In Saccharomyces cerevisiae, Cdc13 has been proposed to mediate telomerase recruitment at telomere ends. Stn1, which associates with Cdc13 by the two‐hybrid interaction, has been implicated in telomere maintenance. Ten1, a previously uncharacterized protein, was found to associate physically with both Stn1 and Cdc13. A binding defect between Stn1‐13 and Ten1 was responsible for the long telomere phenotype of stn1‐13 mutant cells. Moreover, rescue of the cdc13‐1 mutation by STN1 was much improved when TEN1 was simultaneously overexpressed. Several ten1 mutations were found to confer telomerase‐dependent telomere lengthening. Other, temperature‐sensitive, mutants of TEN1 arrested at G2/M via activation of the Rad9‐dependent DNA damage checkpoint. These ten1 mutant cells were found to accumulate single‐stranded DNA in telomeric regions of the chromosomes. We propose that Ten1 is required to regulate telomere length, as well as to prevent lethal damage to telomeric DNA.
Telomeres, the ends of eukaryotic chromosomes, which are composed of TG‐rich sequences elongated by telomerase (Greider, 1996; Lingner and Cech, 1998; Nugent and Lundblad, 1998), are critical for maintaining chromosome stability and genome integrity (Zakian, 1996) and have also been implicated in gene silencing (Lustig, 1998). In the yeast Saccharomyces cerevisiae, mutations in components or regulators of telomerase produce a gradual erosion of chromosomes, which eventually leads to death by senescence (Lundblad and Szostak, 1989; Lundblad and Blackburn, 1993; Lendvay et al., 1996).
Telomeres are capped by proteins that bind to these repeating DNA sequences (Evans and Lundblad, 2000). The S.cerevisiae EST proteins presumably form complexes that regulate telomerase activity and, hence, the length of telomeric tracts (Lendvay et al., 1996; Virta‐Pearlman et al., 1996; Evans and Lundblad, 1999; Hughes et al., 2000b), while others, such as Rap1, the SIR and yKU proteins, appear to be involved not only in telomere length control but also in telomeric silencing and DNA repair (Haber, 1999; Evans and Lundblad, 2000). Some telomeric proteins prevent recombinational events that would otherwise occur frequently between repeating telomeric sequences, and also prevent DNA repair enzymes from inappropriately intervening on the double‐strand breaks naturally present at the ends of telomeres. In yeast, several proteins have been proposed or suspected to play a role in the protection of telomeric ends, including Cdc13, Stn1 (Garvik et al., 1995; Grandin et al., 1997) and telomerase (Lundblad and Szostak, 1989), but also Rap1 (Krauskopf and Blackburn, 1998; Stavenhagen and Zakian, 1998) and the yKu70/yKu80 and Mre11/Rad50/Xrs2 complexes (Bertuch and Lundblad, 1998).
Cdc13, also known as Est4, together with Est1 and Est3, has been implicated previously as the main regulator of telomerase access to telomeric ends (Garvik et al., 1995; Lendvay et al., 1996; Lin and Zakian, 1996; Nugent et al., 1996; Virta‐Pearlman et al., 1996; Evans and Lundblad, 1999; Hughes et al., 2000a; Zhou et al., 2000). In S.cerevisiae, Est2 and Tlc1 represent the reverse transcriptase and RNA template subunits of telomerase, respectively (Singer and Gottschling, 1994; Lendvay et al., 1996; Lingner et al., 1997). Both Cdc13 and Est1 are capable of binding single‐stranded telomeric DNA (Lin and Zakian, 1996; Nugent et al., 1996; Virta‐Pearlman et al., 1996; Bourns et al., 1998; Hughes et al., 2000b). It is not known yet whether the short single‐strand extension of telomeric DNA at the distal end of telomeres acts as an anchoring structure for telomerase (Wellinger et al., 1993; Diede and Gottschling, 1999). Physical association between Est1 and Tlc1, as well as between Est3 and Tlc1, has been demonstrated recently (Hughes et al., 2000a; Zhou et al., 2000). Est1 also associates physically with Cdc13 (Qi and Zakian, 2000). Association of Est1 with Tlc1 does not require Est2, while, in contrast, association of Est3 with Tlc1 does require the presence of Est2 (Hughes et al., 2000a). Finally, immunoprecipitation of Est1 or Est3, but not of Cdc13, resulted in co‐precipitation of telomerase activity (Hughes et al., 2000a), thus confirming that Est1 and Est3 are close regulators of telomerase activity, while Cdc13 is more likely to represent a link between telomerase and telomere ends, possibly via specific interactions with Est1 (Evans and Lundblad, 1999; Qi and Zakian, 2000).
Stn1 has been shown to interact with Cdc13 by two‐hybrid analysis (Grandin et al., 1997). Like Cdc13, Stn1 has been implicated in the physical protection of telomeric ends because stn1‐13 mutant cells, like cdc13‐1 mutant cells, accumulated single‐stranded DNA in telomeric regions of chromosomes when incubated at restrictive growth temperatures (Garvik et al., 1995; Grandin et al., 1997). Moreover, stn1‐13 mutant cells have been shown to be deregulated in telomere length control and to exhibit abnormally long telomeres (Grandin et al., 1997). The Cdc13‐Stn1 complex may be part of a larger complex comprising additional telomeric proteins and may also be the target of as yet unidentified regulators. We report here the isolation of Ten1, a new protein, which we have found to interact physically with both Stn1 and Cdc13. Analysis of ten1 mutants demonstrates that Ten1 is required both for telomere length regulation and telomere end protection.
Isolation of TEN1
To gain further insight into the function of Stn1 and, hence, of Cdc13 (Grandin et al., 1997), we looked for suppressors of temperature‐sensitive (ts) stn1 mutations (see Materials and methods). YLR010 was thus identified as a weak suppressor of the growth defect of stn1‐13 and stn1‐154 mutant cells (Figure 1A). Due to these genetic interactions and to other properties described below, the product of the YLR010 gene was named Ten1, for protein involved in Telomeric pathways in association with Stn1, number 1. YLR010/TEN1, which potentially codes for a 160 amino acid protein, has never been described before (DDBJ/EMBL/GenBank accession No. AJ296344). The Ten1 protein sequence did not display significant homology to other known sequences in databases or any convincing conserved domains or structural motifs (data not shown). Ten1 can therefore be regarded as a novel protein.
Ten1 cooperates with Stn1 for the accomplishment of Cdc13 function
Overexpression of STN1 allowed cdc13‐1 cells to grow at 30°C but not at higher temperatures (Figure 1B), as shown previously (Grandin et al., 1997). On the other hand, overexpression of TEN1 did not complement the loss of function of cdc13‐1 cells, even at 30°C (Figure 1B). Strikingly, however, co‐overexpression of STN1 and TEN1 could partially rescue the loss of function of Cdc13 at temperatures up to 34–37°C (Figure 1B). Thus, although under these conditions the rescue of cdc13‐1 was partial at 37°C, as revealed by the persistence of morphological defects, cells could nevertheless form small colonies (not shown). These observations provide strong genetic arguments to suggest that Ten1 and Stn1 regulate Cdc13 function.
Overexpression of TEN1 reduces telomere length in stn1 mutant cells
stn1 mutant cells exhibit a very dramatic telomere lengthening (Grandin et al., 1997). Overexpression of TEN1 from a multicopy plasmid under the control of its own promoter clearly led to a reduction of telomere lengthening in stn1 mutant cells (Figure 1C). This effect was even more pronounced when TEN1 was overexpressed under the control of the strong, inducible, GAL1 promoter on galactose‐based medium (Figure 1C). We noted that overexpression of TEN1 from the YEp‐GAL1 plasmid also resulted in a decrease in stn1‐associated telomere lengthening even when incubation was on glucose‐based medium (Figure 1C; see also below). On the other hand, overexpression of TEN1 had no apparent effect on telomere length in wild‐type cells (Figure 1C).
Arrest phenotype of ten1 mutant cells
Sporulation of a ten1::kanMX4/TEN1 diploid strain (see Materials and methods) gave rise to only two out of four viable spores, which were always found to be Kan−. Sporulation of a ten1::kanMX4/TEN1 heterozygous diploid previously transformed with YEp195‐TEN1 (URA3) plasmid allowed the recovery of Kan+ spores, which were always also Ura+. Cells derived from such Kan+ Ura+ spores could not grow on 5‐fluoro‐orotic acid (5‐FOA)‐containing medium (5‐FOA is a drug that counterselects for Ura+ cells). These data indicate that the TEN1 gene is essential for vegetative growth.
ten1‐16 and ten1‐31, two ts alleles (see Materials and methods), were found to exhibit a severe growth defect at 37°C. To obtain a tighter arrest, these alleles were recloned into YCp111‐GAL1 (LEU2) and re‐introduced into a ten1Δ strain. The two resulting strains were found to exhibit wild‐type growth and morphology at 25°C when grown on galactose‐based medium. However, after transfer to glucose‐based medium at 37°C, cells of both strains arrested at G2/M, with a dumb‐bell shape, in a very homogeneous manner (Figure 2A). It should be noted that the ten1Δ YCp‐GAL1‐ten1‐16 (and ‐31) mutant cells could still grow on glucose medium at 25°C, albeit with defects (Figure 2A). This was due to the probable presence of intragenic promoter‐like sequences in TEN1, as TEN1 open reading frame (ORF) expressed alone, in the absence of any promoter sequences, could sustain growth of a ten1Δ strain after the plasmid expressing the wild‐type TEN1 gene (ORF + natural promoter sequences) had been shuffled out on 5‐FOA‐containing medium (data not shown). We also noted that the ten1Δ YCp‐GAL1‐ten1‐16 (and ‐31) strains could grow poorly on galactose medium at 37°C, as the probable result of the leakiness of the mutant alleles under these conditions of overexpression driven by the GAL1 promoter (Figure 2A). Therefore, growth of these ten1 mutants on either galactose medium at 37°C or glucose medium at 25°C represents semi‐permissive conditions, as observed on the corresponding fluorescence activated cell sorting (FACS) profiles (Figure 2A), while growth on glucose medium at 37°C represents restrictive conditions, as seen above.
Neither STN1 nor CDC13, when overexpressed from a multi‐copy plasmid under the control of their respective promoters, was able to improve growth of the ten1Δ YCp111‐GAL1‐ten1‐16 (and ‐31) strains on glucose‐based medium at temperatures between 25 and 37°C (data not shown). The nature of the mutations present in the ten1‐16 and ten1‐31 alleles is shown in Table I.
The G2/M arrest conferred by the ten1‐16 and ten1‐31 alleles depends on the DNA damage checkpoint
We next investigated the possibility that the arrest in the ten1‐16 and ten1‐31 alleles might be due to the activation of DNA damage checkpoints. DNA damage has been shown to activate a set of specific genes, including RAD9, whose main function is to halt the cell cycle in order to allow DNA repair, followed by cell cycle resumption if conditions allow (Weinert and Hartwell, 1988). In contrast to the ten1 RAD9+ mutants, ten1 rad9Δ double mutants no longer arrested after a few cell divisions as large‐budded cells, as seen by FACS analysis (Figure 2B). Rather, they continued to divide and formed microcolonies (Figure 2B). The ten1 rad9Δ cells eventually died (data not shown), presumably due to a failure to restrain mitosis in the presence of DNA damage as a consequence of the checkpoint defect (Garvik et al., 1995). Presumably, such cells cannot properly separate their chromosomes and die of mitotic catastrophe, possibly after too many chromosomal aberrations have accumulated. From these experiments, we conclude that ten1‐16 and ten1‐31 cells arrest at the restrictive temperature due to the presence of a functional DNA damage checkpoint.
cdc13‐1 and stn1‐13 mutants accumulate single‐stranded DNA in telomeric regions of the chromosomes when grown at restrictive temperature (Garvik et al., 1995; Grandin et al., 1997). Hybridization of DNA from ten1‐31 cells to a Y′ 32P‐labelled probe under native conditions according to the method described by Wellinger et al. (1993) indicated the presence of abnormally high levels of single‐stranded DNA at 37°C, but not at 25°C (Figure 2C). Application of the in‐gel hybridization method (Dionne and Wellinger, 1996) gave similar results (data not shown). From these experiments, we infer that accumulation of single‐stranded DNA at telomeres in ten1‐16 and ten1‐31 mutants may constitute the DNA damage that causes the activation of the Rad9‐dependent DNA damage checkpoint, as explained above.
Ten1 binds Stn1 in vivo
To verify the possibility that Ten1 and Stn1 might form a complex together, an HA‐His‐tagged version of TEN1 and a myc‐tagged version of STN1 were constructed, which allowed controlled expression by the inducible GAL1 promoter (Figure 3A). Both constructs were fully functional because their expression alone could complement a deletion of the corresponding gene. In cells expressing both constructs simultaneously, immunoprecipitation with anti‐myc antibody revealed the presence of Ten1‐HAHis on anti‐HA western blots. Conversely, immunoprecipitation with anti‐HA antibody allowed Stn1‐myc to be visualized on anti‐myc westerns (Figure 3B). The physical association between Ten1 and Stn1 was confirmed by two‐hybrid analysis (see below).
Since Stn1 has been shown to bind Cdc13 by two‐hybrid analysis (Grandin et al., 1997), a finding recently confirmed by large‐scale two‐hybrid screening (Uetz et al., 2000), we wanted to know whether Cdc13 function was necessary for the association between Stn1 and Ten1 observed above. To this end, the association between Stn1 and Ten1 was assessed in ts cdc13‐1 mutant cells after shifting the culture to the restrictive temperature of 37°C for 4 h prior to preparation of cell extracts. Importantly, under such conditions, Stn1‐myc could still bind Ten1‐GFP and vice versa (Figure 3C).
Ten1 associates with Cdc13 by two‐hybrid analysis
To know whether Ten1 could associate physically with Cdc13, we expressed simultaneously TEN1 and CDC13 from plasmids used to detect protein‐protein interactions in a yeast two‐hybrid system (Fields and Song, 1989). Y190 S.cerevisiae strains simultaneously expressing TEN1 and CDC13 or TEN1 and STN1 were found to be positive in the X‐gal assay, yielding an intense blue colour (Figure 4A). These interactions were confirmed by quantitative assays performed in liquid cultures, using o‐nitrophenyl‐β‐d‐galactopyranoside (ONPG) as a substrate to measure β‐galactosidase activity (Table II). These data demonstrate that Ten1 interacts with both Stn1 and Cdc13 by two‐hybrid analysis, thus supporting the existence within the cell of a putative complex made of Cdc13, Stn1 and Ten1.
The telomeric defect of stn1 mutants is associated with a defect in Ten1 binding
stn1‐13 mutant cells exhibit a ts growth defect at 37°C and abnormally long telomeres at all temperatures between 25 and 37°C (Grandin et al., 1997). Two other ts stn1 mutants, stn1‐101 and stn1‐154, exhibited a tighter arrest at 37°C than stn1‐13 (see Materials and methods), but shared with it the characteristic of possessing long telomeres even at permissive growth temperatures. All three Stn1 mutant proteins were found by co‐immunoprecipitation (Figure 3B) and by two‐hybrid analysis (Figure 4B; Table II) to be defective in their physical association with wild‐type Ten1. Interestingly, we found that Stn1‐13 was also defective in its association with wild‐type Cdc13 in the two‐hybrid system. In both cases, the defective interactions were found to be sensitive to temperature, as is the stn1‐13 mutation (Figure 4B; Table II).
Increasing association between two telomeric proteins interacting directly but weakly within the cell by expression of a fusion (hybrid) protein has been proposed to mimic and enhance the function achieved through the interaction between these two proteins (Evans and Lundblad, 1999). A Ten1‐Stn1 in‐frame fusion protein, placed under the control of the TEN1 promoter in a single‐copy plasmid, was fully functional because it rescued inviability of stn1Δ cells (Figure 3D) as well as that of ten1Δ cells (data not shown). Expression of a Ten1‐Stn1‐13 or a Ten1‐Stn1‐154 fusion protein totally prevented the abnormal telomere elongation conferred by the stn1‐13 or stn1‐154 mutations, respectively (Figure 3D). Importantly, both the Ten1‐Stn1‐13 and Ten1‐Stn1‐154 fusions also restored normal growth at restrictive temperatures (Figure 3D). In these experiments, the abundance of Ten1 was only twice that in wild‐type cells, as the plasmid used for expression of the fusion protein was a single‐copy plasmid, while the abundance of Stn1 was the same as in wild‐type cells because the fusions were expressed in an stn1Δ background.
Altogether, these experiments suggest that a defect in the physical association between Stn1 and Ten1 might be responsible for the long telomere phenotype encountered in the stn1 mutant cells. The nature of the mutations present in the stn1 alleles documented here is shown in Table I.
ten1 mutants with deregulated telomere length
The isolation of the ten1‐3, ten1‐6 and ten1‐13 mutant alleles, which displayed very elongated telomeres, confirmed that Ten1 is involved in telomere length maintenance (Figure 5A). Telomere lengthening in these ten1 mutants was progressive (Figure 5A). All three mutant cells exhibited no morphological or growth defects at temperatures between 25 and 37°C (data not shown). In all three ten1 mutants, telomere length deregulation resulted from a mutation in a single residue (Table I).
We next investigated whether deregulation of telomere length in these mutants was dependent or not on telomerase. Indeed, telomere length can be regulated not only by telomerase, but also by telomerase‐independent mechanisms that rely on Rad52‐dependent homologous recombination (Lundblad and Blackburn, 1993; Teng and Zakian, 1999). To this end, a tlc1Δ ten1Δ YEp195‐TEN1 strain was constructed by crossing, and transformed with a YCp111 (LEU2) plasmid harbouring either ten1‐3, ten1‐6 or ten1‐13. Transformants were further grown on 5‐FOA medium to shuffle out the plasmid containing wild‐type TEN1. We could not recover any 5‐FOA‐resistant tlc1Δ ten1Δ YCp111‐ten1‐3 colony. In addition, the tlc1Δ ten1Δ YCp111‐ten1‐6 or ten1‐13 that were recovered grew poorly. This indicated a severe synthetic defect between TEN1 and TLC1 and, possibly, an accelerated senescence (Figure 5B, right panel). Indeed, the disappearance of non‐Y′ telomeres associated with extreme telomere shortening in these strains strongly suggested that survivors had been recovered (Figure 5B, middle panel, lanes 5 and 6). Since these survivors could not be used to interpret the nature of the telomere length regulation mechanisms taking place in the ten1 tlc1Δ double mutants, we set out to obtain cells at a pre‐survival stage. To achieve this, a TEN1+/ten1Δ TLC1+/tlc1Δ YEp195‐TEN1 zygote was transformed with YCp111‐ten1‐6 (or ‐13) and the resulting transformants further grown on 5‐FOA medium to shuffle out the plasmid containing wild‐type TEN1. Following sporulation, the desired ten1 tlc1Δ spores were selected and further grown for telomere length measurement in parallel with the ten1 TLC1+ and tlc1Δ sister cells. Under such conditions, we could obtain double mutants that, according to the criteria described above, had not yet undergone senescence. Telomere elongation no longer took place in these ten1 tlc1 double mutants (Figure 5B, left panel, lanes 2 and 3), thus indicating that it is dependent on telomerase.
Mutant proteins of Ten1 conferring deregulated telomere length still bind Stn1 and Cdc13
None of the three ten1 mutations documented above affected the binding with Stn1 (Figures 3B, upper panel, lanes 5 and 6, for Ten1‐6‐HAHis and 5C for the Ten1‐3‐HAHis and Ten1‐13‐HAHis proteins). It should be noted that Ten1‐6‐HAHis no longer associated with Stn1‐13‐myc (Figure 3B, upper panel, lanes 7 and 8), which was logical since Stn1‐13‐myc did not bind wild‐type Ten1‐HAHis (Figure 3B, upper panel, lanes 3 and 4). This provided a control for these experiments. Physical association between the Ten1‐6 mutant protein and wild‐type Stn1 was confirmed by two‐hybrid analysis (Figure 4C; Table II). Interestingly, the Ten1‐3 (not shown) and Ten1‐6 mutant proteins also interacted with wild‐type Cdc13 by two‐hybrid analysis (Figure 4C; Table II).
Importantly, wild‐type Ten1‐HAHis, as well as the Ten1‐3‐HAHis and Ten1‐6‐HAHis mutant proteins, still bound wild‐type Stn1‐myc when CDC13 was simultaneously overexpressed within the cell (data not shown). This suggested that overproduction of Cdc13 did not affect the stability of the Stn1‐Ten1 complex and that Cdc13 could not displace Stn1 from Ten1 by direct titration.
To confirm that telomere length deregulation in ten1‐3 and ten1‐6 mutant cells was not the consequence of a defect of binding between Stn1 and Ten1, we used the fusion protein approach described above. We observed that ten1Δ cells expressing YEp‐STN1‐ten1‐3 or YEp‐STN1‐ten1‐6 fusion constructs exhibited telomeres elongated to the same extent as those in the ten1 mutants alone (Figure 5D). The functionality of these fusions was attested by the fact that they rescued inviability of both ten1Δ cells (Figure 5D) and stn1Δ cells (not shown). Therefore, these experiments confirm the view that a defect of binding between the Ten1‐3 or Ten1‐6 mutant proteins and wild‐type Stn1 is not the cause of ten1‐associated telomere elongation.
The results presented here provide genetic and biochemical evidence that Ten1, a novel essential protein, interacts with the telomere maintenance machinery in S.cerevisiae. We find that Ten1 is involved both in telomere length regulation, as attested by the long telomere phenotype conferred by some ten1 mutant alleles, and in telomere end protection, suggested by the presence of lethal, Rad9‐recognized, telomeric DNA damage in other ten1 mutants. Interestingly, both Stn1 and Cdc13, also previously implicated in both telomere length regulation and telomere end protection (Garvik et al., 1995; Grandin et al., 1997), were found here to associate physically with Ten1. We therefore propose the existence of a putative Cdc13‐Stn1‐Ten1 complex functioning both in recruiting telomerase at the telomere ends and in the physical protection of telomere ends. A molecular working model for the Cdc13‐Stn1‐Ten1 complex taking into account the features of the various ten1 and stn1 alleles documented here is presented in Figure 6.
Ten1 has a function in telomere length regulation in association with Stn1 and Cdc13
Three sets of data demonstrate that Ten1 functions in regulating telomere length. First, Ten1 was found by co‐immunoprecipitation to bind Stn1 in vivo when both proteins were overproduced (Figure 3). Ten1 could also associate with Cdc13 (and Stn1) in a two‐hybrid interaction (Figure 4; Table II). Secondly, the ten1‐3, ten1‐6 and ten1‐13 mutations were found to confer abnormal telomere lengthening in a telomerase‐dependent manner (Figure 5). Thirdly, TEN1 was isolated as a partial suppressor of ts stn1 mutations when expressed at a low level. In addition, Ten1 and Stn1 could cooperate in partially substituting for Cdc13 essential function (Figure 1).
We note that overexpression of TEN1 alone could not rescue the cdc13‐1 defect at all, while overexpression of STN1 could, albeit weakly (Figure 1B), unless under the control of the strong GAL1 promoter (Grandin et al., 1997). The nature of the defect in the Cdc13‐1 mutant protein, possibly involving a defect in physically interacting with Stn1 rather than with Ten1, might be at the origin of this difference. Alternatively, particularities of the structure of the putative Cdc13‐Stn1‐Ten1 complex, unknown for the moment, might result in a more efficient recruitment of Ten1 following STN1 overexpression than the converse. We also note that overexpression of CDC13 or inactivation of Cdc13 did not affect the binding between Stn1 and Ten1, which indicates that Cdc13 is unlikely to mediate the association between Stn1 and Ten1.
An important observation was that the Stn1‐13 mutant protein was defective in physically associating with both Ten1 and Cdc13 (Figures 3B and 4B; Table II). Two other Stn1 mutant proteins studied here, Stn1‐101 and Stn1‐154, displayed similar telomere length deregulation to Stn1‐13 and also the same defect in Ten1 binding. Most importantly, expression of a Ten1‐Stn1‐13 or a Ten1‐Stn1‐154 fusion protein totally abolished the stn1‐induced telomere length defect (Figure 3D). Such hybrid proteins are thought to mimic natural interactions between two proteins that are either in close proximity or even bound together, as was found to be the case for Cdc13 and Est1 (Evans and Lundblad, 1999; Qi and Zakian, 2000). Therefore, it is very probable that the function of Stn1 directly involves physical association with Ten1 (Figure 6, model 2), although it is equally possible that Ten1 possesses an additional function in promoting telomerase recruitment independently of Stn1 (Figure 6, model 1). Stn1 possibly functions as a negative regulator of Cdc13 (Grandin et al., 1997), alone or in association with Ten1, while Ten1 might function, alternately, both in promoting telomerase recruitment via interactions with Cdc13 and Est1, and in inhibiting telomerase functioning via recruitment of Stn1 to Cdc13 (Figure 6).
TEN1 is an essential gene whose failure activates the Rad9 DNA damage checkpoint
Like cdc13‐1 and stn1‐13 mutants (Garvik et al., 1995; Grandin et al., 1997), ten1‐16 and ten1‐31 mutants activated the Rad9‐dependent DNA damage checkpoint (Figure 2B). Moreover, ten1‐31 mutant cells, like cdc13‐1 mutant cells, accumulated single‐stranded DNA in telomeric regions of chromosomes (Figure 2C). This characteristic of these ten1 mutants, shared with cdc13‐1 and stn1‐13 mutants (Garvik et al., 1995; Grandin et al., 1997), was expected, given the physical and genetic interactions between Ten1, Stn1 and Cdc13 uncovered here. It is probable that this abnormally high level of single‐stranded DNA at telomeres represents the damage recognized by the Rad9 checkpoint, as suggested previously (Garvik et al., 1995). The involvement of proteins capable of recognizing single‐stranded DNA, such as RPA, in sensing the telomeric DNA damage experienced by these ten1, stn1 and cdc13 mutants, not addressed in the present study or in any previous one, should be the focus of future experiments. Also unexplored at the moment is the possibility that the ten1 mutants described here might exhibit genetic interactions with mutations in DNA replication proteins. Indeed, it is now becoming evident that conventional replication and telomere maintenance are intimately linked (Diede and Gottschling, 1999). Since Cdc13 has been shown recently to interact physically with DNA polymerase α (Qi and Zakian, 2000), it is possible that Ten1, alone or in complex with Stn1, cooperates with Cdc13 in these mechanisms.
We note that the ten1 and tlc1Δ mutations were very synthetic lethal (Figure 5B), as were the stn1‐13 and tlc1Δ mutations at 34 or 37°C (N.Grandin and M.Charbonneau, unpublished results). This led to an accelerated senescence and appearance of survivors, as demonstrated by the disappearance of the non‐Y′ telomeres (Figure 5B). These results suggest that telomerase and the Cdc13‐Stn1‐Ten1 complex play two distinct, but perhaps complementary, roles in telomere protection. It is tempting to speculate that Ten1 and Stn1 function together with Cdc13 in a putative ‘cap’ protein complex that might serve at all times to protect telomeres against inappropriate homologous recombination, for instance. In this hypothesis, the putative Cdc13‐Stn1‐Ten1 complex might represent the functional equivalent of the Oxytricha single‐stranded telomere DNA binding proteins (Froelich‐Ammon et al., 1998; Horvath et al., 1998). It is not yet known whether Stn1 and Ten1 interrupt their binding with Cdc13 during the telomere replication process, an Est1‐mediated reaction (Evans and Lundblad, 1999; Qi and Zakian, 2000). The telomeric phenotypes of stn1 and ten1 mutants support the view that both Stn1 and Ten1 might negatively regulate telomerase recruitment by Cdc13 (Grandin et al., 1997; this study), a model that suggests temporary interruption of the association between Stn1/Ten1 and Cdc13 (Figure 6). However, since neither Stn1 nor Ten1 appears to be able to bind telomeric DNA in vivo or in vitro (our unpublished data), it is not yet known how Stn1 and Ten1 can assume their function of protection during telomerase recruitment. Perhaps the putative regulation of Cdc13 by Stn1‐Ten1 does not necessitate dissociation of the ternary complex. Alternatively, telomere ends might not need physical protection by the Cdc13‐Stn1‐Ten1 complex during telomere replication.
Materials and methods
Plasmids, strains, screening and mutagenesis
General plasmids and media used in this study were as described previously (Grandin et al., 1997). DNA manipulations were performed according to standard procedures (Ausubel et al., 1998). A diploid strain heterozygous for YLR010c [genotype FY; accession No. 10692D; strain name FWEF004(HE)], in which one copy of YLR010c had been disrupted by the kanMX4 marker gene between nucleotides 49 and 435, was purchased from Euroscarf (Frankfurt, Germany). TEN1 was isolated as a low copy suppressor of the growth defect of stn1‐13 cells at 37°C (Grandin et al., 1997) using a genomic YCp50 library (Rose and Broach, 1991). Details on TEN1 isolation, as well as the procedures for construction and selection of ten1 and stn1 alleles, can be found in the Supplementary data (available at The EMBO Journal Online).
FACS analysis and tubulin staining
DNA content was analysed by flow cytometry, as described previously (Hutter and Eipel, 1979). Briefly, cells were fixed overnight, at 4°C, in 70% ethanol, treated with RNase (1 mg/ml) and pepsin (5 mg/ml), stained with propidium iodide (50 μg/ml) and analysed in a Becton Dickinson FACScan analyser. For immunostaining of tubulin of mitotic spindles, rat anti‐tubulin (clone YOL1/34, Sera‐Lab) was used. Nuclei were simultaneously stained with 4′,6‐diamidino‐2‐phenylindole (DAPI).
The pACT2 and pAS2 vectors and the Y190 strain used in the two‐hybrid system of Fields and Song (1989) have been described in Durfee et al. (1993). Y190 cells harbouring the pAS2 and pACT2 constructs were streaked on selective minimal medium. Patches of cells were then replica‐plated from the culture plates on to nitrocellulose membrane and assayed for β‐galactosidase activity using 5‐bromo‐4‐chloro‐3‐indolyl‐β‐d‐ galactopyranoside (X‐Gal; Sigma). The β‐galactosidase activity of strains expressing the different constructs was quantitated in liquid cultures using the ONPG (Boehringer) protocol, as described in Ausubel et al. (1998).
Measurement of telomere length
Genomic DNA was prepared, digested with XhoI and separated by electrophoresis in a 0.9% agarose gel in TBE, as described in Grandin et al. (1997). Southern membranes were hybridized to a 270 bp TG1–3 32P‐labelled probe representing S.cerevisiae telomeric sequences or with a 2.5 kb Y′ 32P‐labelled probe representing subtelomeric sequences from chromosome V‐R (Garvik et al., 1995). Under such conditions, telomere tracts of wild‐type cells appear as a broad band of ∼1.1–1.2 kb, which represents the average length of most chromosomes, those containing Y′ subtelomeric regions (Louis and Borts, 1995). From non‐Y′ chromosomes, XhoI cutting typically generates the fragments indicated by arrowheads near lane 1 of the left panel of Figure 5B, best revealed using the TG1–3 probe (Figure 5B) but also visible using the Y′ probe (in all other figures) due to the presence of a few TG1–3 sequences in the Y′ probe. In senescing cells, the disappearance of the non‐Y′ fragments attests to the fact that survivors have arisen by homologous recombination. Consequently, the presence of these fragments in senescing cells was used as a marker of the state before the acquisition of survivors (Figure 5B). Results were analysed using a Storm PhosphorImager (Molecular Dynamics).
Detection of single‐stranded DNA
To detect single‐stranded TG1–3 DNA, genomic DNA was prepared and digested with XhoI, as described in Grandin et al. (1997), run in a 0.7% agarose gel and subjected to non‐denaturing Southern hybridization, using either a TG1–3 or a Y′ 32P‐labelled probe, as described in Wellinger et al. (1993). Alternatively, single‐stranded DNA was detected using the in‐gel hybridization method described by Dionne and Wellinger (1996).
Supplementary data for this paper are available at The EMBO Journal Online.
We thank Leland Hartwell, Dan Gottschling, Stephen Elledge and Victoria Lundblad for the gifts of strains and plasmids. We thank Suzy Markossian and Armelle Roisin for operating the semi‐automated DNA sequencer. This work was supported by grants from the Association pour la Recherche contre le Cancer, the Centre National de la Recherche Scientifique, programme Génome and the Comités Départementaux de l'Ardèche, la Loire et la Haute‐Savoie de la Ligue Nationale contre le Cancer.
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