA-bound RPA
to attenuate checkpoint responses at telomeres. As the budding yeast telomere capping proteins Cdc13 and Stn1 interact
with the Pola–primase complex (Qi and Zakian, 2000; Grossi
et al, 2004), fission yeast Pot1 and Stn1 might also interact
with the Pola–primase complex and promote efficient recruitment of Pola near telomeric ends. Taz1, previously shown to
promote replication of telomeric repeats (Miller et al, 2006)
and to prevent long G-tail formation at telomeres (Tomita
et al, 2003) in fission yeast, may also have an important
function in regulating the arrival of leading and lagging
strand polymerases at telomeres.
ATR recruitment to telomeres may be especially
important for telomere length regulation on the lagging
strand telomeres
As mutations in RPA and Rad3–Rad26 (ATR–ATRIP) lead to
substantial telomere shortening, cell cycle-regulated accumulation of Rad3–Rad26 and RPA to telomeres is very important
for telomere maintenance in fission yeast (Nakamura et al,
2002; Ono et al, 2003). On the basis of our observations,
differential arrival of DNA polymerases is likely to be the
major cause for accumulation of RPA and Rad3–Rad26 to
lagging strand telomeres. RPA and Rad3–Rad26 could also be
significantly recruited to leading strand telomeres after a
post-replicative resection of the CA-rich strand by the MRN
complex and Dna2 nuclease (Tomita et al, 2003, 2004).
However, as mutations in MRN or Dna2 lead to little to no
telomere shortening, we believe that RPA and Rad3–Rad26
most likely contribute to telomere length regulation primarily
on the lagging strand telomeres in fission yeast (Figure 5B).
On the other hand, as previous genetic analyses have suggested that Rad3–Rad26 and Tel1–MRN pathways contribute
redundantly to telomere capping in fission yeast (Nakamura
et al, 2002), Rad3–Rad26 may contribute to leading strand
telomere maintenance in strains deficient for Tel1–MRN
functions.
Although simultaneous loss of the ATR–ATRIP pathway
(Rad3–Rad26 in fission yeast and Mec1–Ddc2 in budding
yeast) and the ATM–MRN pathway (Tel1–MRN in fission
yeast and Tel1–MRX in budding yeast) has been shown to
cause catastrophic loss of telomere stability in both budding
and fission yeasts, the relative importance of the ATR and
ATM pathways in telomere length maintenance is reversed
between these two yeast species (Naito et al, 1998; Craven
et al, 2002; Nakamura et al, 2002). However, we expect that
the ATM and ATR pathways provide evolutionarily conserved
functions to telomere maintenance in both yeasts. Thus, one
very intriguing model that could explain this difference is that
budding yeast primarily regulates telomere length by controlling Tel1–MRX activity on the leading strand, whereas fission
yeast primarily regulates telomere length by controlling
Rad3–Rad26 activity on the lagging strand. Although further
studies are clearly needed to directly test our hypothesis on
the strand specific actions of the ATR–ATRIP and ATM–MRN
pathways in budding and fission yeasts, studies in mammalian cells have previously suggested that telomerase and the
MRN complex are involved in preferentially extending leading strand telomeres (Chai et al, 2006a, b).
Materials and methods
Yeast strains
The fission yeast strains used in this study were constructed by
standard techniques (Alfa et al, 1993) and are listed in Supplementary Table S1. Sources and construction details of strains are also
available in Supplementary data online.
ChIP analysis
Cells were processed for ChIP analysis as previously described
(Nakamura et al, 2002) with minor modifications. Dynabeads
Protein G (Invitrogen) were added to 2 mg whole cell extracts preincubated with monoclonal anti-myc (9B11; Cell Signaling), anti-HA
(12CA5; Roche) or anti-FLAG (M2-F1802; Sigma) antibodies. For
BrdU incorporation analysis (see Supplementary data online), heat
denatured genomic DNA was pre-incubated with monoclonal antiBrdU antibody (B44; Becton-Dickinson). After extensive washes,
bead-bound DNA was recovered using Chelex-100 resin (Bio-Rad)
(Nelson et al, 2006). Recovered DNA was analysed by triplicate
SYBR Green-based real-time PCR (Bio-Rad) using primers listed in
Supplementary Table S3. Raw percent precipitated DNA values
(percentage raw-precipitation) were then calculated based on DCt
between Input and IP samples. ChIP analyses were also performed
using strains expressing untagged proteins to obtain percentage
background-precipitation values, and they were subtracted from
percentage raw-precipitation values to obtain percentage precipitation values. To compare ChIP data for different proteins and for
different loci, we then converted the percentage precipitation values
to relative precipitation values by setting the maximum percentage
precipitation values from non-HU experiments to be 1.
Supplementary data
Supplementary data are available at The EMBO Journal Online
(http://www.embojournal.org).
Acknowledgements
We thank JP Cooper, TR Cech, SL Forsburg, F Ishikawa, J Kanoh, P
Russell and VA Zakian for strains and plasmids. This work was
supported by UIC start-up fund, Sidney Kimmel Scholar Program
and the NIH grant GM078253 to TMN. LS is supported by a
pre-doctoral fellowship from the American Heart Association. EN
is currently supported by the NIH grant GM077604 and was
previously supported by Leukemia Research Foundation.
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Supplemental information
Differential arrival of leading and lagging strand DNA polymerases at fission yeast telomeres
Bettina A. Moser, Lakxmi Subramanian, Ya-Ting Chang, Chiaki Noguchi, Eishi Noguchi, and Toru
M. Nakamura
Supplemental materials and methods
Yeast strains and plasmids
For mcm2-HA, rad11-FLAG, trt1-myc and trt1Δ::his3+, original strains have been described
previously (Nakamura et al., 1997; Noguchi et al., 2004; Snaith and Forsburg, 1999; Webb and
Zakian, 2008). PCR-based methods (Bähler et al., 1998; Krawchuk and Wahls, 1999) were used to
create carboxy-terminally tagged pot1-myc, stn1-myc, ten1-FLAG, poz1-FLAG, pol1-FLAG, pol2FLAG and pol3-FLAG (Primer used are listed in supplemental Table S2). An amino-terminally
tagged myc-rad26 strain was created in four steps. First, the kanMX6 marker was integrated at the
3’ un-translated region to generate rad26+::kanMX6 strain. Second, hphMX6 marker (Sato et al.,
2005) was integrated to replace kanMX6 to generate rad26+::hphMX6 strain. Third,
rad26+::hphMX6 strain was transformed to integrate loxP-kanMX6-loxP-9myc fragment, PCR
amplified from pOM20 plasmid (Gauss et al., 2005), to the promoter region of rad26+. Finally,
kanMX6 was excised by transforming loxP-kanMX6-loxP-9myc-rad26+::hphMX6 strain with a
PW7 plasmid which encodes the P1 bacteriophage Cre recombinase (Werler et al., 2003) to
generate 9myc-rad26+::hphMX6 strain. Based on cell growth rate, HU sensitivity and stable
telomere length maintenance, all tagged strains were deemed largely intact in their functional roles
for DNA replication, DNA repair, DNA checkpoint, and telomere maintenance (supplemental
Figure S1 and data not shown). For telomere specific factors that showed cell-cycle regulated
changes in telomere binding by ChIP assays (Pot1, Stn1, Trt1 and Taz1), Western blotting
experiments were also performed on the samples collected from synchronized cell cultures to ensure
that these proteins are expressed throughout the cell cycle (supplemental Figure S2). Monoclonal
anti-myc (9B11; Cell Signaling), monoclonal anti-HA (12CA5, Roche) and monoclonal anti-Cdc2
(Y1004, Abcam) were used in these experiments. Previous studies have found that Pol1 (α), Pol3
(δ), Mcm2/Cdc19, and Rad11 (RPA) proteins are expressed throughout the cell cycle (Forsburg et
al., 1997; Kibe et al., 2007; Park et al., 1993). Based on microarray analysis of mRNA expression
1
Moser et al.
levels during the cell cycle (Rustici et al., 2004), Rad26, Nbs1, and Pol2 (ε) proteins are also
expected to present throughout the cell cycle.
Cell cycle synchronization
Fission yeast cells carrying the cdc25-22 mutation were grown overnight in 600 ml YES media
(Alfa et al., 1993) at 25 oC. Exponentially growing cells were shifted to 36 oC for 3 hours, cooled to
25 oC, and further cultured in the absence or presence of 15 mM HU or in the absence or presence
of 50 µM BrdU. Cells were then collected and processed every 20 min. Septation index was also
monitored to ensure comparable synchronization of cultures among all cell cycle experiments.
BrdU incorporation analysis
Genomic DNA was prepared from fission yeast cells expressing hENT1 and TK genes and
incubated with 50 µM BrdU. 200 ng of DNA was denatured at 95°C for 5min, immunoprecipitated
with 1 µg anti-BrdU antibody (B44; Becton-Dickinson) on Dynabeads Protein G (Invitrogen). After
extensive washes with TBSE (10mM Tris-HCl pH7.5, 150mM NaCl, 0.1mM EDTA), TBSE + 1%
Triton X-100, and TE (10mM Tris-HCl pH7.5, 1mM EDTA), bead-bound DNA was recovered
using Chelex-100 resin (Bio-Rad). For determination of relative precipitation values, see ChIP
analysis sub-section in the materials and methods section of the main text.
Co-immunoprecipitation assays
Cells were grown in YES, disrupted by cryogenic milling using Retsch MM301 (frequency 30 1/S,
3 min running time per cycle, total 5 cycles), and resuspended in lysis buffer (50 mM Tris pH8.0,
150 mM NaCl, 10% Glycerol, 5 mM EDTA, 0.5% NP40, 1 mM DTT, 1 mM Na3VO4, 1 mM
PMSF, and Roche complete protease inhibitor cocktail). Lysates were incubated with either
monoclonal anti-myc (9B11; Cell Signaling) or monoclonal anti-FLAG (M2-F1804; Sigma)
antibodies, and protein-antibody complexes were purified with Dynabeads Protein G (Invitrogen).
Purified samples were analyzed on SDS-PAGE and subsequent Western Blot analysis using either
monoclonal anti-myc (9B11) or monoclonal anti-FLAG (M2-F1804) antibodies.
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Yeast. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
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Bähler, J., Wu, J.Q., Longtine, M.S., Shah, N.G., McKenzie, A., 3rd, Steever, A.B., Wach, A.,
Philippsen, P. and Pringle, J.R. (1998) Heterologous modules for efficient and versatile
PCR-based gene targeting in Schizosaccharomyces pombe. Yeast, 14, 943-951.
Forsburg, S.L., Sherman, D.A., Ottilie, S., Yasuda, J.R. and Hodson, J.A. (1997) Mutational
analysis of Cdc19p, a Schizosaccharomyces pombe MCM protein. Genetics, 147, 10251041.
Gauss, R., Trautwein, M., Sommer, T. and Spang, A. (2005) New modules for the repeated internal
and N-terminal epitope tagging of genes in Saccharomyces cerevisiae. Yeast, 22, 1-12.
Hayashi, M., Katou, Y., Itoh, T., Tazumi, A., Yamada, Y., Takahashi, T., Nakagawa, T., Shirahige,
K. and Masukata, H. (2007) Genome-wide localization of pre-RC sites and identification of
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Kanoh, J., Sadaie, M., Urano, T. and Ishikawa, F. (2005) Telomere binding protein Taz1 establishes
Swi6 heterochromatin independently of RNAi at telomeres. Curr Biol, 15, 1808-1819.
Kibe, T., Ono, Y., Sato, K. and Ueno, M. (2007) Fission yeast Taz1 and RPA are synergistically
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Krawchuk, M.D. and Wahls, W.P. (1999) High-efficiency gene targeting in Schizosaccharomyces
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Nakamura, T.M., Morin, G.B., Chapman, K.B., Weinrich, S.L., Andrews, W.H., Lingner, J.,
Harley, C.B. and Cech, T.R. (1997) Telomerase catalytic subunit homologs from fission
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Noguchi, E., Noguchi, C., McDonald, W.H., Yates, J.R., 3rd and Russell, P. (2004) Swi1 and Swi3
are components of a replication fork protection complex in fission yeast. Mol Cell Biol, 24,
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Park, H., Francesconi, S. and Wang, T.S. (1993) Cell cycle expression of two replicative DNA
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Rustici, G., Mata, J., Kivinen, K., Lio, P., Penkett, C.J., Burns, G., Hayles, J., Brazma, A., Nurse, P.
and Bahler, J. (2004) Periodic gene expression program of the fission yeast cell cycle. Nat
Genet, 36, 809-817.
Sato, M., Dhut, S. and Toda, T. (2005) New drug-resistant cassettes for gene disruption and epitope
tagging in Schizosaccharomyces pombe. Yeast, 22, 583-591.
Snaith, H.A. and Forsburg, S.L. (1999) Rereplication phenomenon in fission yeast requires MCM
proteins and other S phase genes. Genetics, 152, 839-851.
Webb, C.J. and Zakian, V.A. (2008) Identification and characterization of the Schizosaccharomyces
pombe TER1 telomerase RNA. Nat Struct Mol Biol, 15, 34-42.
Werler, P.J., Hartsuiker, E. and Carr, A.M. (2003) A simple Cre-loxP method for chromosomal Nterminal tagging of essential and non-essential Schizosaccharomyces pombe genes. Gene,
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3
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Supplemental Figure S1. Telomere lengths of indicated cdc25-22 tagged strains used in this study.
Strains were restreaked multiple times on YES plates incubated at 25 oC to ensure telomere length
equilibrium prior to preparation of genomic DNA. After digestion with EcoRI, genomic DNA was
subjected to electrophesis on a 1% agarose gel, transferred to a nylon membrane, and hybridized to
a telomeric DNA probe (Nakamura et al., 1997).
4
Moser et al.
Supplemental Figure S2. Protein expression levels for indicated telomere specific proteins during
cdc25-22 synchronized cell-cycle experiments. After cells were synchronized in G2/M at 36 oC for 3
h, cell cultures were shifted to 25 oC, cell pellets were collected at times after shift as indicated, and
processed for Western blotting. Cdc2 was used as loading control.
5
Moser et al.
Supplemental Figure S3. DNA replication timing monitored by incorporation of BrdU in cdc25-22
synchronized cells. (A) BrdU incorporation at telomeres is inhibited by addition of 15 mM HU. (B)
BrdU is incorporated into ars2004 with similar kinetics in the presence or absence of 15 mM HU.
Error bars for non-HU data represent standard deviation from three independent experiments, while
HU experiment was done once.
6
Moser et al.
Supplemental Figure S4. Cell cycle regulated recruitment of MCM to telomeres and ars2004. For
Explanation of error bars, see Figure 1. (A, B) Comparison of MCM recruitment (n=2) and BrdU
incorporation (n=3) at telomeres (A) and ars2004 (B). (C, D) Recruitment of MCM to telomeres
(C) and ars2004 (D) in the absence or presence of 15 mM HU (n=2).
7
Moser et al.
Supplemental Figure S5. Cell cycle regulated recruitment of RPA to telomeres and ars2004. For
explanation of error bars, see Figure 1. (A, B) Comparison of RPA recruitment (n=2) and BrdU
incorporation (n=3) at telomeres (A) and ars2004 (B). (C, D) Recruitment of RPA to telomeres (C)
and ars2004 (D) in the absence or presence of 15 mM HU (n=2). (E, F) Comparison of recruitment
timing to telomeres (E) and ars2004 (F) for RPA and MCM. Data from Figure 1B (MCM) and 1C
(RPA) are combined.
8
Moser et al.
Supplemental Figure S6. Cell cycle regulated recruitment of Polα to telomeres and ars2004. For
explanation of error bars, see Figure 1. (A, B) Comparison of Polα recruitment (n=2) and BrdU
incorporation (n=3) at telomeres (A) and ars2004 (B). (C, D) Recruitment of Polα to telomeres (C)
and ars2004 (D) in the absence or presence of 15 mM HU (n=2).
9
Moser et al.
Supplemental Figure S7. Cell cycle regulated recruitment of Polε to telomeres and ars2004. For
explanation of error bars, see Figure 1. (A, B) Comparison of Polε recruitment (n=2) and BrdU
incorporation (n=3) at telomeres (A) and ars2004 (B). (C, D) Recruitment of Polε to telomeres (C)
and ars2004 (D) in the absence or presence of 15 mM HU (n=2).
10
Moser et al.
Supplemental Figure S8. Cell cycle regulated recruitment of Polδ to telomeres and ars2004. For
explanation of error bars, see Figure 1. (A, B) Comparison of Polδ recruitment (n=2) and BrdU
incorporation (n=3) at telomeres (A) and ars2004 (B). (C, D) Recruitment of Polδ to telomeres (C)
and ars2004 (D) in the absence or presence of 15 mM HU (n=2).
11
Moser et al.
Supplemental Figure S9. Polα and Polε are recruited with similar timing at Non-ARS region and
centromeres. (A) Comparison of recruitment timing to non-ARS region (~30 kb Away from
ars2004) for Polα (n=2) and Polε (n=2). (B) Comparison of recruitment timing to centromeres for
Polα (n=2) and Polε (n=2).
12
Moser et al.
Supplemental Figure S10. Pot1 and Stn1 do not form a stable complex. Pot1 can be coimmunoprecipitated with Poz1, a known Pot1 complex subunit of the Pot1 complex (left panel).
Stn1 can be co-immunoprecipitated with Ten1, a known Stn1 complex subunit (right panel).
However, we failed to detect any interaction between Pot1 and Stn1 by co-immunoprecipitation
(both panels).
13
Moser et al.
Supplemental Table S1: S. pombe strains used in this study
Tagged Protein
Strain
Genotype
no tag
TN1741
h- leu1-32 ura4-D18 ade6-M210 his3-D1 cdc25-22
no tag (hENT/TK)
TN4777
h- leu1-32::[hENT1 leu1+] ura4-D18 his3-D1 his7-366::[hsv-TK his7+] cdc25-22
Mcm2-HA
BAM7839
h+ leu1-32 ura4-D18 ade6-M210 cdc25-22 cdc19Δ::[cdc19+-HA leu1+]
Rad11-FLAG
BAM5875
h+ leu1-32 ura4-D18 his3-D1 cdc25-22 rad11+-5FLAG::kanMX6
myc-Rad26
TN7840
h- leu1-32 ura4-D18 his3-D1 cdc25-22 9myc-rad26+::hphMX6
Nbs1-myc
TN7697
h- leu1-32 ura4-D18 his3-D1 cdc25-22 nbs1+::13myc-kanMX6
Pol1-FLAG
TN4284
h- leu1-32 ura4-D18 cdc25-22 pol1+-5FLAG::kanMX6
Pol2-FLAG
TN4782
h+ leu1-32 ura4-D18 ade6-M210 his3-D1 cdc25-22 pol2+-5FLAG::kanMX6
Pol3-FLAG
TN4434
h- leu1-32 ura4-D18 his3-D1 cdc25-22 pol3+-5FLAG::kanMX6
Trt1-myc
TN7708
h- leu1-32 ura4-D18 his3-D1 cdc25-22 trt1+-G8-13myc::kanMX6
Taz1-HA
BAM4643
h- leu1-32 ura4-D18 his3-D1 cdc25-22 taz1+-3HA::ura4+
Rap1-HA
Pot1-myc
TN4688
BAM4650
h- leu1-32 ura4-D18 ade6-M210 his3-D1 cdc25-22 rap1+-3HA::LEU2
Stn1-myc
TN6886
h- leu1-32 ura4-D18 his3-D1 cdc25-22 stn1+-13myc::kanMX6
no tag
TN2411
h- leu1-32 ura4-D18 his3-D1
Pot1-FLAG
BAM4295
h- leu1-32 ura4-D18 his3-D1 pot1+-5FLAG::kanMX6
Stn1-myc
YTC6733
h- leu1-32 ura4-D18 his3-D1 stn1+-13myc::kanMX6
Pot1-FLAG Stn1-myc
TN6944
h- leu1-32 ura4-D18 his3-D1 pot1+-5FLAG::kanMX6 stn1+-13myc::kanMX6
Pot1-FLAG Poz1-myc
TN7011
h- leu1-32 ura4-D18 his3-D1 pot1+-5FLAG::kanMX6 poz1+-13myc::kanMX6
Ten1-FLAG Stn1-myc
TN7503
h- leu1-32 ura4-D18 his3-D1 stn1+-13myc::kanMX6 ten1+-5FLAG-TEV-Avi::kanMX6
h- leu1-32 ura4-D18 his3-D1 cdc25-22 pot1+-13myc::kanMX6
14
Moser et al.
Supplemental Table S2: DNA primers used in strain construction
Strain
Primer Name
Primer Sequence (5’ to 3’)
pot1-myc and
pot1-FLAG
pot1-T7
pot1-B8(x)
pot1-T9(y)
pot1-B10
CGCATTCAGCATCACATATCG
GGGGATCCGTCGACCTGCAGCGTACGAAACAATTTTCGTGCCAAATCC(1)
GTTTAAACGAGCTCGAATTCATCGATGATACAAAACTTACAATAATG(1)
GATATTTCACGTTTCCCCTC
stn1-myc
stn1-tagT
TTTGTCAATTTTTGCTTCGTACAAAAGGGAAATGGAGGCAAGCAAAGAAATATACCTGG
GTGCGAGATAATCAGTTTGTACGGATCCCCGGGTTAATTAA(1)
ATTAACCGCTTATATACCCATGTGTACTTATTGATCTGTTTCCGTAAACATATTCTTAA
ATTAATAGAGGATTGTAATATGAATTCGAGCTCGTTTAAAC(1)
stn1-tagB
ten1-FLAG
ten1-T1
ten1-B2(x)
ten1-T3(y)
ten1-B4
AGGATGCGTGCAATCATATAAGAATGGCAT
GGGGATCCGTCGACCTGCAGCGTACGAATCACATTTTTGACGTTCAGAAACCATTT(1)
GTTTAAACGAGCTCGAATTCATCGATAATCGTGTTAATGTCAGTCTTTATAAT(1)
TGTTGGAAAGACAAAACAAATTCTGGTA
poz1-myc
poz1-T3
poz1-tag(x)
poz1-tag(y)
poz1-B2
AGACTGGGGAAGTCATAACGAA
GGGGATCCGTCGACCTGCAGCGTACGAATTAATGTTTGAGGTAAGCATTTTAACA(1)
GTTTAAACGAGCTCGAATTCATCGATTTTTGGTATCTTTAAATTCCTGGAG(1)
GCTCGTGCAATCGTAACAAATA
pol1-FLAG
pol1CT-5'
pol1CT-3'module
Tpol1-5'module
Tpol1-3'
GTATGTGATGATTCTTCTTGTGG
GGGGATCCGTCGACCTGCAGCGTACGACGATGAAAATATCAGTCCCATATC(1)
GTTTAAACGAGCTCGAATTCTGAAGCTTTAATTCATTCTACCCGTTTAAG(1)
GTGAGTACTTTGCTATCCAGAGC
pol2-FLAG
pol2CT-5'
pol2CT-3'module
Tpol2-5'module
Tpol2-3'
GTGTTTGAGGAAACTTTGGTGGATAAC
GGGGATCCGTCGACCTGCAGCGTACGAGTTCAGCACAGAAAGTATGGACTG(1)
GTTTAAACGAGCTCGAATTCTAACATTTCTCGCCTGACCATGAG(1)
CGAACGTTTAAGAGCATGAGTGG
pol3-FLAG
pol3CT-5'
pol3CT-3'module
Tpol3-5'module
Tpol3-3'
TGAGACCTGTCTTGGATGCAAAGC
GGGGATCCGTCGACCTGCAGCGTACGACCAGGACATTTCATCAAATCTTTTC(1)
GTTTAAACGAGCTCGAATTCGATGGATGTTTTTAATTACTAAATGTG(1)
GCTGGCAAGTGTGCTTTGTCAGT
rad26+::hphMX
rad26-delC-T
rad26-wt-B
rad26-T5
rad26-B6
BAM96
kan-B5
CTCCTTTCGTCAATGTACCCTCAAAATGAA
CTTATTTAGAAGTGGCGCGCCTCACTAAAAATTAGTGTACAACTGCTCC(2)
GTTTAAACGAGCTCGAATTCATCGATTACTTGTTTCTTCAGTGTTGTTCT(1)
TAGTGGTACTTTAATATATTCATTTTCGTT
GCTTGCCTCGTCCCCGCCGGGTC(1)
GGCGGCGTTAGTATCGAATCGAC(1)
myc-rad26
rad26-T10
rad26-B11
rad26-CreLox-T
TAACTGTTAAGGTTTTCCAATTGC
TTTGTTTGTAGCTTTCACTCTCGT
AACAACTATTGTTACGCATAAACGAGAGTGAAAGCTACAAACAAAATGTGCAGGTCGAC
AACCCTTAAT(3)
TTCATCCGAACCCAAACTCTCCAAGTCAAAACTTTCATCAGCCATGCGGCCGCATAGGC
CACT(3)
ATGGCTGATGAAAGTTTTGACTTG
GATTCCCGGACTTTTTGAGAAGAA
rad26-CreLox-B
rad26-T12
rad26-B13
(1)
kanMX6 sequence underlined. (2)adh1 terminator sequence underlined. (3)pOM20 tagging module sequence underlined.
15
Moser et al.
Supplemental Table S3: DNA primers used in ChIP assays
Location
Primer Name
Primer Sequence (5’ to 3’)
References
telomere
(TAS1)
jk380
jk381
TATTTCTTTATTCAACTTACCGCACTTC
CAGTAGTGCAGTGTATTATGATAATTAAAATGG
(Kanoh et al., 2005)
(Kanoh et al., 2005)
ars2004
ars2004-66-F
ars2004-66-R
CGGATCCGTAATCCCAACAA
TTTGCTTACATTTTCGGGAACTTA
(Hayashi et al., 2007)
(Hayashi et al., 2007)
non-ARS
non-ARS-70-F
(Hayashi et al., 2007)
(~30 kb from ars2004)
non-ARS-70-R
TACGCGACGAACCTTGCATAT
TTATCAGACCATGGAGCCCATT
ars3002
ars3002F
ars3002R
TTGGCGCTAAACAATCTCTG
TCCTTGTCGAACTCAATTGC
This study
This study
centromere
(cenH)
cen1-dh-1
cen1-dh-2
CGAGCGATTTGAACATATGCATT
AACGTACTGACCGATTTGATCGT
(Kanoh et al., 2005)
(Kanoh et al., 2005)
16
(Hayashi et al., 2007)
The EMBO Journal (2009) 28, 795–796
www.embojournal.org
|&
2009 European Molecular Biology Organization | Some Rights Reserved 0261-4189/09
Antics at the telomere: uncoupled polymerases
solve the end replication problem
Carolyn M Price*
Department of Cancer and Cell Biology, University of Cincinnati, Cincinnati, OH, USA
*Corresponding author. Department of Cancer and Cell Biology, University of Cincinnati, 3125 Eden Avenue, Cincinnati, OH 45267-0521, USA.
E-mail: [email protected]
The EMBO Journal (2009) 28, 795–796. doi:10.1038/emboj.2009.64
Replicating the end of a linear chromosome poses a problem that can be solved by the combined action of the
general DNA replication machinery, DNA repair factors,
telomere proteins and telomerase. In this issue of The
EMBO Journal, a new study by Moser et al examines the
timing of replication, repair and telomere factor association with fission yeast telomeres. The study demonstrates
the dynamic nature of protein binding and provides
a framework for understanding how leading and lagging
strand polymerases, DNA damage signalling and telomere
factors cooperate during telomere replication.
Telomere replication is an essential process because incomplete
replication leads to telomere shortening and replicative senescence,
whereas failure to duplicate the terminal DNA structure leads to
loss of the G-strand overhang, chromosome fusions and cell death
(Smogorzewska and de Lange, 2004). The basic steps in telomere
replication involve duplication of the telomere duplex by the
standard replication machinery, generation of a G-overhang by
C-strand resection of the telomere replicated by the leading strand
polymerase, addition of G-strand repeats by telomerase and fill-in of
the complementary C-strand by Pol a/primase (Figure 1A) (Gilson
and Geli, 2007). Currently, it is unclear how these steps are
integrated, but given the complexity of a replication fork, it is
most likely that a large number of factors are needed to link the
general replication machinery to the telomere-specific replication
machinery. The ATM and ATR DNA damage signalling pathways
may be used to monitor and regulate this process (Verdun and
Karlseder, 2007; Sabourin and Zakian, 2008).
Past studies with yeast and mammalian cells have established
that a wide variety of proteins bind the telomere, but they do not all
bind simultaneously. Instead, their association and dissociation
seem to be part of a tightly choreographed series of events that
are needed to replicate and then protect the chromosome terminus
(Verdun and Karlseder, 2006; Chan et al, 2008). The current study
by Moser et al (2009) uses a series of timed chromatin immunoprecipitation (ChIP) analyses to provide our first high-resolution
view of these events. The authors performed quantitative ChIP with
synchronized cultures of Schizosaccharomyces pombe harvested at
20-min intervals during progression through S phase. This provided
a detailed picture of the association/disassociation kinetics of
replication, repair and telomere factors. To determine whether
binding of the various factors depended on DNA replication,
hydroxyurea (HU) was used to inhibit the late S-phase replication
of telomeres. By way of comparison, the authors also examined the
timing and level of association of the various factors at an early
firing replication origin (ars2004).
The ChIP analysis indicated that initial replication events are
similar at telomeres and ars2004. The general timing of MCMs and
Pol e recruitment was the same and DNA replication, as monitored
by BrdU incorporation, initiated at the time of Pol e recruitment at
both loci (Figure 1B). However, recruitment of other replication
factors, repair factors and the response to HU treatment were
startlingly different. As expected, the leading strand polymerase
Pol e, and the lagging strand polymerases Pol a and d associated
with ars2004 simultaneously and relatively little RPA or Rad 26
(ATRIP) was present during an unperturbed S phase. However, at
the telomere, binding of Pol a and d was delayed by B20 min
relative to Pol e. Moreover, the amount of telomere-bound RPA and
Rad26 increased in conjunction with Pol e association. Subsequent
binding of Pol a and d coincided with telomerase association and a
Figure 1 Telomere replication. (A) Stages in replication. (B) Dynamics of protein association.
& 2009 European Molecular Biology Organization
The EMBO Journal
VOL 28 | NO 7 | 2009 795
Uncoupled polymerases solve the end replication problem
CM Price
decrease in RPA and Rad26. The conclusion that can be drawn from
these data is that leading and lagging strand replication of the
telomeric tract are temporally separated with leading strand replication occurring first. The remaining template for lagging strand
replication is coated by RPA and thus recruits sensors linked to
the ATR-mediated DNA damage checkpoint. Subsequent lagging
strand replication is then temporally linked to telomerase recruitment.
Analysis of HU-treated cells yielded yet more interesting information. Although the replication block inhibited telomere association of Pol a, d, e, Pot1, RPA and Rad26, binding of telomerase was
only partially blocked and binding of Nbs1 (a component of the
MRN complex) and Stn1 was largely unaffected. Thus, it appears
that telomerase may be able to act independently of DNA replication. Perhaps progression into S phase is sufficient to allow binding
of the MRN complex and subsequent C-strand processing and/or
disruption of the telomeric chromatin allows telomerase access to
the DNA terminus. Once G-strand DNA becomes available it
appears that Stn1 can outcompete RPA and Rad26 for binding.
As Pot1 binding depends on replication but Stn1 binding does not,
one wonders whether the role of Stn1 is to replace RPA on telomeric
DNA in a replication-independent manner.
Taken together, these results begin to reveal how eukaryotic cells
have harnessed both the general replication and DNA damage
response machinery to take care of the end replication problem
on a linear chromosome. Inevitably, the replication fork has to
disappear at the chromosome terminus but it appears that the cell
has devised ways to exploit the resulting separation of leading and
lagging strand synthesis to ensure that the telomeric G-strand can
be extended by telomerase and the C-strand filled in by the lagging
strand replication machinery.
Acknowledgements
This study was supported by NIH grants GM041803 GM073169
to CMP.
References
Chan A, Boule JB, Zakian VA (2008) Two pathways recruit
telomerase to Saccharomyces cerevisiae telomeres. PLoS Genet 4:
e1000236
Gilson E, Geli V (2007) How telomeres are replicated. Nat Rev Mol
Cell Biol 8: 825–838
Moser B, Subramanian L, Chang YT, Noguchi C, Noguchi E,
Nakamura TM (2009) Differential arrival of leading and lagging
strand DNA polymerases at fission yeast telomeres. EMBO J 28:
810–820
Sabourin M, Zakian VA (2008) ATM-like kinases and regulation of
telomerase: lessons from yeast and mammals. Trends Cell Biol 18:
337–346
Smogorzewska A, de Lange T (2004) Regulation of telomerase by
telomeric proteins. Annu Rev Biochem 73: 177–208
Verdun RE, Karlseder J (2006) The DNA damage machinery and
homologous recombination pathway act consecutively to protect
human telomeres. Cell 127: 709–720
796 The EMBO Journal VOL 28 | NO 7 | 2009
Verdun RE, Karlseder J (2007) Replication and protection of telomeres. Nature 447: 924–931
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